PHOTOSYNTHESIS AND SULFUR OXIDATION IN MICROBIAL MATS
UNRAVELLING THE ROLE OF VERSATILE CYANOBACTERIA IN ANCIENT OCEAN ANALOGUES
Dissertation
Zur Erlangung des Grades eines Doktors der Naturwissenschaften Ͳ Dr. rer. nat. Ͳ
dem Fachbereich Biologie der Universität Bremen
Judith M. Klatt Bremen, August 2015 Die vorliegende Arbeit wurde am Max-Planck-Institut für Marine Mikrobiologie in Bremen angefertigt.
1. Gutachter: Prof Dr. Friedrich Widdel 2. Gutachter: Dr. Dirk de Beer 3. Gutachter: Prof. Dr. Michael Kühl
Colloquium date: December 14, 2015 Content
SUMMARY 1
ZUSAMMENFASSUNG 3
I. Introduction
1. PAST AND PRESENT PHOTOSYNTHESIS 9
1.1. Earth’s oxygenation – An overview 10 1.2. Modules involved in photosynthesis and their evolutionary origin 14 1.2.1. Photosynthesis – An overview ………………………………. 14 1.2.2. Light absorption and Light-Harvesting Antennas …..…...... 16 1.2.3. Electron transport ……………………………………………. 17 1.2.4. Electron donors ……………………………………………… 19
1.2.5. CO2 fixation …………………………………………………. 20 1.2.6. (No) Clues about the evolutionary origin ……………….…… 20
1.3. Microbial mats 22 1.3.1. Microbial mats as analogues of ancient ecosystems ……….. 22 1.3.2. Photosynthesis in microbial mats …………………………... 24
1.4. Motivation and Outline 28
1.5. References 30
II. Manuscripts
Overview of Chapters 39
2. STRUCTURE AND FUNCTION OF NATURAL SULPHIDE OXIDIZING MICROBIAL MATS UNDER DYNAMIC INPUT OF LIGHT AND CHEMICAL ENERGY 41
3. ASSESSMENT OF THE STOICHIOMETRY AND EFFICIENCY OF
CO2 FIXATION COUPLED TO REDUCED SULFUR OXIDATION 85
4. HYDROGEN SULFIDE CAN INHIBIT AND ENHANCE OXYGENIC PHOTOSYNTHESIS IN A CYANOBACTERIUM FROM SULFIDIC SPRINGS 145 5. ANOXYGENIC PHOTOSYNTHESIS AND 2-METHYL HOPANOID PRODUCTION: AMODEL CYANOBACTERIUM ISOLATED FROM A PROTEROZOIC OCEAN ANALOG 173
6. ANOXYGENIC PHOTOSYNTHESIS CONTROLS OXYGENIC PHOTOSYNTHESIS IN A CYANOBACTERIUM FROM A SULFIDIC SPRING 215
7. REGULATION OF THE PARTITIONING BETWEEN OXYGENIC AND ANOXYGENIC PHOTOSYNTHESIS IN A CYANOBACTERIUM DOMINATING SULFIDIC SPRING MICROBIAL MATS 253
III. Perspectives
8. DISCUSSION 271
8.1. The advantage of photosynthetic versatility 272 8.1.1. “Detoxification” hypothesis ………………………………. 272 8.1.2. “Energetics” hypothesis ……………………………………. 273 8.1.3. “Direct competition” hypothesis …………………………… 275 8.1.4. “Modulation” hypothesis …………………………………… 275
8.2. Specific adaptations strategies among cyanobacteria to sulfidic conditions 288
8.2.1. Effects of H2S on cyanobacterial photosynthesis – an overview ……………………………………………… 289 8.2.2. Specific adaptation strategies and their advantage in the environment ……………………………...…………. 292
8.3. Implications 297 8.3.1. Stromatolites and oxygenic photosynthesis? ……………….. 297 8.3.2. The importance of sulfur oxidation intermediates in the Proterozoic C-, S- and O-cycle ……………………… 298
8.4. Outlook 299
8.5. References 301 IV.Contributed work
HYPERSPECTRAL IMAGING OF BIOFILM GROWTH DYNAMICS 304
SPATIAL PATTERNS AND LINKS BETWEEN MICROBIAL COMMUNITY COMPOSITION AND FUNCTION IN CYANOBACTERIAL MATS 305
EFFECT OF HIGH ELECTRON DONOR SUPPLY ON DISSIMILATORY NITRATE REDUCTION PATHWAYS IN A BIOREACTOR FOR NITRATE REMOVAL 306
LOW-LIGHT ANOXYGENIC PHOTOSYNTHESIS AND Fe-S-BIOGEOCHEMISTRY IN A MICROBIAL MAT 307
Acknowledgements 308
SUMMARY The capability to perform oxygenic photosynthesis likely evolved in a cyanobacterial ancestor, probably in microbial mats. The cyanobacterial photosynthetic repertoire is not limited to oxygenic photosynthesis. In fact some specialized cyanobacteria can switch to using H2S as an electron donor instead of H2O in a process termed anoxygenic photosynthesis. Such photosynthetic versatility of cyanobacteria might have been an important adaptation strategy to sulfidic conditions in ancient microbial mats. Furthermore, cyanobacterial anoxygenic photosynthesis might have contributed to sustaining ocean euxinia in Proterozoic oceans. Therefore, studying the activity of modern day sulfide-adapted cyanobacteria and the competitiveness of oxygenic with anoxygenic photosynthesis has broad implications. The aim of this thesis was to gain insights into the activity of sulfide-adapted cyanobacteria in microbial mats that represent ancient Earth analogues. In Chapter 2 we describe the interaction between photosynthetically versatile cyanobacteria and chemolithoautotrophic sulphide oxidizing microorganisms in such an analogue system; the
Frasassi sulphidic spring microbial mats. Dependent on the availability of O2 in the water column, two distinct end members of mat structure form; one type with cyanobacteria on top of chemolithotrophs and another type with an inverted structure, which we studied in situ using microsensors. To be able to evaluate the importance of the magnitude of the available and conserved energy for mat stratification, a theoretical background knowledge describing the energy conservation efficiency of sulfur-chemolithoautotrophs was needed, the derivation of which is described in Chapter 3. This showed that mats driven by chemosynthesis conserved substantially less energy than photosynthesis-driven mats, although similar amounts of light and chemical energy were available. The ecosystem does therefore not necessarily develop towards optimal energy usage and the dynamics of energy availability are more important for mat stratification and function than the mere magnitude. During the studies described in Chapter 2, it became apparent that obligate anoxygenic phototrophs are almost entirely absent in the Frasassi sulfidic spring mats. To elucidate the selective advantage of sulfide-adapted cyanobacteria, several physiological studies with cyanobacterial isolates were performed in Chapter 4, 5 and 6. In Chapter 7 similar studies were done on natural mats. In all Chapters we studied the combinatory effect of H2S and light on oxygenic photosynthesis and, if present, on anoxygenic photosynthesis. The study of cyanobacteria isolated from mats is challenging because classical approaches in microbiology, such as growth in a bioreactor, are not always suitable. Benthic
1 cyanobacteria tend to form biofilms, which creates physico-chemical microenvironments. We therefore developed a microsensor-based incubation system, which enabled us to simulate the steep and fluctuating gradients of light, sulfide and O2 occuring in the cyanobacterias’ natural environment (Chapter 4 and 5). In Chapter 4 the activity of an obligately oxygenic phototrophic cyanobacterium isolated from the Frasassi sulfdic springs was studied in the novel setup. We found several stimulatory or inhibitory effects of H2S on its photosynthetic activity. To get insights into the possible mechanisms causing these effects, we developed a model of the photosynthetic redox reactions, which also helped understanding how an obligate oxygenic phototrophic bacterium is so well adapted to diurnally fluctuating light and H2S concentrations. In Chapter 5 similar studies were performed on a cyanobacterium isolated from the Little Salt Spring mats, another ancient Earth analogue. We showed that the cyanobacterium performs both oxygenic and anoxygenic photosynthesis. Dependent on light intensity rates of anoxygenic photosynthetic CO2 fixation were up to twice as high as rates during oxygenic photosynthesis. The two photosynthetic modes were, however, never performed simultaneously. In contrast, Chapter 6, a bioreactor-based study of another cyanobacterial strain obtained from the Frasassi sulfidic springs, showed an elegant transition between oxygenic and anoxygenic photosynthesis, which we explain based on a simple kinetic regulation mechanism controlled by the affinity to H2S. Similar observations were made in Chapter 7, a study of natural mats dominated by a single cyanobacterial morphotype sampled from the Frasassi sulphidic springs. The very high apparent affinity to H2S allows for simultaneous oxygenic and anoxygenic photosynthesis over an astonishingly wide range of
H2S concentrations and irradiances. Overall, this thesis highlights the wide spectrum of adaptations to sulfidic conditions among cyanobacteria. A crucial factor determining success in the environment is the specific effect of the local dynamics of light and H2S on activity. The most successful cyanobacteria ancient Earth analogues are photosynthetically versatile. As exposure to sulphidic conditions is like a red line through the history of cyanobacteria, it seems intuitive that cyanobacterial anoxygenic photosynthesis might be an ancient trait. However, there is currently no robust evidence supporting this hypothesis. This thesis highlights that photosynthetically versatile cyanobacteria might, however, have had an important impact on the ancient biogeochemical cycling. Deeper knowledge concerning the timeline of the emergence of anoxygenic photosynthesis among cyanobacteria has the potential to explain major shifts on the global oxygen budgets and redox state of Earth through history.
2 ZUSAMMENFASSUNG Der evolutive Ursprung oxygener Photosynthese, und damit der einzigen natürlichen Quelle Sauerstoffs unseres Planeten, liegt wahrscheinlich in einem Vorfahren der heutigen Cyanobakterien. Ausgehend von mikrobiellen Matten begann der Siegeszug dieses Prozesses in Form der Cyanobakterien, der die Entwicklung unseres Planeten maßgeblich geprägt hat.
Nebst der einzigartigen Fähigkeit H2O als Electronendonor zu verwenden, sind bestimmte spezialisierte Cyanobakterien zusätzlich zur photosynthetischen H2S-Oxidation, einer Form der anoxygenen Photosynthese, befähigt. Diese Vielseitigkeit könnte eine wichtige Rolle in der frühen Geschichte der Photosynthese in mikrobiellen Matten gespielt haben. Außerdem könnte anoxygene Photosynthese zu der Aufrechterhaltung von Redoxstratifizierung der Ozeane während des Proterozoikums beigetragen haben. Die Charakterisierung der Aktivität zeitgenössischer Sulfid-adaptierter Cyanobacterien und der Konkurrenzfähigkeit oxygener mit anoxygener Photosynthese hat deshalb weitreichende Implikationen. Das Ziel der vorliegenden Arbeit war es Einblicke in die Aktivität von Cyanobakterien in mikrobiellen Matten, im Kontext ihrer historischen Gegenstücke, zu gewinnen. Kapitel 2 beschreibt die Interaktion zwischen photosynthetisch vielseitigen Cyanobakterien und aeroben chemolithotrophen sulfidoxidierenden Mikrooganismen in solchen Matten. Entlang der sulfdischen Quellen von Frasassi differenzieren sich die Matten abhängig von der Sauerstoffkonzentration in der darüberliegenden Wassersäule in zwei grundlegend verschiedenen Strukturen. Während die Cyanobakterien in einer Mattenvariante die oberste Schicht bilden, werden sie in der zweiten Variante von einer Schicht chemolithotropher Bakterien überdeckt. Um Einblicke in die Rolle der Energieverfügbarkeit bei der Strukturbildung und in die aus der Mattestruktur resultierende Energieumwandlungseffizienz gewinnen zu können, mußte, wie beschreiben in Kaptitel 3, eine Theorie zur Berechnung dieser Effizienz für die chemosynthetischen Mikroorganismen hergeleitet werden. Basierend auf diesem theoretischen Konzept, konnten wir zeigen, dass die Matten, in denen Chemosynthese vorherrschte, erheblich weniger Energie konservierten als Photosynthese- dominierte Matten, obwohl sich die Verfügbarkeit von Energie nicht signifikant unterschied. Die mikrobiellen Prozesse stratifizierten sich nicht entsprechend der optimalen Energieausnutzung, sondern in Abhängigkeit von der Dynamik der Energieverfügbarkeit. In Kapitel 2 wird außerdem gezeigt, dass die Photosyntheseaktivität in den mikobiellen Matten von Frasassi nahezu ausschließlich von Cyanobakterien dominiert wird. Um den Selektionsvorteil sulfidadaptierter Cyanobakterien gegenüber obligat anoxygenen Phototrophen näher zu beleuchten, wurden physiologische Studien durchgeführt, die in
3 Kapitel 4, 5 und 6 beschrieben sind. In Kapitel 7 wird eine ähnliche Studie an Cyanobakterien in intakten, natürlich vorkommenden, mikrobiellen Matten vorgestellt. In allen Kapiteln wurde der kombinatorische Effekt von H2S und Lichtintensität auf die oxygene und, wenn vorhanden, anoxygene Photosynthese untersucht. Die Untersuchung von Cyanobakterien stellte sich jedoch als Herausforderung dar, weil traditionelle Methoden der Mikrobiologie, wie das Wachstum in Bioreaktoren, nicht immer übernommen werden konnten. In Biofilmen, zu deren Bildung benthische Cyanobakterien neigen, werden physikalisch-chemische Mikrozonen geschaffen. Um dem Problem der ungewollten Biolfilmbildung Rechnung zu tragen und um die fluktuierenden
Gradienten in Licht, H2S und O2 in mikrobiellen Matten simulieren zu können, wurde ein auf Mikrosensormessungen basierendes Inkubationssystem entwickelt (Kapitel 4 und 5). In Kapitel 4 wird die Regulation der Aktivität eines aus Frasassi isolierten Cyanobakteriums charakterisiert, das ausschließlich zur oxygenen Photosynthese befähigt ist.
H2S hatte sowohl stimulierende als auch inhibierende Effekte auf die Aktivität. Basierend auf einem Model der photosynthetischen Redoxreaktionen wurden mögliche Mechanismen hinter diesen Effekten identifiziert. Gleichzeitig half dieses Model die Reaktion der Cyanobakterien auf Licht- und Sulfidfluktuationen vorherzusagen und ermöglichte somit Einblicke in die mögliche Adaptionsstrategie in der Umwelt. In Kapitel 5 wird ein Cyanobakterium untersucht, das aus mikrobiellen Matten in Little Salt Spring isoliert wurde. Dieses Isolat könnte sowohl oxygene als auch anoxygene Photosynthese betreiben. Je nach Lichtintensität waren die an anoxygene Photosynthese gekoppelten potentiellen CO2-Fixierungsraten bis zu doppelt so hoch wie die Raten während oxygener Photosynthese. Jedoch wurde nie simultane anoxygene und oxygene Photosynthese gemessen. Ganz im Gegensatz dazu, zeigt die Untersuchung eines anderen Cyanobakteriums, isoliert aus Frasassi, in Kapitel 6 einen eleganten Übergang zwischen anoxygener und oxygener Photosythese, für den wir einen einfachen kinetischen Regulationsmechanismus basierend auf der Affinität zu H2S vorschlagen. Von ähnlichen Beobachtungen in einer intakten mikrobiellen Matte, die von einem einzigen cyanobakteriellen Morphotyp dominiert wurde, werden in Kapitel 7 berichtet. Die scheinbar hohe Affinität dieses Morphotypen zu
H2S führte dazu, dass anoxygene und oxygene Photosynthese über eine erstaunliche Spanne von H2S Konzentrationen und Lichtintensitäten simultan auftraten. Diese Arbeit unterstreicht das facettenreiche Spektrum möglicher Adaptionen an sulfidische Bedingungen, das unter den Cyanobakterien zu finden ist. Entscheidend für den
Erfolg einer Spezies in der Umwelt ist der spezifische Effekt der lokalen Licht- und H2S-
4 Dynamik auf die Aktivität. In den hier untersuchten Standorten scheint der kritische erfolgsbestimmende Faktor die metabolische Flexibilität zu sein. Da sich der Kontakt mit sulfidischen Bedingungen wie ein roter Faden durch die Evolutionsgeschichte der Cyanobakterien zieht, drängt sich die intuitive Schlußfolgerung auf, dass diese Flexibiliät und Fähigkeit zu anoxygener Photosynthese ein frühen evolutiven Ursprung haben könnte. Jedoch gibt es zurzeit keine robusten Daten, die dieses Szenario belegen könnten. Schließend aus ihrer Aktivität in zeitgenössischen Analoga, zeigt diese Arbeit jedoch, dass metabolisch flexible Cyanobakterien durchaus eine wichtige Rolle in geo-historischen biogeochemischen Kreisläufen gespielt haben könnten. Weitergehende Studien den evolutiven Zeitablauf der Entstehung anoxygener Photosynthese in Cyanobakterien betreffend, haben somit das Potential globale Veränderungen des Redoxstatus der Erde zu erklären.
5 6
I. Introduction
7
8 I. Introduction
1. PAST AND PRESENT PHOTOSYNTHESIS
Photosynthesis is a process by which plants, algae and phototrophic prokaryotes convert the energy contained in light quanta to chemical forms of energy. The type of photosynthesis that often first comes to mind is oxygenic photosynthesis (OxyP) because it is performed by the plants that surround us and provides us with our life-enabling terminal electron acceptor for respiration: O2. Oxygenic phototrophs are able to couple the reduction of CO2 to the oxidation of water via a series of intermediate redox reactions, which yields biomass and O2. By harvesting and converting light energy this eaction becomes feasible. Life on today’s Earth, with very few exceptions, is almost entirely dependent on OxyP. This is even true for chemolithoautrophic processes, e.g. at deep-water hydrothermal vents, where the microorganisms depend on a suitable electron acceptor for the oxidation of the geothermally derived electron donors. These electron acceptors are - mainly O2 or NO3 (Sievert and Vetriani, 2012). The production of the latter is again dependent on photosynthetically produced O2. Photosynthesis not only provides a flux of chemical energy in the form of disequilibria (e.g., between organic carbon and O2) to other organisms (Raven, 2009); without the activity of oxygenic phototrophs, Earth would have never become a habitable planet. For instance, photosynthesis-derived oxygen initiated the formation of the stratospheric ozone layer that is crucial for UV protection, and the photosynthetic assimilation of CO2 is (still) maintaining Earth’s climate in a fairly stable state (Westerhoff et al., 1997). Among the prokaryotes, cyanobacteria are the only oxygenic phototrophs and all extant OxyP most likely evolved from one cyanobacterial ancestor (Mulkidjanian et al., 2006). The evolution of OxyP was likely predated by forms of anoxygenic photosynthesis (AnoxyP) (Blankenship et al., 2007). The electron donor of OxyP is the ubiquitously present H2O. In contrast, AnoxyP relies on other electron donors, such as H2, Fe(II), - 0 2- As(III), NO2 , formate, oxalate, H2S, S and S2O3 (Widdel et al., 1993; Kulp et al., 2008; Ghosh and Dam, 2009). Intriguingly, some specialized cyanobacteria inhabiting “extreme”, reduced, environments are capable of both OxyP and AnoxyP (referred to as “versatile” in the following) (Cohen et al., 1975). The availability of the potential electron donors for AnoxyP has changed during the history of our planet. Consequently, also the importance of different forms of photosynthesis for global primary productivity must have changed. Thus, the evolution of
9 1. PAST AND PRESENT PHOTOSYNTHESIS photosynthesis and the history of our biogeosphere are closely linked. There are several important gaps in our understanding of the evolution of photosynthesis and how it interfaced with the major developmental stages of the global ecosystem. One of the most intensively studied questions is how and when OxyP succeeded in oxygenating our planet. The geological records are restricted and these questions can often only be approached by studying the physiology of extant microorganisms and their activity in the environment. Such studies allow for modelling of biogeochemical cycling in ancient systems in analogy to the processes observed in contemporary systems. The overarching aim is to provide hypotheses, the testing of which require the identification of signatures in the rock records. In some of these models also photosynthetically versatile cyanobacteria are peripherally mentioned. It seems obvious that phototrophs that can flexibly switch between photosynthetic modes might have played an important role during major transitions in the redox state of the biosphere. Nowadays, their distribution is restricted and their physiology is poorly understood. The following gives an overview on what is currently known about the history of Earth’s oxygenation also stating the gaps in knowledge that might be filled from deeper knowledge of cyanobacterial photosynthesis, their versatility in particular. Second, it is summarized why it is so difficult to gain such insights based on cellular components involved in photosynthesis and their molecular mechanisms. Third, an overview is provided into the studies of biogeochemical cycling in ancient ocean analogues, namely microbial mats. The types of photosynthesis found in microbial mats are introduced and the main environmental factors influencing their competition are described to arrive at the central question of this study: Could photosynthetically versatile cyanobacteria have played an important role in ancient ecosystems inferring from their activity in extant analogues?
1.1. Earth’s oxygenation – An overview
Before the O2 levels in the atmosphere reached today’s levels, our planet’s biogeochemistry and the bioenergetics were quite different. The Archean oceans were likely highly reducing, mainly characterized by ferruginous conditions, and the atmosphere was rich in reduced gases, such as H2 and CH4. Accumulating evidence from the geological records point towards an oxygen increase between 2.8 and 2 billion years ago (Gya), the “Great oxidation event” (GOE) (Lyons et al., 2014) (Fig. 1.1), driven by
10 I. Introduction the oxygenic photosynthetic activity of cyanobacteria. Another oxygen releasing process, the generation from nitric oxide by methane oxidizing bacterium (Ettwig et al., 2010), would have not been able to release sufficient O2 for a modulation of the global oxygen levels (Hohmann-Marriott and Blankenship, 2011). The occurrence of the GOE 2.8 – 2 Ga ago is widely accepted (recently reviewed in Lyons et al., 2014). The exact timing of the GOE and how it relates to the evolution of OxyP is, however, still under debate. The most widely accepted scenario describes the
GOE as the point in time when the rate of O2 production was not subordinate anymore to the rate of O2 reduction by volcanic gases, such as H2 (Holland, 2002). This implies that OxyP likely had already evolved before the GOE (Fig. 1.1.), and it was only upon the decrease of geothermal sources that O2 could persistently accumulate.
Life GOE 100
10-2 (PAL) O2
‘whiffs‘ p 10-4 Neoproterozoic Event Oxidation
Deep ocean Fe2+ H S / Fe2+ Fe2+ O chemistry 2 2
Evolution of Cyanobacteria photosynthesis Purple bacteria Green sulfur bacteria Photosynthetic Eukaryotes Land-based eukaryotes Vascular plants
Dominant AnoxyPversatile OxyP photosynthesis
43 2 10 Earth age (Ga)
Figure 1.1: Evolution of photosynthesis in the context of atmospheric O2 concentration and deep ocean chemistry. On the lower panel predominant photosynthetic modes as speculated by Biddanda et al. (2011) are indicated. pO2 expressed relative to the present atmospheric level (PAL). Note that as opposed to older models of the GOE that suggest a simple 1-step rise in O2, the more recent models based on carbon and sulfur isotope fractionation and uranium data suggest a dynamic increase characterized by a transient maximum (Bekker and Holland, 2012; Planavsky et al., 2012; Partin et al., 2013). Modified from Biddanda et al. (2011), Hohmann-Marriott and Blankenship (2011) and Lyons et al. (2014).
11 1. PAST AND PRESENT PHOTOSYNTHESIS
Because of the highly reduced nature of the atmosphere and oceans, OxyP was likely only able to cause temporary increases (“whiffs”) of O2 before the GOE (Fig. 1.1).
Perhaps the strongest evidence for transient O2 accumulation during the Archean comes from trace metal enrichments in organic-rich shales that point towards oxidative weathering (Anbar et al., 2007). In contrast, non-mass-dependent (NMD) sulfur isotope fractionations during the Archean seem to tell a different story (Farquhar, 2000). NMD fractionations deviate from the behaviour of elements to scale to differences in their masses. As their generation and deposition requires extremely low O2 levels, NMD fractionations have long been considered the solid proof for an O2-deficient Archean biosphere. The value of NMD fractionations was however challenged recently (Reinhard et al., 2013). Also indicators that were long considered as supportive of the hypothesis of an early evolution OxyP, such as biomarkers, banded-iron formations, fossils resembling cyanobacteria and Archean stromatolithes, are also being challenged by more recent studies (Cairns-Smith, 1978; Widdel et al., 1993; Kirschvink and Kopp, 2008; Rasmussen et al., 2008; Bosak et al., 2013; Walter et al., 2014). Based on studies of extant microbial mats it has been suggested that “oxygen oases” in ancient stromatolithes were the main source of the transient O2 rises during the Archean (Lyons et al., 2014). Intriguingly, photosynthetically versatile cyanobacteria capable of both OxyP and AnoxyP are frequently found in extant microbial mats that are considered particularly analogous to their ancient counterparts. While the “whiff”-hypotheses were only generated rather recently, it is well established that it took another 1–2 Ga after the GOE for O2 to reach contemporary levels in another rather abrupt increase during the end-Proterozoic. The long phase between the two steps of increase is often referred to as the “boring billion” (Holland, 2006), which has triggered intensive research aimed at unravelling the mechanisms behind the conspicuous apparent stasis. While the deep oceans towards the end of the Archean were probably stratified with a CO2-rich upper layer and ferruginous deep oceans, the mid- Proterozoic ocean chemistry was likely fundamentally different. Namely, Canfield (1998) stated the existence of stratified oceans with a slightly oxic upper water column above sulfidic deep waters (the “Canfield Ocean”). Accumulating evidence suggests that only the shallow coastal oceans and basins were characterized by euxinia and that ferruginous conditions continued to prevail in the deep oceans (Canfield et al., 2002; Lyons et al., 2009; Poulton et al., 2010; Poulton and Canfield, 2011) (Fig. 1.2). By considering processes in contemporary stratified water bodies, such as the Black Sea, and by arguing
12 I. Introduction that euxinia might have reached the photic mid-depths in the water column in analogy to oxygen minimum zones in the modern ocean, Johnston et al. (2009) proposed that significant contributions to primary production by anoxygenic phototrophs, including photosynthetically versatile cyanobacteria, might have sustained euxinic conditions in the
Proterozoic oceans and tempered O2 production leading to the delay in O2 rise after the GOE (Fig. 1.2).
S0 CO2 O2
Photic zone 2- SO4
H2S CH2O H2O
[H2S] CO2
[O2] S0 +CHO CH O [Fe2+] 2 FeS2 2
Figure 1.2: Current model of the mid-Proterozoic oceans. Johnston et al. (2009) suggested that the redox cline in the Proterozoic oceans might have intruded into the photic zone triggering AnoxyP, which in turn lead to an increase in export production rates of organic matter. The organic matter exported into anoxic water depths might have sustained sulfate reduction at sufficiently high rates to perpetuate euxinia. The model by Johnston et al assumes a global extent of euxinic bottom waters. Accumulating data suggests that the shallow coastal oceans were characterized by euxinic conditions while the open deep oceans were ferruginous. Modified from Lyons et al. (2014) and Johnston et al. (2009).
The timing and cause of the second great rise in O2 concentration 0.8 – 0.5 Ga ago (the “Neoproterozoic Oxidation Event”; Fig. 1.1) is even more heavily debated than the GOE. This is because of contradictory data, a part of which actually points towards continued anoxia in the oceans throughout the Neoproterozoic (Canfield et al., 2008), while other proxies clearly indicate a dramatic rise in atmospheric O2 and completely oxic oceans (Och and Shields-Zhou, 2012). Thus, similarly to the GOE, the Neoproterozoic Oxidation Event was possibly rather dynamic and not a simple 1-step increase.
13 1. PAST AND PRESENT PHOTOSYNTHESIS
Overall, the reconstruction of Earth’s oxygenation based on the rock record is continuously evolving and a more and more refined picture is forming. The geological records are limited and only allow temporally and spatially restricted glimpses. Also, the majority of the rock records would not be interpretable without considering the full spectrum of possible biological processes that might have left their signatures. Therefore, comprehensive research from diverse disciplines is needed to enlighten when and how different metabolisms, especially photosynthesis, evolved and how they interfaced with the development of our biosphere.
1.2. Modules involved in photosynthesis and their evolutionary origin
In the absence of a robust fossil record, insights into the milestones of metabolic design might be gained by studying extant phototrophs that represent the winners of billions of years of evolution. In the following we will therefore look at the different types of photosynthesis that exist today and why it is particularly hard to understand the timing of their origin.
1.2.1. Photosynthesis – An overview The only bacterial representatives of oxygenic phototrophs are cyanobacteria. In contrast, the capability to perform anoxygenic photosynthesis is scattered over the phylogenetic tree. Anoxygenic phototrophs are generally categorized into six groups: Purple bacteria, green sulfur bacteria, filamentous anoxygenic phototrophs (FAP; previously known as green non-sulfur bacteria), heliobacteria and acidobacteria. In contrast to the other phototrophs, acidobacteria seem to lack CO2 fixation pathways and are therefore not able to perform photosynthesis in the sense of synthesizing organics from CO2. The capability to perform AnoxyP is not limited to obligate anoxygenic phototrophs, i.e., the phototrophs that cannot produce O2; cyanobacteria have also been shown to use alternative electron donors for their photosynthesis (Cohen et al., 1975).
Photosynthetic CO2 fixation can be described by the net redox reactions performed by the photosynthetic organism. Sulfide-driven AnoxyP yielding zero-valent sulfur, for instance, ideally proceeds according to 0 2 H2S + CO2 Æ S + CH2O + H2O while in OxyP the oxidation of water is coupled to CO2 fixation according to
H2O + CO2 Æ O2 + CH2O. These net reactions actually occur in two completely independent steps: light-driven redox carrier reduction (“light reactions”) and the actual CO2 fixation using the carbon
14 I. Introduction fixation pathway (“dark reactions”). In Table 1, the redox midpoint potentials of some redox carrier couples involved in photosynthesis and electron donors are listed. Clearly, due to the very positive redox potential of the H2O/O2 couple the energy, and thus photon, requirements for OxyP are substantially higher than for any type of AnoxyP.
Table 1.1: Redox midpoint potentials of some redox carrier couples and external electron donors (bold) involved in photosynthesis. From Hohmann-Marriott and Blankenship (2011).
Redox couples Redox midpoint potential at pH 7 [V]
Fdred / Fdox í0.430
+ H2 /2 H í0.420
NADPH / NADP í0.320
0 H2S / S í0.240
MQred / MQox í0.070
UQred/UQox 0.1
PQred/PQox 0.1
RFeSred/RFeSox (Rieske FeS cluster) 0.100–0.270
2+ Fe /Fe(OH)3 0.15
H2O/ 1/2 O2 0.815
Something all phototrophs have in common is that their photosynthesis is initiated by the absorption of photons by pigments. Most of this light energy is absorbed by accessory (also: antennae) pigments and then transferred in the form of excitation energy to the reaction center (RC) chlorophyll (Chl) or bacteriochlorophyll (BChl) molecules. Via various intermediate redox reactions the excited state reaction center Chls/BChls of some phototrophs drive the oxidation of the external electron donor, such as H2O in OxyP, and the reduction of redox carriers, such as NAD+. During electron transport, additionally a proton motive force (pmf) is generated, which drives the formation of ATP. Thus, during these “light reactions” of phototrophs with a linearly organized electron transport chain the excitation energy is converted into chemical energy in the form of reduced electron carriers and ATP. In phototrophs that exclusively rely on cyclic electron
15 1. PAST AND PRESENT PHOTOSYNTHESIS transport the excited state BChls are only involved in pmf and ATP generation, which in term drive redox carrier reduction through “reverse electron transport”. Independent of the type of electron transport reactions ATP and reduced electron carriers can then be used for CO2 fixation. Each group of phototrophs possesses characteristic pigments, RCs, other electron transport chain components and CO2 fixation pathways. The following sections first briefly describe how these modules differ among phototrophs. Equipped with this background information, it is then summarized what can and cannot be learnt concerning the evolutionary history of AnoxyP, OxyP and versatile cyanobacteria.
1.2.2. Light absorption and Light-Harvesting Antennas The light-harvesting accessory pigments in photosynthetic organisms absorp photons and transfer their energy to RC complexes where the actual photochemistry (primary charge separation) is initiated by special RC Chls/BChls. Accessory pigments in phototrophs are often arranged in complexes. In cyanobacteria, these are phycobilisomes (PBS) that are made of the phycobiliproteins. These are generally associated to photosystem II (PSII), while photosystem I (PSI) is mainly provided with excitation energy by other Chls (Chl a, Chl b, Chl d, Chl f). RCs of obligate anoxygenic phototrophs are also mainly associated to BChl. Green sulfur bacteria and the Chloroflexaceae and Oscillochloridaceae among the FAP mainly use BChl c, which are arranged in complex specialized comportments, the so called chlorosomes. Alternatively, also BChl a and BChl e are employed by green bacteria (Hohmann-Marriott and Blankenship, 2007). The main accessory pigment of purple sulfur bacteria is BChl a, while some representatives of the purple non-sulfur bacteria alternatively employ BChl b. Overall, the antenna complexes are highly diverse. This refers to both the intrinsic structure of the employed pigments as well as to the arrangement in the complexes. The diversity of the absorption spectra of the actual chlorophyll and non-chlorophyll pigments and absorption property modification by the protein environment is reflected in the multiplicity of in-vivo absorption spectra of phototrophic cells. Also, phototrophs are not only spectroscopically diverse, their efficiency of light harvesting is also highly variable (Manske et al., 2005). As opposed to the various pigments with accessory function, the structure of the charge-separating Chls and BChls of oxygenic and anoxygenic phototrophs in the RCs is very restricted (Hohmann-Marriott and Blankenship, 2011). In PSI and PSII of oxygenic
16 I. Introduction phototrophs the primary charge separation is driven by Chl a molecules. Analogously, charge separation in anoxygenic phototrophs is exclusively driven by BChl a. The only exception are Heliobacteriaceae, which employ BChl g in their RCs (Heinnickel and Golbeck, 2007).
1.2.3. Electron transport The reaction centers (RCs) are complexes of proteins, Chls/BChls, quinones, cytochromes and FeS clusters. In these complexes the primary charge separation, and thus the conversion of physical into chemical energy, and the first electron transport reactions occur. The RC Chls/BChls are generally denoted as P followed by their absorption maximum in the red/near infra-red region, e.g. P680 for the pair of Chls in PSII of oxygenic phototrophs (Fig. 1.3). Using excitation energy harvested by accessory pigments that has migrated to the Chls and BChls, charge separation between the Chls/BChls and primary electron acceptor (generally another Chl/BChl or a (bacterio)pheophytin (Pheo/BPhe)) occurs. The term charge separation describes the formation of a radical pair, e.g. P680•+ and Pheo•- in PSII. Thus, the RC Chl/BChl becomes oxidized. The initial electron acceptor (e.g., Pheo) rapidly reduces a quinone. This is the initiation of the electron transport reactions. The quinone donates electrons to a next redox carrier, e.g. another quinone (in so called Q-type RCs) or to an FeS cluster (in so called FeS-type RCs) (Fig. 1.3). For green sulfur bacteria and heliobacteria it is not resolved if the excited BChls directly transfer an electron to the FeS cluster while skipping the quinone, or if there is a quinone acting as an electron carrier between BChl and the FeS clusters. The flow of electrons continues from the primary electron acceptors of the RCs down the redox potential to either reduce a redox carrier (linear electron transport) or re- reduce the RC Chls (cyclic electron transport). In organisms equipped with Q-type RCs the primary electron accepting quinone (QA) donates its electrons to another quinone
(QB), which then becomes part of a mobile quinone pool that leaves the RC. Cyanobacteria are the only bacterial phototrophs that use plastoquinone (PQ) for these steps (Fig. 1.3). All other phototrophs with a Q-type RC employ menaquinone (MQ) or ubiquinone (UQ). MQ is exclusively used during photosynthesis under anaerobic conditions and has been suggested to represent the most native quinone type. In FeS-type reaction centers the primary electron accepting quinone is often denoted as A1. In PSI A1 is a phylloquinone, while the other FeS-type reaction centers of anoxygenic phototrophs use MQ as the electron acceptor (Fig. 1.3).
17 1. PAST AND PRESENT PHOTOSYNTHESIS
Q-type RCs FeS-type RCs Ͳ1.5 P840* A0 P870* A1 Ͳ1.0 FeͲSx BChl FeͲSA BPh FeͲSB Fd FNR Ͳ0.5 NAD QA H2S MQ QB hQ cytbc1 [V] 0.0 QP m cytc E hQ cytbc1 P840 cytc cytc2 0.5 P870
1.0 Purple bacteria Green S bacteria
1.5
Ͳ1.5 P700* A0 Ͳ1.0 A1 FeͲSx P680* FeͲSA FeͲSB Pheo Ͳ0.5 Fd FNR H2S NADP QA SQR [V] 0.0 QB m
E PQ hQ cytb6f PC 0.5 P700 hQ 1.0 H2OOEC P680 1.5 Oxygenic phototrophs Figure 1.3: Overview on the different reaction center (RC) types and associated electron transport reactions. Em is the redox midpotential at standard physiological conditions. Shaded blocks refer to the two types of RC complexes. Any pigment employed for light harvesting ultimately transfers its excitation energy (exciton) to a “special pair” of chlorophyll molecules, denoted as P followed by their absorption maximum, in the RC, which causes the transition from the ground state to the excited state (P*). Q-type RCs are found among purple bacteria and oxygenic phototrophs in the form of PSII. In PSII the oxygen evolving complex (OEC) oxidises water oxidation and donates electrons to the P680 RC. In all Q-type RCs electrons are transferred from P* to quinones (QA and QB) via pheophytin (Pheo) or bacteriopheophytin (BPh). Quinols leaving the RC become part of the mobile plastoquinone (PQ) or Qp pool and reduce the cytochrome (cyt) b6f or the bc1 complex. FeS-type RCs are found among green sulfur bacteria and oxygenic phototrophs (PSI). Electrons from the external electron donor, e.g., H2S, enter the electron transport chain at the level of PQ in cyanobacteria or menaquinone (MQ) in green sulfur bacteria. In FeS-type RCs the electrons are first transferred to quinones (A1). Electron transport within the RC then continues over Iron-sulfur (FeS) clusters. NADP+ or NAD+ are reduced via ferredoxin (Fd) and a ferredoxin-NAD(P) oxidoreductase (FNR).
18 I. Introduction
The mobile membrane-associated quinones, such as PQ in oxygenic phototrophs, transport electrons from the RCs to large protein complexes (quinol-acceptor oxidoreductases), and couple the re-oxidation of these complexes to the reduction of the next electron acceptor. There are three major types of complexes: the cytochrome b6f complex of oxygenic phototrophs, the alternative complex III of FAP (and Acidobacteria) and the cytochrome bc1 complex, which is used by all other anoxygenic phototrophs (Bryant et al., 2012) (Fig. 1.3). The redox reactions at these complexes are coupled to proton translocation and thus to energy generation (primarily the pmf is used to drive all energy consuming membrane coupled processes, e.g. generation of ATP). The FeS-type RCs additionally participate in the generation of reducing equivalents, such as NADPH and NADH that are used to reduce CO2. In oxygenic phototrophs a Q-type RC (PSII) and an FeS-type RC (PSI) operate in series, which enables them to “extract” electrons from water in the PSII- coupled oxygen-evolving complex (OEC), to directly reduce NADP+ and to generate ATP (Fig. 1.3).
1.2.4. Electron donors Except for OxyP, electrons in photosynthesis are extracted from the external electron donor, such as H2S (Table 1.1.), by enzymes that are not directly associated with the RC. The entry sites for the electrons are generally mobile electron carriers such as cytochromes or quinones. For instance, in cyanobacteria that are capable of AnoxyP a sulfide:quinone:oxidoreductase (SQR) directly couples the reduction of the PQ pool to the oxidation of H2S to zero-valent sulfur (Fig. 1.3). Phototrophs are often able to use more than one electron donor. Most versatile cyanobacteria seem to exclusively rely on an SQR, and thus the supply of H2S, for their anoxygenic photosynthetic activity. For some cyanobacteria reduced products of sulfide oxidation other than S0 (e.g., thiosulfate (de Wit and van Gemerden, 1987; Rabenstein et al., 1995)) have been reported and an Anacystis nidulans was even found to use also other sulfur compounds and hydrogen as electron donors (thiosulfate, Peschek (1978)). Additionally, there are some indications that some cyanobacteria might be able perform Fe2+-driven photosynthesis (Cohen, 1989; Parenteau et al., 2011). However, SQR is still the only identified enzyme involved in cyanobacterial AnoxyP. The type of electron donor and its redox potential affects the level at which electrons can enter the electron transport chain. The electrode potential of the electron acceptor in the electron transport chain relates to the energy that can be conserved in the
19 1. PAST AND PRESENT PHOTOSYNTHESIS form of the pmf (Table 1.1). In versatile cyanobacteria, for instance, SQR reduces PQ, which can then transfer the electrons to the pmf-generating b6f complex. In some anoxygenic phototrophs, ATP is additionally gained by substrate level phosphorylation during the complete oxidation of reduced sulfur compounds to sulfate via sulfite (Dahl, 1996).
1.2.5. CO2 fixation To date, six metabolic pathways that perform carbon fixation have been identified. These are the Calvin–Benson–Bassham (Calvin) cycle, the reductive tricarboxylic acid (rTCA) cycle (Evans et al., 1966), the reductive acetyl-CoA (also Wood-Ljungdahl) pathway (Ragsdale and Pierce, 2008), the 3-hydroxypropionate cycle (Herter et al., 2002); the 3- hydroxypropionate/4-hydroxybutyrate cycle (Berg et al., 2007) and the dicarboxylate/4- hydroxybutyrate cycle (Huber et al., 2008). The Calvin cycle is used by the vast majority of autotrophic organisms. Among phototrophs it is found in the cyanobacteria and most purple bacteria (Thauer, 2007). FAP and green sulfur batcteria employ the 3- hydroxypropionate cycle and rTCA cycle, respectively. Heliobacteria were so far not found to be able to grow autotrophically at all (Asao and Madigan, 2010).
1.2.6. (No) Clues about the evolutionary origin Deciphering the history of the evolution of photosynthesis from its modularly arranged network of pigments and redox reaction mediating electron carriers, enzymes and complexes seems an insurmountable task. Among the quinol-acceptor oxidoreductases, for instance, complex III is entirely distinct, while the bc1 and b6f complexes are very similar. The major difference between the latter two complexes lies in the subunit that either contains a cytochrome c or f, the evolutionary origin of which “remains a mystery” (Hohmann-Marriott and Blankenship, 2011).
The reactions involved in the different pathways of CO2 fixation differ dramatically. The capability to perform CO2 fixation was therefore likely “invented” multiple times in separate lineages. As there is no correlation of pathway occurrence to other modules employed for photosynthesis, it has been hypothesized that the earliest phototrophs grew heterotrophically and only later during evolution acquired CO2 fixation genes from chemoautotrophic organisms by lateral gene transfer (Blankenship, 2010). Similarly, the nature of the different antennae systems is highly diverse and complex, which suggests that they evolved separately and were later spread by lateral gene transfer (Blankenship, 2010).
20 I. Introduction
In contrast to the pigments in the antennae complexes, BChl a and Chl a seem to be basic constituents of photosynthesis that likely evolved early during the history of photosynthesis. This has raised the question of which one of them is the actual ancestor of all charge separating chlorophylls. No consensus exists on this question. The main argument in favour of Chl a as the “original”, is based on the pathway of Chl synthesis. Namely, BChl synthesis requires the action of two homologous enzymes for double bond reduction at two separate sites, while during Chl a synthesis only one enzyme is required. However, the argument that a simpler pathway must precede a more complex one is weak. As pointed out by Hohmann-Marriott and Blankenship (2011) the distribution of
O2-dependent steps also cannot give valuable insights into the evolutionary timing of the onset of Chl a and BChl a synthesis due to the complex nature of extant Chl biogenesis that might be the result of various more recent modulations. The conspicuous co-occurrence of a type 1 and type 2 RC in cyanobacteria poses the question if both RCs evolved in an ancestral phototroph and then were selectively lost in bacteria that are nowadays called obligate anoxygenic phototrophs (selective loss hypothesis) or if the RCs evolved in two separate lineages and were later fused in a cyanobacterial ancestor during evolution (fusion hypothesis). One variation of the selective loss hypothesis suggests that the “proto-cyanobacterium” already possessed two different photosystems that were not expressed at the same time. This is reminiscent of extant versatile cyanobacteria that only employ PSI during AnoxyP and adjust the synthesis of PSII dependent on the availability of H2S (Garcia-Pichel and Castenholz, 1990). A major step forward in understanding the evolution of photosynthesis would be to find a “primitive” cyanobacterium that selectively expresses both photosystems individually. According to Allen and Martin (2007) such anoxygenic cyanobacterium “may have adapted to present-day environments, such as eutrophic, anaerobic lakes with intermittent sulfide influx.”. To date, there is no consensus on the question as to when, where and how the different RCs evolved. At least it seems clear that all RCs evolved from a single ancestral RC (the “UrRC”) and that RCs diversified by gene duplication and lateral gene transfer events, which finally also gave rise to their homo- or heterodimeric structures (Hohmann-Marriott and Blankenship, 2011). In case of the pathways employed for external electron donor reduction, the situation is even more complicated. For each electron donor, often multiple pathways exist that are scattered among photosynthetic and non-photosynthetic prokaryotes, which is especially true for reduced sulfur oxidation pathways. The enzymatic steps utilized by
21 1. PAST AND PRESENT PHOTOSYNTHESIS present bacteria are likely the result of horizontal gene transfer, wherein it is a matter of debate if the most ancient pathways initially evolved in chemo- or phototrophic ancestors (Ghosh and Dam, 2009). Also the only one enzyme involved in cyanobacterial AnoxyP, SQR, that has been identified so far does not display congruent phylogenies with 16S rRNA of sulfur chemolithotrophs, which indicates a history of extensive horizontal gene transfer, and it is therefore not suitable for inferring evolution and divergence times of the traits of specific lineages (Griesbeck et al., 2000).
1.3. Microbial mats 1.3.1. Microbial mats as analogues of ancient ecosystems
Understanding the history of photosynthesis and O2 is a multi-disciplinary task, requiring research in the fields of geology, biology, chemistry and physics. The input of biologically oriented disciplines generally starts with the identification of possible metabolic processes. A prominent example is the discovery of ferrotrophic 2+ microorganisms that are capable of Fe oxidation in the absence of O2 (Widdel et al., 1993). This process has lent itself as an alternative explanation for banded iron formations
(BIFs) that have long been considered as a solid proxy for the O2 concentrations in the ancient oceans. Once a process of interest has been identified, biological research can go into several directions. One way is to look deeply into the metabolic design of the extant processes or to use specific subunits of the metabolism, such as protein complexes, as “molecular clocks” to infer upon their origin. As mentioned in section 1.2.6, this seems, so far, an insurmountable task in case of photosynthesis. Another important way to approach questions that are of importance in the context of understanding the history of microbially mediated processes, is to study the physiology of extant microorganisms, how metabolic rates are regulated, and what kind of signatures this might leave. Similarly, studying extant ecosystems and the interaction between processes can reveal the drivers and feedback mechanisms that better constrain the current models of ancient ecosystems, which in turn can provide (testable) hypotheses on how the ancestral forms of the studied processes might have modulated the ancient biosphere. Among extant systems microbial mats are of great interest because they may be direct analogues for ancient stromatolithes, and represent condensed analogues of stratified water bodies, such as the mid-end Proterozoic oceans (Fig. 1.2). Also, despite their large metabolic diversity, a detailed investigation of processes and their interaction in microbial mats is possible within a reasonable time frame due to their rapid response and small size.
22 I. Introduction
The fact that the oldest known fossils have a similar structure as modern stromatolithes and microbial mats is a simple argument, and a debatable one, for metabolic similarities. All processes in microbial mats are an interplay of several feedback mechanisms that shape the effective diurnal O2 budget. This highly complex way of microbial life had likely already evolved by the time of Earth’s oldest sedimentary record, which is inferred from the fact that modern microbial mats and ancient lithified stromatolithes and MISS (microbially induced sedimentary structures) have a laminated structure in common (Stal, 2002; Noffke and Awramik, 2013). OxyP likely evolved in such microbial structures, creating local oxygen oases long before the first rise in atmospheric oxygen concentration (the great oxidation event, GOE) (Buick, 2008). As microbial mats are thought to have covered most of the oceans’ shallow regions at least until the end of the Proterozoic (Fig. 1.4), such localized oxygen hotspots might explain how biology drove the intermittent “whiffs of oxygen” before the GOE (e.g., Lyons et al., 2014) (section 1.1.; Fig. 1.1.). Therefore, understanding the control of the effective oxygen budget of contemporary microbial mats, might also inform the geological records.
Figure 1.4: An artist’s illustration of the Archean Oceans with stromatolites covering the shallow coastal regions. Image Credit: Peter Sawyer / Smithsonian Institution
However, there are some clear differences between modern microbial mats and ancient stromatolites. Most prominently, the vast majority of modern mats do not lithify
23 1. PAST AND PRESENT PHOTOSYNTHESIS
(Walter et al., 1992). Still, it needs to be considered that extant microbial mats are tremendously diverse and it is very likely that also only a small fraction of Archean and Proterozoic mats actually lithified and are available in the rock record. The apparent shortcoming of the missing similarity between modern and ancient mats, thus actually underpins the need for a deeper understanding of extant mats to eventually be able to track them through the geological record.
1.3.2. Photosynthesis in microbial mats Phototrophic microbial mats are mainly driven by light energy that is transferred to the chemical realm by photosynthesis. This creates new thermodynamic disequilibria, the energy of which is further exploited by other metabolisms. Both light and chemical energy are characterized by a specific average magnitude and spatio-temporal variability, which together impact the structure and function of the mat. As a result of the directionality of the external energy sources, mass transfer phenomena and thermodynamics, microbial mats are generally organized in strata perpendicular to the direction of the energy fluxes. The stratification of specialized functional groups is mainly governed by the energy yields associated with each available energy source, the efficiency at which the energy is exploited and converted into biomass, and by the kinetics of the reaction performed by the different microorganisms. Still, the mat-specific arrangement of groups of microorganisms is hard to predict because it is finally determined by characteristic physiological properties, such as regulation of metabolic rates, switching between different metabolisms, motility, or induction of defence mechanisms (e.g. toxin production) (Jørgensen and Postgate, 1982; Garcia-Pichel et al., 1994; Schwedt et al., 2011). These properties shape the microorganisms’ niche in the environment, which is therefore characterized by a seemingly endless number of parameters. Here, the interplay between the time scales of energy fluctuations and the speed at which the microorganisms can adapt will also be important. Also, feedback effects of microbial activity on other processes and specific interactions – competitive or beneficial – between microorganisms have to be considered to understand structure and function of microbial mats. Despite the complexity of process interaction, certain general patterns are shared among microbial mats. For instance, in the absence of an external supply of potential electron donor for AnoxyP, microbial mats are ultimately driven by OxyP. Organic material resulting from excretion by and death of the oxygenic phototrophs is then used in fermentation, or consumed by aerobic and anaerobic respiratory processes, such as sulfate
24 I. Introduction reduction (SR). SR liberates hydrogen sulfide into the mat, which can be used by chemo- or phototrophic microorganisms. As such microbial mats are driven by an energy source, light, that is diurnally fluctuating and exhibits steep gradients in the mat and also steep and fluctuating chemical microgradients, e.g., of O2 and sulfide concentration, develop. The phototrophic mat inhabitants thus have to cope with enormous changes of the local physicochemical parameters over a diurnal cycle.
1.3.2.1. Anoxygenic phototrophs in microbial mats In microbial mats driven by OxyP, the main sulfide source, SR, is generally located in the deeper layers of microbial mats. Obligate sulfur-phototrophs are therefore faced with the dilemma of opposing gradients of their energy source, light, and electron donor for photosynthesis, which displaces dense populations from the uppermost position in the mats. The light absorption spectra of anoxygenic phototrophs are well tuned to the local light environment underneath oxygenic phototrophs. Even though both oxygenic and obligate anoxygenic phototrophs rely on light energy, these functional groups do not directly compete for it because they utilize complementary wavelengths of the light spectrum (Fig. 1.5). According to ecological theory, this differentiation of spectral niches promotes their coexistence (Gause, 1934; MacArthur and Levins, 1967). Besides light, nutrients can also limit photosynthesis in microbial mats (Pinckney et al., 1995; Al-Thani et al., 2014). In this scenario the different photosynthetic functional groups compete for this limiting factor. Here, the direction of supply is of major importance for the outcome of this competition. Specifically, if the limiting nutrient is mainly externally supplied or if nutrient remineralisation mainly occurs in the aerobic layers of the mat, OxyP is favoured over AnoxyP. In reverse, if anaerobic remineralisation accounts for the major fraction of nutrient supply to photosynthetic microorganisms in the upper layers, anoxygenic phototrophs are potentially able to establish a nutrient “gauntlet”, thus tempering OxyP (Johnston et al., 2009). Intriguingly, this concept does not fully apply if bioavailable nitrogen limits productivity because this would favour growth of cyanobacteria capabable of N2 fixation, unless additional trace metal scarcity limits their diazotrophic activity (Anbar and Knoll, 2002; Glass et al., 2009). Anoxygenic phototrophs also have to directly compete with sulfur- chemolithotrophs for electron donor. Large sulfur bacteria seem to generally have higher affinities towards sulfide and can therefore outcompete anoxygenic phototrophs dependent on the temporal dynamics of the depth of the oxic-anoxic interface (Jørgensen and Marais, 1986). Some species among the purple sulfur bacteria can perform
25 1. PAST AND PRESENT PHOTOSYNTHESIS
chemolithotrophic growth with O2 as electron acceptor. There are no known examples of permanently chemosynthetic growth of purple sulfur bacteria, possibly because of the low affinity to sulfide (Overmann and Garcia-Pichel, 2013). Also among the different groups of anoxygenic phototrophs there can be competition for electron donor and consequently the interplay between the light utilization efficiency and affinity towards sulfide are expected to substantially shape the fitness of the phototrophs. Also, the toxicity of H2S has to be considered. For instance, purple non-sulfur bacteria can only utilize sulfide when present at low concentrations but would be toxified under high-sulfide conditions where purple and green sulfur bacteria thrive.
0.8 1 2 3
0.6 3
0.4 1 2 0.2 Absorpion (rel. units)
0.0 400 500 600 700 800 900 O (nm)
Figure 1.5: In vivo absorption spectra of a filamentous cyanobacterium (1), purple sulfur bacterium (2) and filamentous anoxygenic phototroph (3) from the Frasassi sulfidic spring microbial mats. Spectra were obtained by hyperspectral imaging on a cellular level (Polerecky et al., 2009).
Yet another factor that is expected to affect the activity of obligate anoxygenic phototrophs and that also plays an important role in the competition among the different groups of anoxygenic phototrophs, is the local O2 concentration and the resulting photooxidative stress. Triplet excited photosynthetic pigments, such as bacteriochlorophylls, act as photosensitizers, i.e., their excitation energy can be transferred to ground state O2 leading to the direct formation of singlet oxygen (Glaeser et al., 2011). If not quenched by secondary pigments, carotenoids, the singlet oxygen and other resulting reactive oxygen species (ROS; e.g., superoxide, peroxide, hydroxyl radicals) cause severe cell damage by reacting with diverse biomolecules, such as
26 I. Introduction proteins, DNA, RNA and lipids (Asada, 1996). The simultaneous presence of bacteriochlorophylls and O2 may therefore be lethal. The development of ROS is counteracted in most purple sulfur bacteria by an inhibitory effect of high light intensities and O2 on the expression of photosynthesis genes (Oh and Kaplan, 2001; Beatty, 2002). Intriguingly, this does not imply that the anoxygenic photosynthetic activity of purple sulfur bacteria is directly inhibited by O2. In the dynamic mat microenvironment the photosynthetic apparatus synthesized during anoxic and dark conditions in the night can operate at low oxygen tension during the day and activity can be maintained due to complex defence response mechanisms to the photooxidative stress (Berghoff et al., 2011). In contrast, anoxygenic photosynthetic activity in green sulfur bacteria is directly suppressed by O2. Specifically, in this group of phototrophs energy transfer from the BChl c in the chlorosomes to the reaction centers, and therefore photosynthesis, appears to be substantially decreased under aerobic conditions likely due to redox-dependent quenching of the excited BChl c in the chlorosomes (Frigaard and Matsuura, 1999). This quenching prevents both the direct formation of singlet O2 in the reaction center, as well as superoxide generation by reaction of O2 with reduced ferredoxin. Such O2 dependent quenching mechanisms were not observed in Chloroflexus aurantiacus, a filamentous anoxygenic phototroph (FAP) (Frigaard et al., 1999). Instead their reaction center has been hypothezised to form less reactive oxygen species than green sulfur bacteria and ferredoxin is not involved in their photosynthetic electron transport and CO2 fixation pathway, which makes them substantially more resistant towards O2 (Blankenship and Matsuura, 2003). Also, FAP upregulate carotenoid synthesis under aerobic conditions (Pierson and Castenholz, 1974) and they are able to switch between photoautotrophy, photoheterotrophy and aerobic respiration dependent on the local availability of light, sulfide, O2 and cyanobacterial excudates (van der Meer et al., 2005), which also applies to purple non-sulfur bacteria. In the latter phototrophs respiration is completely inhibited by light (Scherer, 1990).
1.3.2.2. Cyanobacteria in microbial mats Oxygenic phototrophs have also developed diverse strategies to cope with the dynamics of light, and O2 and sulfide availability. In research on oxygenic photosynthesis, sulfide, specifically the species H2S, was initially studied because of its toxicity. Namely, it has been shown that it inhibits PSII activity and various other enzymes indirectly involved in photosynthesis, such as antioxidant enzymes and carbonic anhydrases (Evans, 1967; Coleman, 2000; Badger and Price, 2003; Miller and Bebout, 2004), and that it promotes
27 1. PAST AND PRESENT PHOTOSYNTHESIS the generation of reactive oxygen species under aerobic intracellular conditions (Chen and Morris, 1972; Misra, 1974; Hoffman et al., 2012). Indeed, the vast majority of cyanobacteria are highly sensitive to sulfide as it irreversibly blocks the OEC in PSII. Only certain specialized cyanobacteria are able to inhabit environments where they are regularly exposed to sulfidic conditions, e.g., microbial mats. The adaptation strategies among such cyanobacteria to sulfide are highly diverse, as already recognized by (Cohen et al., 1986). The authors defined a simple classification scheme for the adaptations based on four types of responses to sulfide: (1) Irreversible inhibition of photosynthesis by traces of sulfide (2) Continued operation of oxygenic PS in the presence of sulfide
(3) Simultaneous oxygenic and anoxygenic PS – PSII is not sensitive to H2S. CO2
fixation rates are maximal at low H2S when anoxygenic and oxygenic PS are performed simultaneously
(4) Capable of both oxygenic and anoxygenic PS - PSII is partially inhibited by H2S.
CO2 fixation rates are maximal when exclusively oxygenic P or exclusively anoxygenic P are performed. Accordingly, during simultaneous operation of both
photosynthetic modes at intermediate H2S concentrations CO2 fixation rates are decreased. Thus, cyanobacteria seem to be equipped with a multifaceted spectrum of adaptations to sulfide. Their anoxygenic photosynthetic activity has often been classified as a detoxification mechanism (e.g., in Stal (2002)). In the past years however cyanobacterial AnoxyP has been increasingly recognized as a process that might have played an important role in the carbon and sulfur cycle in the redox stratified mid-end Proterozoic oceans (Johnston et al., 2009).
1.4. Motivation and Outline Research into the evolution of photosynthesis and its role in Earth’s history has recently focussed on refining our understanding of when crucial events occurred (Anbar et al., 2007; Buick, 2008; Sessions et al., 2009). So far however, only a few studies have aimed to characterize the microbial interactions that may have influenced the evolution of photosynthesis. Therefore, extant ecosystems might still reveal insights into the factors that directly select for organisms with enhanced performance or fitness under given sets of conditions that are analogous to ancient ecosystems. Extant phototrophic microbial mats can be considered as direct analogues to
28 I. Introduction ancient microbial mats, which is where OxyP likely evolved, and as condensed analogues of the water column in ancient stratified oceans. It is, however, crucial to identify suitable microbial mat systems distributed over geochemical/energetic gradients corresponding to the historical timeframes that we wish to explore. I investigated the phototrophic microbial mats distributed along the flow path of the Frasassi sulfidic spring water and the Little Salt Spring (LSS) mats as ancient Earth analogues. The overall aim of this thesis was to address the question of which mat-intrinsic and external factors shape the competition among functional groups of phototrophs and with other autotrophs, such as chemolithotrophic sulfur oxidizing bacteria. Specifically, we aimed for a deeper understanding of the importance of diurnal fluctuations in the physico-chemical conditions for the outcome of this competition and how the diurnal O2 budget in the analogue systems is affected. In the course of our studies on phototrophs from the sulfidic environments we found a multifaceted palette of adaptation strategies to the diurnally changing sulfidic conditions and we aimed to analyse which strategy is optimal: obligate AnoxyP, versatility allowing switching between AnoxyP and OxyP or obligate sulfide-resistant OxyP. This was linked to several questions: Is versatility the key to success in such environments? Might photosynthetically versatile cyanobacteria also have played an important role in ancient microbial mats or the water column of euxinic Proterozoic oceans? Would it have mattered? We hypothesized that the balance between photosynthetic modes plausibly has a substantial effect on other processes in the mat, such as sulfate reduction, and the overall diurnal O2 budget – independent of whether AnoxyP and OxyP are performed by different functional groups or by a single functional group, i.e., the cyanobacteria (Johnston et al., 2009). To better understand the advantage of each mode and overall photosynthetic versatility, the regulation of the transition between OxyP and AnoxyP dependent on the dynamic physico-chemical microenvironmental conditions that cyanobacteria are exposed to in microbial mats was studied in more depth. This is addressed by physiological studies on cyanobacterial isolates, and by in-situ measurements in natural cyanobacterial mats. As a synthesis of this thesis, I used the physiological data to construct metabolic models of biogeochemical cycling (O, C and S) in contemporary and ancient systems, to obtain insights into the questions raised above.
29 1. PAST AND PRESENT PHOTOSYNTHESIS
1.5. References
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37 38 Overview of Chapters
CHAPTER 2 – STRUCTURE AND FUNCTION OF NATURAL SULPHIDE OXIDIZING MICROBIAL MATS UNDER DYNAMIC INPUT OF LIGHT AND CHEMICAL ENERGY Klatt, Judith M.; Meyer, Steffi; Häusler, Stefan; Macalady, Jennifer L.; de Beer, Dirk; Polerecky, Lubos
Status: In press in The ISMEJ
CHAPTER 3 – ASSESSMENT OF THE STOICHIOMETRY AND EFFICIENCY OF CO2 FIXATION COUPLED TO REDUCED SULFUR OXIDATION Klatt, Judith M.; Polerecky, Lubos
Status: Published in Frontiers in Microbiology
CHAPTER 4 – HYDROGEN SULFIDE CAN INHIBIT AND ENHANCE OXYGENIC PHOTOSYNTHESIS IN A CYANOBACTERIUM FROM SULFIDIC SPRINGS Klatt, Judith M.; Haas, Sebastian; Yilmaz, Pelin; de Beer, Dirk; Polerecky, Lubos
Status: Published in Environmental Microbiology
CHAPTER 5 – ANOXYGENIC PHOTOSYNTHESIS AND 2-METHYL HOPANOID PRODUCTION: AMODEL CYANOBACTERIUM ISOLATED FROM A PROTEROZOIC OCEAN ANALOG Trinity L. Hamilton, Judith Klatt, Laurence M. Bird, Katherine H. Freeman, Dirk de Beer, and Jennifer L. Macalady
Status: In preparation Contribution: Microsensor measurements, data analysis, modelling, writing
CHAPTER 6 – ANOXYGENIC PHOTOSYNTHESIS CONTROLS OXYGENIC PHOTOSYNTHESIS IN A CYANOBACTERIUM FROM A SULFIDIC SPRING Judith M. Klatt, Mohammad A. A. Al-Najjar, Pelin Yilmaz, Gaute Lavik, Dirk de Beer, Lubos Polerecky
Status: Published in Applied and Environmental Microbiology
CHAPTER 6 – REGULATION OF THE PARTITIONING BETWEEN OXYGENIC AND ANOXYGENIC PHOTOSYNTHESIS IN A CYANOBACTERIUM DOMINATING SULFIDIC SPRING MICROBIAL MATS Klatt, Judith M.; Häusler, Stefan; de Beer, Dirk; Polerecky, Lubos
Status: In preparation
39 40 II. Manuscripts
2. STRUCTURE AND FUNCTION OF NATURAL SULPHIDE OXIDIZING MICROBIAL MATS UNDER DYNAMIC INPUT OF LIGHT AND CHEMICAL ENERGY
Klatt, Judith M.1*; Meyer, Steffi1; Häusler, Stefan1; Macalady, Jennifer L.2; de Beer, Dirk1; Polerecky, Lubos1,3*
1Max Planck Institute for Marine Microbiology, Celsiusstr.1, 28359 Bremen, Germany 2Pennsylvania State University, University Park, Pennsylvania, USA 3Department of Earth Sciences – Geochemistry, Faculty of Geosciences, Utrecht University, Princetonplein 9, 3584 CC, Utrecht, The Netherlands
Accepted in the ISMEJ
* corresponding authors [email protected] [email protected]
41 2. ENERGY CONVERSION IN ILLUMINATED SULPHIDIC SPRINGS
Abstract
We studied the interaction between phototrophic and chemolithoautotrophic sulphide oxidizing microorganisms in natural microbial mats forming in sulphidic streams. The structure of these mats varied between two end-members: one characterized by a layer dominated by large sulphur oxidising bacteria (SOB; mostly Beggiatoa-like) on top of a cyanobacterial layer (B/C mats) and the other with an inverted structure (C/B mats). C/B mats formed where the availability of oxygen from the water-column was limited (<5 μM). Aerobic chemolithotrophic activity of the SOB depended entirely on oxygen produced locally by cyanobacteria during high light conditions. In contrast, B/C mats formed at locations where oxygen in the water-column was comparatively abundant (>45 μM) and continuously present. Here, SOB were independent of the photosynthetic activity of cyanobacteria and outcompeted the cyanobacteria in the uppermost layer of the mat where energy sources for both functional groups were concentrated. Outcompetition of photosynthetic microbes in the presence of light was facilitated by the decoupling of aerobic chemolithotrophy and oxygenic phototrophy. Remarkably, the B/C mats conserved much less energy than the C/B mats, although similar amounts of light and chemical energy were available. Thus, ecosystems do not necessarily develop towards optimal energy usage. Our data suggest that when two independent sources of energy are available, the structure and activity of microbial communities is primarily determined by the continuous rather than the intermittent energy source, even if the time-integrated energy flux of the intermittent energy source is greater.
Keywords: Anoxygenic photosynthesis / oxygenic photosynthesis / cyanobacteria / large sulphur oxidizing bacteria / chemolithoautotrophy / aerobic sulphide oxidation / microbial mat / oxygen oases / stratification
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Introduction Life is driven by essentially two types of energy: chemical energy, in the form of chemical disequilibria, and physical energy carried by light of suitable wavelengths. On geological time-scales, changes in energy fluxes and the development of new strategies of life are closely coupled. One of the most important changes in energy availability was the evolution and proliferation of oxygenic photosynthesis. The onset of this process introduced the thermodynamically most favourable electron acceptor oxygen to the wider reduced environment. Oxygenic photosynthesis therefore provided previously unexplored thermodynamic disequilibria to other microorganisms thereby allowing for the evolution and diversification of aerobic metabolisms (Dismukes et al., 2001; Castresana & Saraste, 1995; Catling et al., 2005). Insights into the revolutionary milestones associated with the evolution and proliferation of oxygenic photosynthesis can be gained by studying modern model ecosystems in environments whose geochemical/energetic characteristics exhibit spatial gradients that are analogous to the temporal transition from a reduced to an oxidized state of the Earth's surface. The thin microbial mats forming at the bottom of light-exposed cold sulphidic springs at Frasassi, Italy (Galdenzi et al., 2008), are such an analogue system as they are distributed across a gradient from reduced to oxidized conditions in the overlying water column. These contemporary phototrophic microbial mats are of particular interest as they represent analogues to ancient cyanobacterial mats (e.g., stromatolites) that are thought to have been extensive in shallow waters throughout the Proterozoic and possibly already in the Archean (Schopf, 2012; Grotzinger & Knoll, 1999; Seckbach & Oren, 2010; Ward et al., 1992; Allwood et al., 2009). In this study, we used microsensors to quantify energy fluxes available for and conserved by the dominant processes in the mats – photosynthesis (P) and aerobic sulphide oxidation (SO) – with the aim to understand how the interaction between these processes under a fluctuating input of chemical and light energy determines the structure and function of the mats. We hypothesized that under the different conditions the mats develop towards a system that conserves the available energy optimally. We discuss the important role of energy dynamics on microbial mat structure and possible implications of our results in the context of Earth’s oxygenation.
43 2. ENERGY CONVERSION IN ILLUMINATED SULPHIDIC SPRINGS
Materials and Methods
Study site This study was performed in streams and pools forming where the sulphidic waters from the Frasassi Cave system in the Frasassi Gorge, Italy, emerge and mix with waters of the Sentino river (Galdenzi et al., 2008). The study sites included the outflows of two perennial springs, the Fissure Spring and the Main Spring (43°24’4’’N, 12°57’56’’E).
Water chemistry Sampling for bulk water chemistry analysis was done in May and September 2009 and in September 2012. Samples were taken during night and around midday within a few cm above the microbial mat patches and were preserved immediately in gas-tight vials containing a mixture of 100 μl of 20% ZnCl2 and 50 μl of saturated HgCl2 solution. The vials were filled with water samples until there was no head-space, and kept at room temperature until quantification in Bremen. Dissolved inorganic carbon (DIC) and ammonium were determined using flow injection analysis (Hall & Aller, 1992). The sum of nitrate and nitrite was quantified according to Braman & Hendrix (1989) using an NOx analyzer equipped with a chemiluminescence detector (Model CLD 66, Eco Physics, 2- - Switzerland). pH and concentrations of dissolved O2 and total sulphide (Stot = [S ]+[HS
]+[H2S]) were determined with microsensors. Temperature at the mat surface was measured with a PT1000 mini-sensor (Umweltsensortechnik, Germany).
Microscopy The dominant members of the mat community were identified by microscopy. Imaging by bright field, phase contrast and fluorescence microscopy was done using an Axiophot epifluorescence microscope (Zeiss, Germany). Photopigments were identified on a single- cell level by hyperspectral imaging (Polerecky et al., 2009).
Microsensors
O2, pH and H2S microsensors with a tip diameter of 10–80 μm and response time of <1 s were built, calibrated and used as described previously (Revsbech, 1989; Jeroschewski et al., 1996; de Beer et al., 1997). Calibration of the H2S microsensors was performed in acidified spring water (pH<2) to which NaS2 was added stepwise. The total sulphide concentrations, Stot, in the calibration solutions were determined according to Cline
(1969). Calculation of Stot from the local H2S concentrations and pH values measured
44 II. Manuscripts
with microsensors was done according to Millero (1986), using the pK1 value of 7.14- 7.17 depending on temperature (Jeroschewski et al., 1996; Wieland & Kühl, 2000).
In situ microsensor and light measurements over a diel cycle
In situ measurements of O2, pH and H2S in the mats were done using previously described microsensor setups (Weber et al. 2007, www.microsen-wiki.net). Parallel O2, pH and H2S profiles were measured under naturally variable light conditions by measuring continuously over a complete diel cycle on a cloudless day. In parallel to microsensor measurements, downwelling irradiance of the photosynthetically available radiation (PAR) was recorded using a calibrated light logger (Odyssey Dataflow Systems, New Zealand) or a calibrated scalar irradiance microprobe (Lassen et al., 1992) placed next to the microsensors. The irradiance microprobe was additionally used to quantify reflectance of the mats, as previously described (Al-Najjar et al., 2010).
Microsensor measurements under controlled light conditions Measurements under controlled light conditions were performed both in-situ and ex-situ.
In-situ we first measured steady state profiles of O2, H2S and pH in the dark and at an incident irradiance of 650 μmol photons m-2 s-1 generated by a halogen lamp (KL1500, Schott, Germany) corresponding to the natural illumination around midday.
Subsequently, volumetric rates of gross oxygenic P were determined using the O2 microsensor-based light-dark shift method of Revsbech & Jorgensen (1983). Last, the same light-dark shift approach was applied using H2S and pH microsensors instead of a
O2 microsensor, which allowed quantification of volumetric rates of gross anoxygenic P in terms of consumed Stot (Klatt et al., 2015). For this measurement the light was switched on and off over a few minute intervals while the signals were recorded in 0.3 s intervals. All of these measurements were done in the same spot of the mat. To confirm that the cyanobacterial population in the mats was able to perform simultaneous oxygenic and anoxygenic P, similar measurements were performed ex-situ.
The mat sample was placed in a temperature-controlled (15°C) flow chamber and O2, H2S and pH were measured in parallel in the same spot of the mat under variable illumination from the halogen lamp. To assess the potential role of obligate anoxygenic phototrophs that can use light in the near infrared (NIR) part of the spectrum and H2S as the electron donor for anoxygenic P, similar measurements were additionally performed using NIR
45 2. ENERGY CONVERSION IN ILLUMINATED SULPHIDIC SPRINGS
light emitting diodes (maximal emission at Ȝmax = 740 and 810 nm; H2A1 series, Roithner-Lasertechnik, Austria). The measurements with and without the additional NIR light were done before and after the addition of 5 μM of DCMU (3-(3,4-Dichlorophenyl)- 1,1-dimethylurea; Aldrich, Germany), a specific inhibitor of oxygenic P.
Calculation of fluxes and daily energy budgets
Diffusive fluxes of O2 and Stot were calculated from the measured concentration gradients multiplied by the corresponding diffusion coefficients and factors correcting for the azimuthal angle at which the profiles were measured (Berg et al., 1998; Polerecky et al., 2007). Diffusion coefficients, D, were corrected for temperature, salinity and, in case of
Stot, for the composition of the total sulphide pool as previously described (Sherwood et al., 1991; Wieland & Kühl, 2000). Specifically, diffusion coefficients were 1.78 × 10-5 2 -1 -5 2 -1 cm s for O2 and 1.35 × 10 cm s for Stot, varying by not more than 3% depending on the temperature in each particular measurement. Fluxes of incident light energy were estimated from the measured downwelling irradiance, assuming that the average energy content of photons of PAR is 217.5 kJ (mol photons)-1 (Al-Najjar et al., 2010). The fraction of the incident light energy absorbed by the cyanobacterial population in the mats was estimated from the measured reflectance of the mats (see Supplement 1). Fluxes of energy conserved by oxygenic and anoxygenic P as well as by aerobic
SO were estimated by multiplying the estimated CO2 fluxes with the molar energy required for CO2 fixation by the respective process. For oxygenic and anoxygenic P the
CO2 flux was estimated from the measured fluxes of O2 produced and Stot consumed in the photosynthetically active zone assuming the stoichiometry of O2/CO2 = 1 and Stot/CO2
= 2, respectively. For CO2 fixation coupled to aerobic SO we employed the approach of Klatt & Polerecky (2015), which allows the prediction of the complete stoichiometry of autotrophic SO based on the measured Stot/O2 flux ratios. Prerequisite for these 0 2- calculations is the relationship between the S /SO4 production ratio and the energy conservation efficiency, i.e., the ratio between the energy demand for CO2 reduction and the energy available from O2 reduction. To derive this relationship we made two main 2- assumptions: (i) When SO is in a steady state, sulphide is oxidized completely to SO4
(Jørgensen et al., 2010); (ii) When the Stot/O2 consumption ratio in the SO layer varies strongly over a diel cycle, the highest measured Stot/O2 consumption ratio occurs when 0 sulphide is oxidized incompletely to S whereas the lowest measured Stot/O2 consumption
46 II. Manuscripts
2- ratio corresponds to complete sulphide oxidation to SO4 . These assumptions enabled us 0 2- to determine the energy conservation efficiency as a function of the S /SO4 production ratio, based on which we estimated the fluxes of CO2, energy gained and energy conserved by SO from the measured fluxes of Stot and O2 at different tiimes during the diel cycle. Details of the CO2 and energy flux calculations are given in Supppplement 2. Daily budgets were calculated by integrating the instantaneous fluxes over the 24 h period for all relevant reactants and energy-harvesting processes involved.
Results
Mat structure vs. chemical composition of the spring-water The light-exposed microbial mats frrom the Frasassi sulphidic springs are dominated by two functional groups: cyanobacterria, identified by the presence of the characteristic cyannobacterial pigments chlorophyll a and phycocyanin in the cells revealed by hyperspectral microscopy (Supplementary Fig. S1), and large Beggggiatoa-like sulphur oxidizing bacteria (SOB; filamentous, containing sulphur inclusions, motile through gliding) (Fig. 1A and 1B). Inferrring from microscopy, hyperspectral imaging and reflectance measurements of the mats (Fig. S2), the abundance of phototrophs other than cyannobacteria was not significant in the studied mat layers.
Figure 1: Microscopic images of the dominant cyanobacterial and SOB morphotypes in sulphide oxidizing microbial mats from the light-exposed sulphidic streams in Frasassi, Italy, and photographs of end-member mat structures. (Scale bars: A–B: 10 μM, C: 20 cm. D: 5 cm).
A close inspection revealed thhat the structure of the mats fell eessentially between two end-members: mats with a distinnct cyanobacterial layer on top of a distinct SOB layer (Fig. 1C), and mats with SOB on top of cyanobacteria (Fig. 1D). Thesse two mat types are hereafter referred to as C/B and B/C mats, respectively. In the spring outflow streams, the distribution of these two mat types was patchy and varied on a centimetre to decimetre scale. Water chemistry and llight measurements
47 2. ENERGY CONVERSION IN ILLUMINATED SULPHIDIC SPRINGS revealed that the physico-chemical parameters directly above the mats also changed at this scale. This was mainly due to mixing of the spring water with the freshwater from the Sentino river under highly spatially variable flow conditions, which lead, for instance, to the formation of stagnant anoxic pools next to an aerated water column. The formation of a certain mat type did not correlate with the daily light dose, with temperature, pH, nitrate, ammonium, DIC, H2S or Stot concentrations in the water column during day and night, nor with the maximum H2S and Stot concentrations measured inside the mat (Table 1 and S1). However, the locations harbouring the two end-member mat types clearly differed with respect to the dissolved O2 concentration in the water column above the mats, with the C/B and B/C mats exclusively found in areas where the O2 concentration during the night was below 5 μM and above 45 μM, respectively (Table 1). Changes in the water column O2 concentration during the day, which occurred due to the high rates of oxygenic P in the mats when the overlying water column was stagnant, had no effect on the formation of a certain mat type (compare data for mats C/B-1 and C/B-2 in Fig. S3 and Table S1).
Photosynthetic activity of the cyanobacterial community
Laboratory microsensor measurements revealed light-induced production of O2 and consumption of H2S in the cyanobacterial layer of C/B (Fig. 2) and B/C mats (data not shown). While O2 production was due to oxygenic P, removal of H2S could be caused by three processes: chemical or biologically mediated oxidation with the photosynthetically - 2- produced O2, a shift in the equilibrium of sulphide speciation (H2S, HS and S ) induced by a pH increase associated with the uptake of DIC by oxygenic P, or by anoxygenic P that uses H2S as the electron donor. First, H2S oxidation with O2 could be excluded because the H2S dynamics changed abruptly upon light-dark transitions and were independent of O2 concentrations. Additionally, the depth-integrated rate of gross oxygenic P matched the net rate derived from steady state diffusive profiles (see below), showing that there is negligible oxygen reduction activity (such as due to the reaction with H2S) in the photosynthetically active layer. Second, assuming that total sulphide concentrations, Stot, stayed constant during the light-dark transitions and pH varied as measured (by about 0.005 pH units; Fig. 2), the calculated variation in H2S would be about 1-2 orders of magnitude lower than measured (data not shown). Therefore, we exclude also the second possibility and conclude that the measured light-induced variation in H2S was a direct result of anoxygenic P in the cyanobacterial layer of the mat.
48 II. Manuscripts
T [°C] [°C]
) -1 s -2 6 93 14 14 93 133 14 14 133 124 13 124 PAR a (SOB) layer, as derived from from derived layer, as a (SOB) n in Table S2. S2. n in Table S2. mol photons m mol ȝ
( ht. ht.
3,5 x 60 60 M] 3.6 3.6 34.4 34.4 NO μ [ (+/- 1.8) (+/- 1.9) (+/- 0.07)
+ 3,5 4 56 56 77 77 M] 206 206 NH μ [ (+/- 4.7) (+/- 3.3) (+/-10.2)
2 8.0 8.0 7.3 - - 7.3 - 163 13 8.0 8.0 7.5 7.5 pH
average photon flux during 15 h of lig during average photon flux from samples taken during the day are show during taken samples from
3,5 3.9 3.9 4.7 4.7 4.2 4.2 4.4 4.4 DIC [mM]
(+/- 0.6) (+/-0.11) (+/-0.18) (+/-0.15) and layer dominated by large sulphur oxidizing bacteri oxidizing sulphur by large layer dominated and
umn and in the studied mats. 2 night. The values obtained during the day are shown in Table shown day are obtained during the values The night. S concentrationS and pH. 2 2 5 0 5 8.9 6.9 150 < 1 78 14.5 < 150 6.9 5 8.9 M] 65 65 45 45 O μ [
4
36 36 73 73 95 95 510 510 435 435 (88) (88) (353) (353) (681) (681) (175) (3339)
given as the (PAR), available radiation inside the cyanobacterial (C) cyanobacterial the inside ) 1 2 tot SOB s taken during night. The values determined determined The values during night. s taken 4
M] 61 34 80 - 8.3 < 1 50 50 1 - 15 80 34 61 < - 8.3 36 36 81 81 345 345 415 415 125 125 μ S (S (287) (287) (149) (149) (642) (642) (390) (390) [ 2 (2755) (2755) H C 4
(n = 3 – 6). deviations andard
, was calculated from the measured H the from calculated , was 16 16 23 23 73 73 50 50 89 89 401 401 (92) (92) (248) (248) (151) (582) (628) (278) (278) tot
Main Spring Fissure Spring
Concentrations in the water column (W) and (W) water column Concentrations in the C/B-3 Fissure Spring C/B-3 Fissure B/C-3 C/B-2 Main Spring C/B-2 Main C/B-1 Main Spring C/B-1 Main Mat type Site Spring B/C-2 Main B/C-1 W Total concentration, S sulfide photosynthetically Downwelling irradiance of the Values are derived from microsensor measurements performed during performed measurements microsensor from derived are Values water sample from determined Values were to st refer parentheses in Values Table 1: Physico-chemical parameters in the overlying water col 1 2 3 4 6 5 measurements. microsensor
49 2. ENERGY CONVERSION IN ILLUMINATED SULPHIDIC SPRINGS
This conclusion was confirmed by ex situ measurements in a C/B mat treated with
DCMU, a specific inhibitor of oxygenic P but not of anoxygenic P. Specifically, H2S concentrations in the DCMU-treated mat decreased and increased upon the addition and removal of light, respectively, while the effect on pH was marginal (data not shown) and
O2 was below detection limit (Fig. 3A). For the incident irradiance of 100 μmol photons m-2 s-1 the volumetric rates of anoxygenic P derived from light-dark shift microsensor -3 -1 measurements ranged between 1–4 mmol Stot m s (Fig. 3B) and their depth-integrated -2 -1 value (1.53 μmol Stot m s ) closely matched the flux of Stot removed by the cyanobacterial anoxygenic P derived from the steady state microprofiles (1.55 μmol Stot m-2 s-1). This implies that anoxygenic P was the only significant sink of sulphide in the cyanobacterial layer.
7.100 10 I=30 dark I=30 134 240
9 132 235 8 130 pH 7.095 230 (μM) tot S (μM)
7 128 2 S H
Oxygen (μM) Oxygen 6 126 225
5 124 7.090 220 0 5 10 15 20 time (s)
Figure 2: Light-induced dynamics of O2, H2S, Stot and pH inside the cyanobacterial layer of a freshly collected C/B mat. Total sulphide concentrations, Stot, were calculated from the measured -2 -1 H2S concentrations and pH. The incident irradiance was 30 μmol photons m s . Distance between sensor tips was about 5 mm.
The observed light-induced variability of Stot could also be due to the activity of obligate anoxygenic phototrophs. The activity of such phototrophs is, however, expected to be affected by the local oxygen concentration. For example, filamentous anoxygenic phototrophs (also known as green non-sulphur bacteria) can switch from sulphide-driven anoxygenic P to photoorganoheterotrophy or aerobic respiration when oxygen and cyanobacterial excudates are available in the light (van der Meer et al., 2005; Polerecky et al., 2007). Additionally, anaerobic anoxygenic phototrophs, such as green and purple sulphur bacteria, are expected to be poisoned by high oxygen concentrations.(van Gemerden & Mas, 1995). In our measurements, however, the net and gross rates of anoxygenic P before the addition of DCMU to the mat sample, i.e., in the presence of oxygenic P, were almost equal to those measured after inhibition of oxygenic P by DCMU (data not shown), suggesting that obligate anoxygenic phototrophs were not significantly contributing to the light-driven sulphide consumption in the cyanobacterial
50 II. Manuscripts
layer. This was confirmed by the observation that exposure to near infrared light, which specifically targets bacteriochlorophylls of obligate anoxygenic phototrophs, did not induce sulphide consumption (Fig. S4).
O2 (μM) 0 5 10 15 20 -1.0 -1.0 A B -0.5 -0.5
0.0 0.0
0.5 0.5 depth (mm) 1.0 1.0
1.5 1.5
2.0 2.0 0 200 400 600 -5 -4 -3 -2 -1 0 H S (μM) S (μM) -1 2 tot Anoxygenic P (μM Stot s )
Figure 3: Ex situ microsensor measurements in a C/B mat after the addition of DCMU. (A) Steady-state depth profiles of O2 and H2S and Stot measured in the light (open symbols) and in the dark (filled symbols). (B) Depth profile of volumetric rates of anoxygenic photosynthesis, as derived from the light-dark shift method adapted for Stot. For both panels, total sulphide concentrations, Stot, were calculated from the measured H2S concentrations and pH, and the incident irradiance during the light measurements was 100 μmol photons m-2 s-1.
Together these data show that the cyanobacterial community in the studied mats is able to perform oxygenic and sulphide-driven anoxygenic P simultaneously and that cyanobacterial anoxygenic P is fully responsible for the measured light-induced H2S variation in the cyanobacterial layer of the mats.
Activity of C/B mats Continuous in situ micro-profiling in C/B mats under naturally variable illumination gave consistent results with those obtained in the laboratory. In the dark, sulphide consumption rates were minute or below detection limit, as implied by essentially flat Stot profiles across the mats during the night, and O2 was not detected (Supplementary Fig. S3). At sunrise, while still at low light, the rate of sulphide removal due to anoxygenic P in the cyanobacterial layer sharply increased, resulting in a gradual decrease in H2S concentrations (Fig. 4A). Around midday, when the incident irradiance exceeded about 300 μmol photons m-2 s-1, sulphide concentrations within the cyanobacterial layer decreased and the photosynthetic production of oxygen sharply increased (Fig. 4A). These dynamics occurred essentially in reverse order at sunset.
51 2. ENERGY CONVERSION IN ILLUMINATED SULPHIDIC SPRINGS
) )
-1 500 -1
) 500 -1
s B/C mat, C-layer s 3.5 A C/B mat, C-layer 3.5 C -2 -2 s -2 m m 3.0 3.0 400 tot tot m 400 2 ol ol 2.5 2.5 ol 300 300 (μM)
(μ 2.0 (μ 2.0 S S (μM) (μ 2 2 H
1.5 200 1.5 200 H 1.0 1.0 100 100 0.5 0.5 OxygenicP O AnoxygenicP S AnoxygenicP S 0.0 0 0.0 0 23:00 02:00 05:00 08:00 11:00 14:00 17:00 20:00 19:00 22:00 01:00 04:00 07:00 10:00 13:00 time of the day (hh:mm) time of the day (hh:mm)
3.0 3.0
) 3.5 C/B mat, SOB-layer )
) 3.5 B/C mat, SOB-layer B ) D -1 -1 -1 -1 s s s 2.5 s
-2 2.5 -2 -2 3.0 -2 3.0 m m m m 2 2 tot 2.5 2.0 tot 2.5 2.0 O O S S ol ol ol 2.0 ol 2.0
1.5
1.5 2 1.5 2 /O 1.5 /O tot 1.0 1.0 tot S flux (μ S flux (μ flux (μ flux (μ 2 2 tot 1.0 tot 1.0 O O S 0.5 S 0.5 0.5 0.5
0.0 0.0 0.0 0.0 23:00 02:00 05:00 08:00 11:00 14:00 17:00 20:00 19:00 22:00 01:00 04:00 07:00 10:00 13:00 time of the day (hh:mm) time of the da y (hh:mm)
Figure 4: Activity of the cyanobacterial and SOB populations in the C/B and B/C mats exposed to natural light fluctuations over a 24 h period. The fluxes were derived from in situ microsensor profiles of O2 and H2S concentration (Supplementary Figs. S5 and S6) and pH. The corresponding average H2S concentration in the cyanobacterial layer, and the Stot/O2 flux ratio in the SOB layer, are also shown. Open and shaded areas correspond to light and dark, respectively.
In situ measurements under controlled light conditions revealed additional insights into the activity of C/B mats. At high incident irradiance (650 μmol photons m-2 s-1), the cyanobacterial community performed both oxygenic and anoxygenic P simultaneously, as shown by the light-dark shift measurements (Fig. 5B). Aerobic respiration in the photosynthetically active zone was negligible, as the net diffusive flux of oxygen from the zone was very similar to the gross rates of oxygenic P depth-integrated over the zone (2.43 and 2.47 μmol m-2 s-1, respectively). Although oxygenic P was detectable at depths 0-0.6 mm, anoxygenic P was detectable only at depths 0-0.4 mm, i.e., in zones of the cyanobacterial layer where both H2S and light were abundant. The overlap between the zone of anoxygenic photosynthetic activity and the decrease in Stot from the overlying water indicated that the sulphide used by the cyanobacteria originated exclusively from the overlying water. This was supported by the close match between the depth-integrated -2 -1 rates of gross anoxygenic P (1.56 μmol Stot m s ) and the downward Stot flux from the -2 -1 water column (1.55 μmol Stot m s ) (Fig. 5A and 5B). When the oxygenic P was, however, not active (i.e., in the morning and later in the afternoon), the flux of Stot
52 II. Manuscripts
consumed by anoxygenic P had significant contributions from both the downward and upward components of the diffusive Stot flux.
-1 O2 (μM) Oxygenic P (μM O2 s ) 0 50 100 150 200 250 -6 -4 -2 0 2 4 6 A B -0.5 A B C/B mat
0.0 water column
Cyanobacteria 0.5
SOB 1.0
depth (mm)
1.5
2.0 0 200 400 600 -6 -4 -2 0 2 4 6 -1 Stot (μM) Anoxygenic P (μM Stot s ) 6.8 7.0 7.2 7.4 pH
-1 O2 (μM) Oxygenic P (μM O2 s ) 0204060 -6 -4 -2 0 2 4 6
-0.5 C D B/C mat water column 0.0 SOB
0.5 Cyanobacteria
1.0
depth (mm)
1.5
2.0 0 25 50 75 100 125 150 -6 -4 -2 0 2 4 6 -1 Stot (μM) Anoxygenic P (μM Stot s ) 6.85 6.90 6.95 pH Figure 5: Microsensor measurements in the C/B and B/C mats under artificially controlled light conditions. Measurements in the C/B (panels A and B) and B/C mats (panels C and D) were performed in-situ and ex-situ, respectively. Left panels show steady-state depth profiles of O2, Stot and pH in the light (open symbols) and in the dark (closed symbols). Right panels show the corresponding depth profiles of volumetric rates of oxygenic and anoxygenic photosynthesis, as derived from light-dark shift microsensor measurements of O2 and Stot, respectively. For all panels, total sulphide concentrations, Stot, were calculated from the measured H2S concentrations (not shown) and pH. The incident irradiance during the light measurements in the C/B and B/C mats was 650 and 710 μmol photons m-2 s-1, respectively. For orientation, approximate structures of the mats are also shown.
Sulphide consumption in the SOB layer was detectable only after the onset of oxygenic P by the cyanobacteria (Figs. 4B and 5A). This observation is consistent with the absence of an external supply of an electron acceptor, i.e., O2, from the water column, - and additionally indicates that anaerobic sulphide oxidation with NO3 as the electron acceptor was not important. Once O2 became available, sulphide consumed by the SOB
53 2. ENERGY CONVERSION IN ILLUMINATED SULPHIDIC SPRINGS originated exclusively from below, while the sulphide entering the mat from above was consumed by the cyanobacteria (see above). Analysis of the profiles in Fig. 5A revealed that the photosynthetically produced -2 -1 O2 (downward flux 0.85 μmol O2 m s ) and the sulphide supplied from below (upward -2 -1 flux 0.48 μmol Stot m s ) were consumed at a Stot/O2 flux ratio of about 0.6. However, a detailed analysis of the complete sets of profiles measured in the C/B mat over a diel cycle (Fig. S3) revealed that the Stot/O2 consumption ratio changed substantially over the course of the day, reaching about 2 after O2 became available, gradually decreasing to about 0.7 before increasing back to about 2.6 just before O2 depletion towards the night
(Fig. 4B). Similar patterns in the diurnal variations of the Stot/O2 consumption ratio between 0.7 and 2.6 were also observed in a replicate C/B mat (Supplementary Fig. S5A– B).
The observed diurnal variation in the Stot/O2 consumption ratio in the SO zone can be explained by assuming that the SOB activity varied between incomplete oxidation of 0 2- sulphide to S and complete oxidation of sulphide to SO4 when the Stot/O2 ratio reached the maximum and minimum value, respectively. This assumption made it possible to estimate the energy conservation efficiency of the SOB community. Specifically, using 0 the maximum value of Stot/O2 = 2.6 (oxidation to S ) and additionally taking into account the measured local concentrations of the substrates and products involved, we found the efficiency values of 16.6% and 16.5% for the two replicate C/B mats. Conversely, the 2- minimum value of Stot/O2 = 0.7 (complete oxidation to SO4 ) measured in the respective mats gave the efficiency values of 17.2% and 17.1% (calculation details given in Supplement 2.2). These calculations suggest that the energy conservation efficiency of the SOB community in the studied mats was around 16.9% and did not significantly depend 0 2- on the S /SO4 production ratio.
The Stot/O2 consumption ratio in both replicate C/B mats was significantly 2 negatively correlated with the downward O2 flux from the cyanobacterial layer (R =0.91, 2 P<0.0001) as well as with the local concentration of O2 (R =0.91, P<0.0001) and H2S 2 2 (R =0.66, P=0.008), but not correlated with the upward Stot flux (R =0.27, P=0.15; Supplementary Fig. S6). In one of the C/B mats the availability of sulphide from below was low and thus the minimum Stot/O2 consumption ratio of 0.7, which indicates complete oxidation of sulphide to sulphate (see above), was reached early during the day (Fig. S5B). When this ratio was reached we observed a pronounced downward shift of the zone where sulphide was aerobically consumed (Fig. S5C). This downward shift did not affect
54 II. Manuscripts
the correlation between the Stot/O2 consumption ratio and the O2 flux or the local O2 and
H2S concentrations. Taken together, our data suggest that in the layer underneath the photic zone aerobic sulphide oxidation by SOB was the only significant sink of oxygen in the layer while aerobic heterotrophic activity was negligible. Therefore, we conclude that the SOB population in the C/B mats most likely varied its activity between incomplete and 0 2- complete aerobic sulphide oxidation to S and SO4 , respectively (i.e., at a variable 0 2- S /SO4 production ratio), whereby this variability was regulated by the availability of O2 produced locally by the overlying cyanobacteria and by the local H2S concentration but not by the flux of sulphide from below.
Activity of B/C mats Despite the fact that B/C mats were exposed to similarly high incident light intensities as the C/B mats (Table 1), sulphide removal in these mats due to anoxygenic P was not very pronounced (Fig. 4C). Also oxygenic P was very low or even below detection limit in the B/C mats (Fig. 4C; Fig. S7). This was confirmed by ex situ measurements under controlled light conditions. Specifically, the depth-integrated rates of gross anoxygenic and oxygenic P were about one order of magnitude lower than in C/B mats at similar irradiances, and remained low even when the applied irradiance exceeded the maximal values measured in-situ (Fig. 5D; Fig. S7). Additionally, the rate of O2 consumption in the cyanobacterial layer (such as by aerobic respiration), calculated as the difference between the gross and net rates of O2 produced in the layer (Fig. 5C and 5D), was not significant in the B/C mats. Also, the depth-integrated rates of gross anoxygenic P (0.20 -2 -1 -2 -1 μmol Stot m s ) matched closely the net Stot flux (0.19 μmol Stot m s ) consumed in the cyanobacterial layer, indicating that anoxygenic P was the only significant sink of sulphide in this layer. The source of this photosynthetically removed sulphide was exclusively from below. Sulphur and oxygen cycling in B/C mats was dominated by the light-independent aerobic chemolithotrophic SO in the top SOB layer (Fig. 4D, Fig. 5C). In the dark, sulphide for aerobic SO originated from both the water column and the underlying sediment. During high light conditions, the downward sulphide flux from the water- column was the main source, while the upward sulphide flux was mostly consumed in the cyanobacterial layer (see above; Fig. 5C). Additionally, during the maximum incident irradiance, consumption of O2 produced locally by the underlying cyanobacterial layer
55 2. ENERGY CONVERSION IN ILLUMINATED SULPHIDIC SPRINGS
only contributed less than 15% to the overall O2 consumption in the SOB layer. Thus, SO in the top layer relied mostly on O2 supplied externally from the overlying water column.
Over the course of the day, the Stot/O2 consumption ratio remained constant at about 0.67 (Fig. 4D). Thus, aerobic SO by the SOB community in the B/C mats was in steady state and therefore most likely proceeded completely to sulphate (Nelson et al., 1986; Jørgensen et al., 2010). Using this stoichiometry, we estimated the energy conservation efficiency of the SOB community in the two replicate B/C mats to be 16.4 and 17.1% (calculation details given in Supplement 2.2), which is very similar to the value estimated for the SOB community in the C/B mats (see above).
Light absorption by the cyanobacteria C/B and B/C mats back-reflected 4.5% and 81% of the incident irradiance, respectively. This suggests that the estimated fractions of the incident flux of light energy absorbed by the cyanobacterial populations were 95.5% and 19%, respectively (see Supplement 1). However, these values likely define the upper limits of the light energy utilized by cyanobacteria in the mats, since light absorption could occur also due to abiotic components in the mats (Al-Najjar et al., 2012).
Carbon and energy budgets Assuming that photosynthesis by cyanobacteria and aerobic sulphide oxidation by SOB were the only processes responsible for oxygen and sulphide cycling, we used the measured fluxes of light, oxygen and sulphide together with the estimated energy conservation efficiency of the SOB populations to estimate daily carbon and energy budgets in the two studied mat types (Table 2). The C/B mats conserved about 2.42% of the incident light energy directly by P (split into 1.83% by oxygenic and 0.59% by anoxygenic P). This conservation efficiency was increased to about 2.54% when additionally considering that the photosynthetically produced oxygen is used for SOB- driven carbon fixation coupled to aerobic chemosynthetic SO. When expressed in terms of fixed carbon, the exploitation of the thermodynamic disequilibrium between O2 and sulphide, which was internally generated by oxygenic P, thus increased the total primary productivity in the C/B mats by 15% (from 67.5 to 79.5 mmol C m-2 d-1).
56 II. Manuscripts
(0.12%; 0.25%) (0.12%; oxidation (SO)
5.41 (31.43%; 68.6%) 5.41 (31.43%; -23.38 (18.5%; 81.5%) -23.38 (18.5%; 1477.34 (98.5%; 1.5%) 1477.34 (98.5%; 0.37%
-8.63 -37.67 -46.30 -8.63 -37.67 1455.74 21.68
1.70 (100%; 0%) 1.70 (100%; 3.71 0.12% (0.12%; 0) 17.1% -4.32 (100%; 0%) -4.32 (100%; -19.06
for this process. ergy available (2.40%; 0.14%) (2.40%; osynthesis (P, anoxygenic and oxygenic) and by aerobic sulphide 40.34 (94.5%; 5.5%) 40.34 (94.5%; consumed, respectively. consumed,
15.1%) -79.53 (84.9%; 2.54% 1585.75 (99.2%; 0.8%) 1585.75 (99.2%;
3
to represent also the en sumed nd is produced and nd is
P. oxygenic of as a consequence ecosystem the 43.93 -34.11 9.82 0.0 -56.30 -56.30 -47.12 -32.28 -79.34 -47.12 -32.28 1572.53 13.22
P (anoxyP;oxyP) SO Total (PS; SO) P (anoxyP;oxyP) SO Total (P; SO) 38.10 (24.4%; 75.6%) 75.6%) 38.10 (24.4%; 2.24 65.1%) -67.49 (34.9%; -12.04 used when the compou used divided by energy available. available. by energy divided
2.42% (0.59%; 1.83%) 16.9% 4
2
)
) -1 ) -1 -1 d d -2 d -2 -2 m
2
m ) ) m 1
2
-1 -1 tot 1 1 d d
-2 -2 flux 2 flux flux 2 tot mmol O mmol S mmol CO Energy gained by aerobic sulphide oxidation (SO) was as Energy gainedby aerobic(SO) sulphide oxidation ( Energy flux available S ( ( (kJ m (kJ m (kJ Conservation efficiency CO O C/B mat B/C mat Energy flux conserved Positive and negative values are Positive and within internally generated energy Represents energy conserved Calculated as 3 in the two end-member mat types. Table 2: Daily budgets of compounds and energy utilized by phot 1 2 4
57 2. ENERGY CONVERSION IN ILLUMINATED SULPHIDIC SPRINGS
In contrast, the B/C mats conserved only about 0.12% of the incident light energy by P (mainly by anoxygenic P), although the daily flux of light energy available for both mat types was similar. Instead, energy conservation in the B/C mats was dominated by chemosynthesis (about 69% of the total energy conserved), although the daily flux of light energy available to the system was about 67-fold larger than the flux of chemical energy (in the form of oxygen and sulphide diffusing from the water-column, utilized by aerobic SO). Thus, due to the small contribution of P, the overall energy conversion efficiency in the B/C mats was only 0.37%, i.e., 7-fold lower than in the C/B mats. This grossly inefficient utilization of the available light energy in the B/C mats is also reflected in the estimated overall primary productivity, which was about 3-fold lower than in the C/B mats (23.4 vs. 79.5 mmol C m-2 d-1; Table 2). Analysis of the replicate measurements led to similar conclusions although with slightly different numerical values (Supplementary Table S2).
Discussion The dominant functional groups in the Frasassi spring mats, photosynthetic cyanobacteria and aerobic chemolithoautotrophic SOB, are directly coupled as both oxygen and sulphide are involved in their energy generation pathways. The two functional groups, however, depend on different energy sources, i.e., on light and chemical energy, respectively. Our results show that activity in the mats is driven by both energy sources. However, depending on the direction and temporal dynamics of the energy supply, the mats stratified in essentially two distinct structures, C/B mats and B/C mats, characterized by substantially different activity patterns and, most strikingly, utilization efficiencies of the externally available energy.
C/B mats In C/B mats, cyanobacteria inhabiting the photic layer switched between anoxygenic and oxygenic P over a diurnal cycle, with microzones performing oxygenic and anoxygenic P simultaneously (Figs. 4 and 5). Since C/B mats formed in areas characterized by a very low O2 concentration (<5 μM) in the overlying water (Table 1), oxygenic P was the exclusive provider of electron acceptor for aerobic SO. Due to the fluctuating availability of light the supply of oxygen to the SOB population residing under the cyanobacterial population was also fluctuating. A possible adaptation of SOB to live under such conditions is to rapidly adjust their SO
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0 2- stoichiometry, i.e., to vary the S /SO4 product ratio, depending on the availability of O2 and Stot, whereby the range of Stot/O2 consumption rations that can be covered by the SOB through the variation between the complete and incomplete oxidation of sulphide to sulphate and zero-valent sulphur, respectively, is determined by the corresponding energy conservation efficiency (Klatt & Polerecky, 2015). Our data suggest that this strategy was adopted by the SOB in the C/B mats. Specifically, we estimated that the dominant SOB performed aerobic SO with an energy conservation efficiency of ~16.9%, which allowed them to adjust their Stot/O2 consumption ratio to the range of Stot/O2 flux ratios imposed by the environment (between ~0.7 and ~2.6) and thus harvest the available fluxes of sulphide and oxygen optimally.
Interestingly, when the Stot/O2 consumption ratio reached the minimum value of 0.7, i.e., when sulphide oxidation proceeded entirely to sulphate, a further increase in the rate of oxygen supply by oxygenic P led to a downward migration of the SOB. We suggest that this is because the SOB could not adjust the Stot/O2 consumption ratio below the minimum value determined by their energy conservation efficiency (Klatt & Polerecky, 2015), leaving the adjustment of their position as the only option to maintain optimal utilization of the available substrates in the dynamically changing gradients of sulphide and O2. There are two plausible hypotheses concerning the exact trigger for migration: (i) the biomass-dependent maximum rate of O2 consumption was reached, or
(ii) the rate of O2 consumption by the SOB became limited by the supply of H2S. In both cases any further increase in oxygenic P would lead to an increase in the local O2 concentration triggering downwards migration due to a phobic response to O2 (Møller et al., 1985).
Intriguingly, the Stot/O2 consumption ratio was only determined by the O2 flux and the local H2S concentration but not by the Stot flux (Fig. S6). This is consistent with the 0 2- expectation that SOB should adjust their S /SO4 production ratio so as to maximize utilization of the available O2. This is because the carbon yield per oxygen is expected to 0 2- be almost constant irrespective of the S /SO4 production ratio, i.e., the O2 reduction rate directly translates into growth, as suggested by fact that the energy conversion efficiencies estimated for the incomplete and complete SO in the SOB inhabiting the C/B mats were almost equal (Klatt & Polerecky, 2015). 0 2- Together, the flexible O2-dependent adjustment of the S /SO4 production ratio seems to allow the SOB to efficiently exploit the chemical energy in the system and to additionally build up storage of intracellular S0 that might serve as an electron acceptor
59 2. ENERGY CONVERSION IN ILLUMINATED SULPHIDIC SPRINGS
for anaerobic respiration during night when the O2 is not available (Schwedt et al., 2011). The dominant SOB therefore appear to be highly adapted to co-exist with oxygenic phototrophs. Overall, under the oxygen-limited conditions in C/B mats, both photosynthetic and aerobic chemosynthetic activity are regulated by light energy supplied to the system.
Despite the dependence of both anoxygenic P and chemolithotrophy on H2S as an electron donor, its light-dependent depletion was not disadvantageous for the SOB because the chemolithotrophic and phototrophic layers did not compete for the same sulphide pool. Specifically, chemolithotrophic activity was exclusively supplied by the sulphide flux from below while the photosynthetically oxidized sulphide originated from the overlying water column. Aerobic SO by SOB, or for that matter any aerobic activity, was directly coupled to both oxygenic and anoxygenic P, the latter required to locally deplete the reductant (H2S) so as to enable the former. Therefore, both anoxygenic and oxygenic photosynthetic activities were beneficial for the SOB, as they together allowed for the life-enabling production of oxygen required for the SOB to thrive under the particular oxygen limiting conditions where C/B mats developed.
B/C mats In B/C mats, SOB formed a layer on top of the cyanobacteria, where they could access a continuous supply of energy, H2S and O2, from the water column throughout the entire diel cycle (Fig. 4D). In contrast, the energy source (light) for the cyanobacteria in the B/C mats was discontinuous. We hypothesize that the continuity of the energy supply was the main advantage that the SOB had over cyanobacteria in the B/C mats. Specifically, in the absence of light during the night the cyanobacteria were not triggered to assert themselves in the uppermost position of the mat, i.e., closest to the energy sources for both functional groups during the day. This has been taken advantage of by the SOB, whose chemolithotrophic activity in this position could continue uninterrupted. During the day the position of the cyanobacteria remained in the lower part of the mat, where the light availability was significantly reduced (at least 5-fold) due to intense back-scattering in the overlying layer of SOB (Supplement 1). Therefore, the cyanobacteria were not able to harvest the light optimally. It remains unclear how exactly the very distinct layering was maintained during the day and why the cyanobacteria were not able to invade the SOB layer. An important factor was probably faster motility of the SOB that made them more successful competitors for the 'prime spot' on top of the mats.
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Importance of energy flux direction and dynamics for mat structure and function When oxygen provided externally from the water column was limiting, light availability regulated the activity and spatial organization of the dominant functional groups, leading to the formation of C/B mats. In these mats primary productivity was exclusively driven by light. Namely, light energy was directly utilized by anoxygenic and oxygenic P and indirectly facilitated productivity of the SOB that exploited the chemical energy in the form of the thermodynamic disequilibrium driven by photosynthetically produced O2. Consequently, processes stratified predictably according to the direction and magnitude of the energy source, and the dominant functional groups, photosynthetic cyanobacteria and aerobic SOB, beneficially interacted. This led to an efficient utilization of the available external energy. B/C mats, on the other hand, were exposed to two energy sources, both of which had the same direction, i.e., they were externally supplied from the water column, and were therefore most abundant at the upper surface of the mat. One of these sources, namely chemical energy in the form of oxygen and sulphide, was continuous, while the other, light, was diurnally fluctuating. As the functional groups competed for the space closest to their energy sources, specific adaptation mechanisms (e.g., motility) and phenotypic features (e.g., light-scattering S0 globules) gained in importance. Intriguingly, the organisms specialized in utilizing chemical energy, SOB, outcompeted all photosynthetic microbes from the position closest to the light, even though the availability of light energy per day was orders of magnitude higher than that of chemical energy. This means that the continuously available chemical energy was used preferentially, while the additional potential for gaining oxygen via internal recycling of the other available external energy source (light) was neglected, or even suppressed. As a consequence, the competition for a favourable position in the mat led to a comparatively inefficient use of the bulk external energy. It is likely that also nitrogen retention, storage and removal in the two mat types strongly differ due to the differences in the O2 concentrations in the water-column overlying the C/B and B/C mats and the differences in the activity of the SOB and cyanobacteria in the mats. To understand possible effects of this on the mat stratification, more detailed measurements of bioavailable nitrogen in the mats would be required, which could, however, not be done during this study due to technical limitations. Nevertheless, the lack of correlation between the mat type and the water-column
61 2. ENERGY CONVERSION IN ILLUMINATED SULPHIDIC SPRINGS
- + concentrations of NO3 and NH4 (Table 1) suggests that nitrogen cycling is an unlikely factor that determines the formation of a certain mat type.
Implications The scenario in C/B mats resembles what is thought to have been the arena for the evolutionary transition from anoxygenic to oxygenic P in the ancient phototrophic microbial mats in isolated shallow-water environments. The environment where these critical steps in evolution occurred was chemically reduced (e.g., Tice & Lowe, 2004; Sessions et al., 2009; Lyons et al., 2014). Anoxygenic P using the available reductants, e.g. H2S, is expected to be an important process in such environments. However, H2S can become locally depleted, which provides a selective advantage to cyanobacteria that are able to switch from anoxygenic to oxygenic P (Cohen et al., 1986; Jørgensen et al., 1986; Klatt et al., 2015) and that are therefore never limited by electron donors. In C/B mats this versatility leads to the introduction of aerobic hotspots in the otherwise reduced environment. The C/B mats also demonstrate the revolutionary consequences of creating such aerobic hotspots. Oxygen as the most favourable electron acceptor offers a myriad of thermodynamic strategies for other microorganisms and oxygenic P has therefore set the basis for the co-evolution of aerobic organisms. Initially, as the proliferation of oxygenic P and aerobic metabolisms took place in otherwise anoxic and reduced environments, these two processes were spatially and temporarily closely coupled. This is illustrated in C/B mats, where the aerobic SO was completely dependent on the internal conversion of light energy into chemical energy driven by the photosynthetic production of oxygen. Despite the substantial consumption of oxygen by aerobic chemolithotrophy in the layer underneath the photic zone, C/B mats were net sources of oxygen to the water column. It is assumed that in ancient, more reduced oceans, excess reductants have initially scavenged the photosynthetically produced oxygen, and it was only upon depletion of these sinks that oxygen could persistently accumulate in the atmosphere and upper water column of the oceans during the Great Oxidation Event (GOE). However, the GOE was possibly predated by transient accumulations of oxygen in the atmosphere (the ‘whiffs of oxygen’) (e.g. Anbar et al., 2007; Lyons et al., 2014) and it is not unlikely that, despite reductant availability, oxygen had also persistently accumulated in the shallow water column above microbial mats, similarly to the scenario in the Frasassi sulphidic springs. Independent of the exact timing, water column oxygenation freed aerobic
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organisms from consortia and microenvironments where O2 was provided exclusively through the close proximity to oxygenic phototrophs (i.e., oxygen oases; e.g. Buick 2008). A paradoxical consequence of this is strikingly demonstrated in the Frasassi B/C mats. Although these mats were exposed to light and contained significant populations of oxygenic phototrophs, they became a net sink for oxygen once it accumulated in the water column. This is because they were mainly driven by an aerobic, light-independent process (SO). This demonstrates that an aerobic metabolism that had, by necessity, evolved dependent on light, could even outcompete the oxygenic phototrophs that formerly served as the exclusive provider of oxygen for its energy metabolism. The oxygenation of the Earth’s atmosphere represents an event where oxygen has evolved from a fluctuating internal source of chemical energy into a continuous external source. We suggest that aerobic chemosynthesis might have become competitive against photosynthesis, thus tempering photosynthesis-driven primary productivity and providing a negative feedback on the proliferation of oxygenic phototrophic organisms. As demonstrated by the Frasassi mats, the effect of this negative feedback could have been as radical as a turn from a hugely productive ecosystem that acts as a net O2 source (C/B mat) into a considerably less productive ecosystem that acts as a net O2 sink (B/C mat). Further research is required to identify possible biosignatures of this competitive interaction between phototrophs and aerobic chemolitotrophs that could be searched for in geological records to explore whether the negative-feedback hypothesis could be generalized beyond the highly localized scale of this study.
Acknowledgements We thank Daniel S. Jones (University of Minnesota) for invaluable help in the field, Alessandro Montanari and Paula Metallo for providing laboratory facilities and enjoyable atmosphere at the Osservatorio Geologico di Coldigioco, and Tim Ferdelman for fruitful discussions. We thank the technicians of the microsensor group for microsensor construction. This work was financially supported by the Max-Planck-Society, by a 2011 “For Women in Science Award” to JMK, and by NASA Astrobiology Institute (PSARC, NNA04CC06A) and Hanse-Wissenschaftskolleg Fellowship funds to JLM.
Conflict of interest statement The authors declare no conflicts of interest.
63 2. ENERGY CONVERSION IN ILLUMINATED SULPHIDIC SPRINGS
Supplementary information is available at The ISME Journal’s website.
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STRUCTURE AND FUNCTION OF NATURAL SULPHIDE OXIDIZING MICROBIAL MATS UNDER DYNAMIC INPUT OF LIGHT AND CHEMICAL ENERGY
Klatt, J. M.; Meyer, S.; Häusler, S.; Macalady, J.L.; de Beer, D.; Polerecky, L.
Supplementary Information
Content
1. Absorption of light energy by the cyanobacterial population in the studied microbial mats
2. Conversion of fluxes of O2, Stot and light to fluxes of fixed CO2 and conserved energy
2.1. Photosynthesis (P)
2.2. Aerobic chemolithoautotrophic sulphide oxidation (SO)
3. Supplementary Figures
4. Supplementary Tables
5. Supplementary References
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1. Absorption of light energy by the cyanobacterial population in the studied microbial mats
Reflectance of the C/B mats and B/C, denoted as RC/B and RB/C, respectively, was measured as previously described (Al-Najjar et al., 2010), using a fibre-optic field radiance microsensor (Jørgensen & Marais, 1988; Kühl & Jørgensen, 1994). To arrive at expressions for the fraction of the incident light energy absorbed by the cyanobacterial population in the mat, AC, we modeled light propagation in the two mat types as schematically shown in Figure S8.
For the C/B mats we assumed that light attenuation occurred only due to absorption by cyanobacteria (Figure S8A). Thus we calculated AC as 1 – RC/B.
In the B/C mats the light path and its interaction with the microorganisms were more complicated and therefore AC was estimated as follows. First, we assumed that light attenuation in the top layer dominated by large sulphur oxidizing bacteria (SOB) occurred only due to scattering. Second, approximating the B/C mat by a two-layer optical structure, we assumed that propagation of the incident light energy flux, I0, through the mat followed boundary conditions summarized in Figure S8B. Specifically, at the upper interface of the SOB layer, the upwelling light flux was a sum of the component that was back-scattered from the SOB layer, I0RB, and the component that was twice transmitted 2 through the SOB layer and once back-scattered from the cyanobacterial layer, I0TB RC. Consequently, the reflectance of the B/C mat, defined as the ratio between the upwelling and downwelling light fluxes, was approximated as
2 RB / C = RB + RCTB . (1)
The assumption of no light absorption in the SOB layer implied that RB=1–TB, which made it possible to rewrite Eq. 1 as a quadratic equation for TB,
2 0= RCTB TB + (1 RB / C ) , (2) from which TB can be calculated as
1± 1 4RC 1 RB / C TB = . (3) 2RC
In this equation, only the expression with the '–' sign was considered as physically meaningful, because the one with the '+' sign yielded TB>1.
Following Figure S8B, AC in the B/C mats is given by
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AC = TB (1 RC ) , (4) which accounts for the fact that the light absorbed (i.e., not back-scattered) by the cyanobacteria (factor 1–RC) must have been first transmitted through the SOB layer
(factor TB).
Because it was not possible to determine the reflectance of the cyanobacterial layer in the B/C mat without destroying the mat, we assumed that it was equal to the experimentally determined reflectance of the cyanobacteria in the C/B mat, i.e., RC = RC/B.
Combining Eqs. 3 and 4, AC in the B/C mat was therefore estimated as
1 RC / B 1 1 4RC / B 1 RB / C AC = . (5) 2RC / B
Experimentally determined reflectance values were RC/B = 0.045 and RB/C = 0.81, which implied AC = 0.955 for the C/B mat and AC = 0.183 for the B/C mat. Since the cyanobacterial layer in the B/C mats was thinner and less dense than in the C/B mats, RC was most likely lower than RC/B. Furthermore, some light absorption probably occurred also in the SOB layer. Thus, the fraction of incident light energy absorbed by cyanobacteria in the B/C mats was probably lower than the upper limit of 0.183.
2. Conversion of fluxes of O2, Stot and light to fluxes of fixed CO2 and conserved energy
Here we provide the details of how we estimated the fluxes of CO2 fixed and energy conserved by the two dominant processes in the studied mats – photosynthesis and aerobic sulphide oxidation – from the measured fluxes of O2 and Stot. We assumed that the conserved energy comprises two components: (i) energy required for the reduction of the electron carrier (e.g., NADP+) that is used as the electron donor for the reduction of
CO2, and (ii) energy required in the actual CO2 fixation pathway.
2.1. Photosynthesis (P)
Photosynthetic CO2 fixation by cyanobacteria was assumed to follow reactions
H2O + CO2 Æ O2 + CH2O, (6) 0 2 H2S + CO2 Æ 2 S + CH2O + H2O, (7) where the first and second reaction corresponds to oxygenic and anoxygenic P, - respectively. We assumed H2S (not HS ) to be the substrate for, and elemental sulphur
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(S0) to be the product of, anoxygenic P (Cohen et al., 1986; Arieli et al., 1991; Griesbeck et al., 2000; Klatt et al., 2015).
-2 -1 To quantify the net flux of C fixed by oxygenic P (JC,oxy-P, in mol C m s ), we used the measured O2 flux in combination with the CO2/O2 stoichiometry of 1, i.e.,
JC,oxy-P = JO2,oxy-P. (8) -2 -1 Here, JO2,oxy-P (in mol O2 m s ) is the sum of the upward and downward diffusive fluxes from the cyanobacterial layer into the water-column and into the SOB layer, respectively, derived from in situ microsensor profiles. Analogously, to quantify the net flux of C -2 -1 fixed by anoxygenic P (JC,anoxy-P, in mol C m s ), we used the measured Stot flux together with the CO2/Stot stoichiometry of 1/2, i.e.,
JC,anoxy-P = 0.5 JStot,,anoxy-P (9) -2 -1 Here, JStot,,anoxy-P (in mol Stot m s ) is the total sulphide flux consumed in the cyanobacterial layer, as derived from in situ microsensor profiles.
-2 -1 To quantify the flux of energy conserved by oxygenic P (JE,oxy-P, in kJ m s ), we multiplied the CO2 flux with the energy demand of oxygenic P, i.e.,
JE,oxy-P = ǻEr(oxy-P) JC,oxy-P. (10)
Here, the energy demand ǻEr(oxy-P) was calculated by considering that during oxygenic
P two separate energy-demanding reactions are performed. Specifically, CO2 reduction in the Calvin cycle requires conversion of ATP into ADP and Pi. Assuming an ATP energy -1 content of 41 kJ (mol ATP) , the energy requirement of the Calvin cycle is ǻGr(CO2 fix) -1 = 123 kJ (mol CO2) (Supplemental material of Bar-Even et al., 2010). ATP is generated via the light energy-driven membrane-associated electron transport reactions. These reactions additionally have to provide two moles of reducing equivalents NADPH required for the reduction of one mole of CO2. Using the Gibbs free energies of formation given by Thauer et al. (1977) and (Alberty, 1998) energy demand for NADP+ reduction + -1 -1 coupled to H2O oxidation is 533 kJ (2 mol NADP ) = 533 kJ (mol CO2) . Thus, by -1 summing up these two contributions we obtained ǻEr(oxy-P) = 656 kJ (mol CO2) .
Analogously, we quantified the flux of energy conserved by anoxygenic P -2 -1 (JE,anoxy-P, in kJ m s ) by multiplying the energy demand of anoxygenic P with the CO2 -2 -1 flux consumed by anoxygenic P (JC,anoxy-P, in mol CO2 m s ), i.e., -2 -1 JE,anoxy-P (kJ m s ) = ǻEr(anoxy-P) JC,anoxy-P. (11)
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+ Here, ǻEr(anoxy-P) is the sum of the energy demand for NADP reduction by H2S (74 kJ -1 -1 (mol CO2) ) and for CO2 fixation (123 kJ (mol CO2) ), i.e., ǻEr(anoxy-P) = 197 kJ (mol -1 CO2) .
2.2. Aerobic chemolithoautotrophic sulphide oxidation (SO)
Chemolithoautotrophic microorganisms derive their energy for CO2 reduction from a chemical reaction with negative ǻGr. For aerobic SO this reaction is H2S oxidation with
O2. H2S additionally serves as the electron donor for the reduction of CO2. To calculate the fluxes of CO2 fixed and energy gained and conserved by SO in the studied mats, we followed the generic theoretical framework described in Klatt & Polerecky (2015).
According to this framework, the generalized equation for aerobic SO coupled to CO2 reduction is
H2S + y (2 – 1.5x) O2 + (1 – y) (2 – 1.5x) CO2 + (1 – y) (2 – 2.5x) H2O Æ 0 2- + x S + (1 – x) SO4 + (1 – y) (2 – 1.5x) CH2O + y x H2O + (2 – 2x) H , (12)
0 2- where x and (1 – x) describe the fractions of the sulphide pool oxidized to S and SO4 , respectively, and y and (1 – y) describe the fractions of the sulphide pool used for energy generation and CO2 reduction, respectively (both x and y are between 0 and 1).
To calculate the flux of CO2 fixed by aerobic SO, we note that the stoichiometric coefficients for Stot, O2 and CO2 in Eq. 12 are related as S 1 tot = (13) O 2 y 2 1.5x and CO 1 y 2 = , (14) O 2 y Thus, if the parameter x is known, Eq. 13 makes it possible to calculate the parameter y from the measured ratio of the Stot and O2 fluxes consumed by aerobic SO, which can -2 -1 subsequently be used to calculate the flux of CO2 fixed (JC,SO, in mol C m s ) from the -2 -1 flux of O2 consumed (JO2,SO, in mol O2 m s ) by aerobic SO from Eq. 15, namely as 1 y J = J . (15) C,SO y O2,SO
-2 -1 To calculate the flux of energy gained (JE,SO,gained, in kJ m s ) and conserved -2 -1 (JE,SO, in kJ m s ) by aerobic SO, we additionally took into account the efficiency of energy conservation by SO, denoted hereafter as İ. As described in Klatt and Polerecky
71 2. ENERGY CONVERSION IN ILLUMINATED SULPHIDIC SPRINGS
(2015), this efficiency can be calculated once the parameters x and y are known using the function aerob_efficiency available at http://nanosims.geo.uu.nl/SOX. Thus, the flux of energy conserved by aerobic SO was calculated as
JE,SO = (1 – y) ǻGr(SO, CO2 red)JStot,SO, (16) -2 -1 where JStot,SO is the flux of Stot consumed by aerobic SO (in mol Stot m s ) and ǻGr(SO, - CO2 red) is the energy required for the reduction of CO2 in aerobic SO (in kJ (mol H2S) 1). This energy requirement was calculated as the sum of the energy needed to reduce + NAD in the reverse electron transport reaction and of the energy needed to fix CO2 in the Calvin cycle (see Supplementary Figure S1 in Klatt & Polerecky, 2015). Finally, the flux of energy gained by aerobic SO to fuel CO2 fixation was calculated as
JE,SO,gained = –JE,SO /İ, (17) as follows from Eq. 30 in Klatt and Polerecky (2015).
In the B/C mats we observed that the Stot/O2 consumption ratio in the SOB layer was approximately constant during the diurnal cycle (Fig. 4D, Fig. S5). This suggested that aerobic SO in these mats was in a steady state, which made it possible to assume that 2- sulphide was oxidized completely to SO4 (Jørgensen et al., 2010). Using Eqs. 12–13, -1 this implied that x = 0 and y = [2(Stot/O2)] . For the two replicate B/C mats, the averaged values of the Stot/O2 consumption ratio measured in the SOB layer were 0.669 and 0.654 (Table S3), which translates into y = 0.747 and y = 0.765, respectively. Using these values together with the local H2S and O2 concentrations and pH measured in situ with microsensors (averaged over the SOB layer), and additionally assuming that the local sulphate concentrations (2 mM) and temperature (13–15 oC) in the SOB layer were the same as in the water column (see Tables 1 and S1), we obtained the energy conversion efficiency values of 17.1% and 16.4% for the SOB populations in the respective replicate B/C mats (Tables 2 and S2). These values fall between the efficiencies previously calculated for Beggiatoa str. MS-81-6 (12.3%) and Beggiatoa str. MS-81-1c (32.5%) (Hagen & Nelson, 1997; Klatt & Polerecky, 2015). Using these values of x, y and İ, we calcualted the fluxes of CO2 fixed, energy conserved and energy gained by aerobic SO from the measured fluxes of Stot and O2 consumed in the SOB layer of the respective B/C mat using Eqs. 14–17.
In contrast to B/C mats, the Stot/O2 consumption ratio in the SOB layer of C/B mats varied strongly over the diurnal cycle (Fig. 4B and S5B). We assumed that this was 0 2- due to the variable end product of SO (S versus SO4 ), as parameterized by x in Eq. 12.
To estimate the fluxes of CO2 and energy under this scenario, we assumed that the highest
72 II. Manuscripts
measured Stot/O2 consumption ratio occurred when sulphide was oxidized incompletely to 0 S (i.e., when x = 1), whereas the lowest measured Stot/O2 ratio corresponded to complete 2- sulphide oxidation to SO4 (i.e., x = 0) (see Eq. 13). Using the same approach as described above, we calculated the energy conservation efficiency İ for the two extreme cases. Specifically, the highest Stot/O2 consumption ratios measured in the two replicate C/B mats were 2.574 and 2.577, which gave the efficiency values of 16.6% and 16.5%, respectively. In contrast, the lowest Stot/O2 consumption ratios were 0.685 and 0.681, which gave the efficiency values of 17.2% and 17.1% in the respective mats. This striking similarity of the calculated values suggested that the SOB population in the studied mats performed aerobic SO with an efficiency that was independent of the end product (i.e., of 0 2- the S /SO4 ratio). In our subsequent calculations we assumed that this efficiency was equal to the average of the calculated values, i.e., 16.9%.
Therefore, to calculate the fluxes of CO2 fixed, energy gained and energy conserved by aerobic SO in the SOB layer of the C/B mats, we first used the O2 or Stot concentration profiles determined in situ with microsensors to determine in every time- point (i) the average concentrations of H2S and O2 and pH in the overlapping zone of H2S and O2 and (ii) the fluxes of Stot and O2 consumed in this zone. Subsequently, we used Eq.
13 together with the average energy conservation efficiency of 16.9% and the Stot/O2 flux ratio to determine the values of x and y in Eq. 12, which we finally used together with
Eqs. 15–17 to calculate the fluxes of CO2 fixed, energy gained and energy conserved by SO. Detailed results obtained for one of the replicate C/B mats are shown in Table S3. The final results integrated over 24 h for the first and second replicate C/B mat are summarized in Table 2 and Table S2, respectively (column SO).
73 2. ENERGY CONVERSION IN ILLUMINATED SULPHIDIC SPRINGS
3. Supplementary Tables wn in Table1. Table 1. Table 1. T [°C] [°C] T 3,4 M] μ [ x
NO 3,4 M] μ [ asurements during night are sho are night during asurements + 4 NH 2 pH 3,4 2) 2) 7.5 199 (+/-5.2) 3.6 (+/- 0.07) 13 1) 1) 8.0 79 (+/- 2.0) 33.4 (+/- 1.8) 14 died mats measured during day. DIC [mM] the day. The corresponding me corresponding The day. the S concentrationS and pH. 2 The corresponding values determined for the night are shown in are shown night for the determined values corresponding The 2 M] μ [ 2 O 2 M] μ ) [ 1 tot S (S 2 andard deviations (n = 3 – 6). deviations andard , was calculated from the measured H the from calculated , was tot C/B-1 Main Spring C/B-2 Main Spring 73 (582) C/B-3 Fissure Spring 13 (103) B/C-1 0 401 (628) Main Spring B/C-2 864 4.1 (+/-0.3 Main Spring 89 (278) 5 B/C-3 3.2 (+/- 0.8) 8.0 Fissure Spring 50 (92) 52 (+/- 3.9) 45 16 (248) - 50 (+/- 8.9) 3.9 (+/-0.2 14.5 65 6.9 80 4.6 (+/-0.09) 7.3 - - - 8.3 - - - 14.5 13 - 15.5 Mat type Site H Total concentration, S sulfide Values are derived from microsensor measurements performed during performed measurements microsensor from derived are Values day. during the taken water samples from determined Values were to st refer parentheses in Values Table S1: Chemical parameters in the water column above stu 1 2 3 4
74 II. Manuscripts xidation (SO) in the (0.12%; 0.13%) (0.12%; 0.24% ) ) -12.02 69.4%) -17.30 (30.6%; 14.94 0.8%) 1929.38 (99.2%;
18.9%) 2.46 4.72 (47.9%; 52.1%) (47.9%; 4.72 18.9%) 2.46 -9.29 -25.54 -34.84 -9.29 -25.54 le for this process. le
(90.5%; 9.5%) (90.5%; 87.7% -5.30 (12.3%; (2.62%; 0.09%) (2.62%; 0.12% (0.10%; 0.02%) 16.4% 0 (96.8%; 3.2%) 0 (96.8%; 2.26 (81.1%; also the energy availab tosynthesis (P, anoxygenic and oxygenic) and aerobic sulphide o 2.71% 0.5%) 1093.06 (99.5%; 1914.44 3 oduced and consumed, respectively. oduced and consumed, 36.05 -12.90 23.15 36.05 -12.90 -38.41 0.65 -39.06 -25.44 -14.38 -39.82 -25.44 -14.38 1087.5 5.56 within the mat through oxygenic P. oxygenic through mat the within idation (SO) was assumed to represent was assumed (SO) idation used when the compound is pr is when the compound used 28.66 (17.5%; 82.5%) 82.5%) 28.66 (17.5%; 0.94 29.6 -48.77 (26.1%; 73.9%) -48.77 (26.1%; -5.09 -53.86 divided by energy available. available. by energy divided 2.64% (0.46%; 2.17%) 16.9% 4
2
)
) -1 ) -1 -1 d Shown are results for the second replicate of each mat type. d P (anoxyP;oxyP) SO C/B-2 mat Total (P; SO) P (anoxyP;oxyP) SO Total (P; SO) B/C-2 mat -2 d -2 -2 m
2
m ) ) m 1
2
-1 -1 tot 1 1 d d -2 -2 flux 2 flux flux 2 tot mmol O mmol S mmol CO Represents energy generated internally Represents energy Conservation efficiency Energy gainedby aerobic sulphide ox
Energy flux available ( S ( Energy flux conserved ( O (kJ m (kJ m (kJ CO Positive and negative values are Positive and energy conserved Calculated as 3 studied mats. Table S2: Daily budgets of reactants and energy utilized by pho 1 4 2
75 2. ENERGY CONVERSION IN ILLUMINATED SULPHIDIC SPRINGS
4
E,SO,gained J 3 red) 2 (SO, O (SO, r on (SO) in the studied mats. G ǻ
4 E,SO 07.02 -780.11 630.72 1 J 3 erved erved gained Energy red) 2 104.15 29.74 -179.70 104.15 29.74 176.27 175.41 72.36 -347.14 175.41 72.36 429.59 -411.22 203.60 69.81 414.14 -253.88 135.67 40.21 239.57 389.14 3.71 -770.55 21.68 409.48 2.46 -764.72 14.94 281.70 78.67 -590.00 281.70 78.67 466.53 -697.75 327.81 97.29 574.39 363.38 -649.86 307.74 84.17 498.10 132.85 51.65 -248.94 132.85 51.65 306.49 (SO, CO (SO, r E ǻ
+ + + +
+ + + +
Æ Æ Æ Æ Æ Æ O + 0.23 H O + 0.55 H O + 0.76 H O + 0.24 H O O O O O O 2 2 2 2 2 2 2 2 2 2
+ + O Æ Æ Æ Æ Æ
2 2 2 2 2 2 O + 1.36 H O + 1.71 H O + 1.99 H O + 1.56 H 2 2 2 2 + 0.51 H + 0.47 H + 0.08 H + 0.33 H + 0.53 H + 0.22 H O + 2 H O + 2 H 2 2 2 2 2 2 2 2 O + 0.73 H O + 0.50 H O + 0.35 H O + 0.72 H 2 2 2 2 O + 0.89 H + 0.16 CO + 0.23 CO + 0.27 CO + 0.16 CO + 0.11 CO 2 2 2 2 2 2 + 0.40 CH + 0.47 CH + 0.53 CH + 0.44 CH 2- 2- 2- 2- 4 4 4 4 + 0.51 CO + 0.47 CO + 0.40 CO + 0.47 CO + 0.53 CO + 0.44 CO + 0.51 CH + 0.47 CH 2 2 2 2 2 2 + 0.16 CH + 0.23 CH + 0.27 CH + 0.16 CH 2- 2- 4 4 2- 2- 2- 2- 4 4 4 4 + 0.11 CH S + 0.51 O S + 0.69 O S + 0.80 O S + 0.52 O S + 0.39 O 0 2 2 2 2 2 S 1 SO 1 SO H H H H H + 0.68 SO + 0.86 SO + 0.99 SO + 0.78 SO 0 0 0 0 S + 1.49 O S + 1.53 O S + 1.12 O S + 1.31 O S + 1.46 O S + 1.23 O 2 2 2 2 2 2 H H H H H H + 0.11 SO + 0.28 SO + 0.38 SO + 0.12 SO 0 0 0 0 0.32 S 0.32 S 0.14 S 0.01 S 0.22 0.89 S 0.89 0.72 S 0.72 S 0.62 S 0.88 energy gained and energy conserved by aerobic sulphide oxidati
1 C,SO
x y J x y for the C/B mat. for the -1 s -2 for the C/B mat. for the pH pH -1 s 2 -2 mol m S 2 μ
H M. 2 μ 2 O in
2 /O tot S 1 -1 -1 for the B/C mats and mats B/C for the -1 O2,SO S) J 2 for the B/C mats and mJ m and mJ for the B/C mats d -1 Measured with microsensors Measured with microsensors Calculated Stoichiometry Energy cons -2
1 d -2 1.08 1.35 1.25 135 20 8.0 0.618 0.746 0.294 0.746 0.618 8.0 1.25 135 20 1.35 1.08 0.476 0.739 0.321 7.9 0.89 150 24 1.07 1.20 2.57 10 8.1 0.055 32 1.27 0.49 1 0.776 0.83 1.62 1.96 50 30 8.1 0.887 0.760 0.133 0.760 1.96 0.887 50 8.1 30 1.62 0.83 1.47 1.12 0.76 180 24 7.9 0.143 0.735 0.696 0.735 0.143 7.9 0.76 180 24 1.12 1.47 0.856 0.733 0.006 7.9 0.69 183 23 1.10 1.61 0.104 0.761 1.93 0.879 50 8.0 32 1.24 0.64 1.13 1.65 1.46 80 30 8.0 0.724 0.750 0.258 0.750 1.46 0.724 80 8.0 30 1.65 1.13 1.28 1.04 0.81 120 25 7.9 0.220 0.737 0.562 0.737 0.220 7.9 0.81 120 25 1.04 1.28 Stot,SO J in mmol in mmol m Averagedover the SO layer, H (mol in kJ kJ m in Shown are results for two B/C mats and one C/B mat. B/C mats Shown are results for two 2 Table S3: Theoretical stoichiometry together with the values of 1 4 3
Mat B/C-2 25.54 39.06 0.654 40 30 6.5 0 0.765 12.00 C/B-1 76 B/C-1 37.67 56.3 0.669 95 73 7.1 0 0.747 19.06 II. Manuscripts
4. Supplementary Figures
Figure S1: Hyperspectral imaging of two dominant cyanobacterial morphotypes found in the studied microbial mats. Shown are transmission spectra with clear spectroscopic signatures of cyanobacteria-specific pigments chlorophyll a (maximal in vivo absorption at wavelengths 680 nm and 450 nm) and phycocyanin (at 625 nm). Images of the filaments were constructed by assigning the curvature of the transmission spectrum at 680 nm to the intensity of the green channel (Polerecky et al., 2009).
1.0
0.8
0.6
Reflectance 0.4
0.2
0.0 400 500 600 700 800 900
O (nm) Figure S2: Example reflectance spectrum of a C/B mat. The spectrum shows spectroscopic signatures corresponding to cyanobacteria-specific pigments chlorophyll a and phycocyanin (maximal in vivo absorption at wavelengths 680 nm and 625 nm, respectively). The lack of spectral signatures of bacteriochlorophylls (absorption maxima between 700 and 890 nm; Polerecky et al. 2009) indicates that the abundance of obligatory anoxygenic phototrophs such as green or purple sulphur/non-sulphur bacteria in the studied mats was negligible.
77 2. ENERGY CONVERSION IN ILLUMINATED SULPHIDIC SPRINGS
Figure S3: In situ measurements in the C//B mats over a 24 h period. Shown are downwelling irradiance of the photosynthetically available radiation (PAR) and vertical profiles of H2S and O2 concentrations as a function of time in two replicate mats. All measurements were done simultaneously, with microsensor tips separated by not more than 5 mm. Zero depth corresponds to the mat surface.
78 II. Manuscripts
VIS VIS VIS NIR NIR NIR VIS I=289 I=289 I=215 I=13 I=41 I=194 I=289 I=215 400 70
60 350 50 300 40
(μM)
(μM)
2
tot
S O 30 250 20 200 10
0 150 012345678910 time (min)
Figure S4: Light-induced dynamics of O2, H2S, Stot and pH inside the cyanobacterial layer of a C/B mat. Total sulphide concentrations, Stot, were calculated from the measured H2S concentrations and pH. The irradiance was either in the visible (VIS; 400-700 nm) or near infrared (NIR; a mixture of light emitted by light-emitting diodes with emission maxima at 740 and 810 nm) part of the spectrum. Incident irradiance (I, in μmol photons m-2 s-1) applied during the indicated time intervals is also shown (intervals shaded with gray correspond to darkness). Distance between sensor tips was < 5 mm.
79 2. ENERGY CONVERSION IN ILLUMINATED SULPHIDIC SPRINGS
Figure S5: Activity of cyanobacterial and SOB populations in the C/B-2 mat over a 24 h period. The fluxes were derived from in situ microsensor profiles of O2 and H2S concentration (Fig. S3) and pH. The corresponding average H2S concentration in the cyanobacterial layer, the Stot/O2 flux ratio in the SOB layer, and the depth interval of the zone in the maatt where sulphide was oxidized with oxygen are also shown. Grey bars in panel C correspond to the time-points when the Stot/O2 consumption ratio was <0.7. In all panels shaded and non-shaded areas correspond to dark and light, respectively.
80 II. Manuscripts
3.0 C/B−2 C/B−1 B C/B−1 2.5 A C/B−2 P = 0.1998 P = 0.1447 2.5 P <0.0001 P <0.0001 2 2 2 R = 0.2226 R = 0.2779 R = 0.9097 R 2= 0.9178 2.0 2.0 2 2 O O
/ 1.5 /
ot ot t t 1.5 S S 1.0 1.0 0.5
0.5 0.0 0.00.51.01.52.0 0.00.51.01.52.0 µm ol S m −2 s−1 µm ol O m −2 s−1 tot 2
2.5 C 2.5 D C/B−1 C/B−2 P <0.0001 P <0.0001 C/B−2 R 2= 0.9089 R 2= 0.9162 2.0 P <0.0001 2.0 R 2= 0.9162 2 2 1.5 O O /
/ 1.5
ot ot t t S S 1.0 1.0 C/B−1 P = 0.0083 0.5 0.5 R 2= 0.665 0.0 20 25 30 35 0 50 100 150 200 [H S] (µM ) [O ] (µM ) 2 2
Figure S6: Regulation of aerobic SO in C/B mats. Shown are ratios between the fluxes of the total sulphide (Stot) and O2 consumed in the SOB layer (Stot/O2) as a function of the flux of Stot consumed in the SOB layer (panel A), flux of O2 consumed in the SOB layer (panel B), average concentration of H2S in the SOB layer (panel C) and average concentration of O2 in the SOB layer (panel D). All values were calculated from the concentration depth profiles measured in situ. Lines and legends show the results of the respective correlation analyses.
81 2. ENERGY CONVERSION IN ILLUMINATED SULPHIDIC SPRINGS
Figure S7: In situ measurements in the B/C mats over a 24 h period. Shown are downwelling irradiance of the photosynthetically available radiation (PAR), and vertical profilees of H2S and O2 concentrations as a function of time in two replicate mats. All measurements were done simultaneously, with microsensor tips separated by not more than 5 mm. Intervals when microsensor profiles were not recorded are shown in black. Zero depth corresponds to the mat surface.
82 II. Manuscripts
I water water A 0 B I0 2 I0 R C I0 R B I 0 R C T B
Cyanobacteria BeggiatoaSOB absorbed: I0 (1 -R C) I0 TB RC Beggiatoa SOB (1 Cyanobacteria absorbed: I0 TB (1 -R C)
Figure S8: Schematic diagrams of the models of light propagation through a C/B and B/C mat. Shown are fluxes of incident, transmitted, absorbed and back-scattered (“reflected”) light. Transmission and reflectance of the SOB layer is denoted by TB and RB, respectively, reflectance of the cyanobacterial layer as RC, light energy flux as I0.
5. Supplementary References
Alberty RA. (1998). Calculation of standard transformed formation properties of biochemical reactants and standard apparent reduction potentials of half reactions. Arch Biochem Biophys 358:25–39.
Al-Najjar MAA, de Beer D, Jørgensen BB, Kühl M, Polerecky L. (2010). Conversion and conservation of light energy in a photosynthetic microbial mat ecosystem. ISME J 4:440– 449.
Arieli B, Padan E, Shahak Y. (1991). Sulfide-induced sulfide-quinone reductase activity in thylakoids of Oscillatoria limnetica. J Biol Chem 266:104–11.
Bar-Even A, Noor E, Lewis NE, Milo R. (2010). Design and analysis of synthetic carbon fixation pathways. Proc Natl Acad Sci U S A 107:8889–94.
Cohen Y, Jørgensen BB, Revsbech NP, Poplawski R. (1986). Adaptation to hydrogen sulfide of oxygenic and anoxygenic photosynthesis among cyanobacteria. Appl Environ Microbiol 51:398–407.
Griesbeck C, Hauska G, Schütz M. (2000). Biological sulfide oxidation: Sulfide-quinone reductase (SQR), the primary reaction. Recent Res Dev Microbiol 4:179–203.
Hagen KD, Nelson DC. (1997). Use of reduced sulfur compounds by Beggiatoa spp.: Enzymology and physiology of marine and freshwater strains in homogeneous and gradient cultures. Appl Environ Microbiol 63:3957–3964.
83 2. ENERGY CONVERSION IN ILLUMINATED SULPHIDIC SPRINGS
Jørgensen BB, Dunker R, Grünke S, Røy H. (2010). Filamentous sulfur bacteria, Beggiatoa spp., in arctic marine sediments (Svalbard, 79 degrees N). FEMS Microbiol Ecol 73:500–13.
Jørgensen BB, Marais DJ Des. (1988). Optical properties of benthic photosynthetic communities: fiber-optic studies of cyanobacterial mats. Limnol Oceanogr 33:99–113.
Klatt JM, Al-Najjar MAA, Yilmaz P, Lavik G, de Beer D, Polerecky L. (2015). Anoxygenic photosynthesis controls oxygenic photosynthesis in a cyanobacterium from a sulfidic spring. Appl Environ Microbiol AEM.03579–14.
Klatt JM, Polerecky L. (2015). Assessment of the stoichiometry and efficiency of CO2 fixation coupled to reduced sulfur oxidation. Front Microbiol 6. doi:10.3389/fmicb.2015.00484.
Kühl M, Jørgensen BB. (1994). The light field of microbenthic communities: radiance distribution and microscale optics of sandy coastal sediments. Limnol Oceanogr 39:1368– 1398.
Polerecky L, Bissett A, Al-Najjar MAA, Färber P, Osmers H, Suci PA, et al. (2009). Modular spectral imaging system for discrimination of pigments in cells and microbial communities. Appl Environ Microbiol 75:758–771.
Thauer RK, Jungermann K, Decker K. (1977). Energy conservation in chemotrophic anaerobic bacteria. Bacteriol Rev 41:100–180.
84 II. Manuscripts
ASSESSMENT OF THE STOICHIOMETRY AND EFFICIENCY OF
CO2 FIXATION COUPLED TO REDUCED SULFUR OXIDATION
Klatt, Judith M.1*, Polerecky, Lubos1,2*
1Max Planck Institute for Marine Microbiology, Celsiusstr.1, 28359 Bremen, Germany
2Department of Earth Sciences – Geochemistry, Faculty of Geosciences, Utrecht University, Princetonplein 9, 3584 CC, Utrecht, The Netherlands
Published in
Frontiers in Microbiology (2015)
*Corresponding authors
85 3. STOICHIOMETRY AND EFFICIENCY OF SULFUR OXIDATION
Abstract Chemolithoautotrophic sulfur oxidizing bacteria (SOB) couple the oxidation of reduced sulfur compounds to the production of biomass. Their role in the cycling of carbon, sulfur, oxygen and nitrogen is, however, difficult to quantify due to the complexity of sulfur oxidation pathways. We describe a generic theoretical framework for linking the stoichiometry and energy conservation efficiency of autotrophic sulfur oxidation while accounting for the partitioning of the reduced sulfur pool between the energy generating and energy conserving steps as well as between the main possible products (sulfate versus elemental sulfur). Using this framework, we show that the energy conservation efficiency varies widely among SOB with no apparent relationship to their phylogeny. Aerobic SOB equipped with reverse dissimilatory sulfite reductase tend to have higher efficiency than those relying on the complete Sox pathway, whereas for anaerobic SOB the presence of membrane-bound, as opposed to periplasmic, nitrate reductase systems appears to be linked to higher efficiency. We employ the framework to also show how limited rate measurements can be used to estimate the primary productivity of SOB without the knowledge of the sulfate-to-elemental-sulfur production ratio. Finally, we discuss how the framework can help researchers gain new insights into the activity of SOB and their niches.
1. Introduction
Autotrophic sulfur oxidizing bacteria (SOB) comprise a phylogenetically diverse group of microbes that obtain the energy required for growth from the oxidation of reduced sulfur compounds. Prominent natural habitats of SOB are hydrothermal vents, where these bacteria live in symbiotic association with invertebrates or, when free-living, form microbial mats (e.g. Sievert and Vetriani, 2012). SOB are found also on top of organic-rich marine sediments, where they typically form conspicuous and often quite extensive mats. For instance, on the continental shelf off the Namibian coast, such mats cover an area comparable to the size of Austria (>80,000 km2; Brüchert et al., 2006). In addition to natural habitats, SOB are important also in industrial applications, where they are used, for instance, for waste water treatment by biodesulfurisation (Janssen et al., 2009).
A common feature of environments inhabited by SOB is the encounter of sulfide or other reduced sulfur species with a terminal electron acceptor (TEA) such as oxygen or
86 II. Manuscripts nitrate. This encounter creates a chemical disequilibrium from which energy can be harvested and conserved in the form of biomass. While the general validity of this concept is well established, there are still significant gaps in our understanding of systems driven by chemical energy derived from reduced sulfur oxidation. This is because the reactions performed by autotrophic SOB are highly complex and the assessment of the stoichiometry of sulfur oxidation is not trivial, even for cultivated SOB.
In chemolithoautotrophic SOB known to date the energy gained from the reduced sulfur oxidation drives reverse electron transport within the membrane, which generates sufficiently negative electron potential to reduce an electron carrier (e.g. NAD+) with the reduced sulfur compound as the electron donor (e.g. Harold 1986). Thus, the reduced sulfur compound is used as the electron donor for the reduction of both CO2 (via the electron carrier) and TEA. The partitioning of the reduced sulfur pool between these two reductive processes, which is related to the efficiency of energy conservation and the growth rate per substrate utilized, is poorly understood and varies substantially among the different cultivated SOB (Kelly, 1999). Additionally, the end product of sulfur oxidation (e.g., sulfate versus elemental sulfur) is also variable and appears to exhibit a degree of flexibility even within individual organisms, as shown for several Beggiatoa strains (Nelson et al., 1986; Hagen and Nelson, 1997; Berg et al., 2014).
This complexity is echoed in the diversity of pathways by which SOB oxidize the reduced sulfur compound. As summarized in Figure 1, three main pathways have been identified so far (reviewed, e.g., by Ghosh and Dam, 2009): (i) the Sox pathway mediated by the thiosulfate-oxidizing multi-enzyme (TOMES) complex, (ii) the tetrathionate (SI4) pathway of thiosulfate oxidation, and (iii) the rather recently described “branched” pathway. The occurrence of individual enzymes of the traditional pathways does not appear to be linked to the phylogenetic identity of SOB. This phenomenon is generally explained by horizontal gene transfer, which is possibly also responsible for the co-occurrence and linkage of specific enzymes or even several complete pathways in the same organism (e.g. Ghosh and Dam, 2009). In addition to the diverse sulfur oxidation pathways, the complexity of autotrophic sulfur oxidation by SOB is further increased by the fact that they can employ different terminal oxidases for TEA reduction and two possible pathways for CO2 fixation (namely the Calvin-Benson-Bassham cycle, hereafter referred to as the Calvin cycle, or the
87 3. STOICHIOMETRY AND EFFICIENCY OF SULFUR OXIDATION reverse tricarboxylic acid (rTCA) cycle; e.g., Sievert and Vetriani, 2012).
2- 0 S2O3 S H2S
Qred Qox soxXA cyt cox cyt cred cyt cred cyt cox soxYZ FCC
soxXA soxXA SQR PQ soxYZ ox PQred soxYZ cyt cred cyt cox 2- S4O6 “S0“ S0
cyt cox cyt cox soxB soxCD GSSH soxB
cyt cred cyt cred H2S
cyt bred cyt cred 2- SO cyt bred 4 O cyt cred 2 cyt b ox SDO cyt cox SOAR
ATP cyt box cyt cox Q APS ox rDSR
Q SO 2- red 3
Figure 1: Scheme of the main pathways of reduced sulfur oxidation. Green arrows indicate the traditional Sox pathway, blue arrows the tetrathionate (SI4) pathway and red arrows correspond to the branched pathway for thiosulfate oxidation. Possible entry sites of H2S that are not part of these traditional pathways are shown with a grey dotted arrow. FCC = flavocytochrome c:oxidoreductase; SQR = sulfide:quinine:oxidoreductase; GSSH = S-sulfanylglutathione; APS = adenosine-5’- phosphosulfate; SOAR = sulfite:cyt c:oxidoreductase; rDSR = reverse dissimilatory sulfate reductase, SDO = sulfur dioxygenase; soxXA, soxYZ and soxB are subunits of the thiosulfate-oxidizing multi- enzyme (TOMES) complex.
The complexity of SOB physiology and sulfur oxidation biochemistry hampers our ability to quantify the contribution of SOB to sulfur, carbon, nitrogen and oxygen cycling in a given environment, as well as to understand their niches. This is especially true for environments with fluctuating conditions, where also the products and intermediates of sulfur oxidation vary. Additionally, this complexity hinders the development of industrial applications that use SOB for processes such as waste water biodesulfurisation, which require precise control and predictability of SOB activity.
In this study we describe a generic theoretical framework for a rapid quantitative assessment of chemolithoautotrophic sulfur oxidation. First, we demonstrate how to use it for the quantification of the stoichiometry and energy conservation efficiency of autotrophic
88 II. Manuscripts sulfur oxidation from rate measurements of the reactants involved. Second, we apply it on literature data and identify possible links between the energy conservation efficiency and the different oxidative and reductive pathways employed by SOB. Third, we suggest how it can be used to estimate chemolithoautotrophic primary productivity by SOB from limited rate measurements. Finally, we discuss how our framework makes it possible to formally relate the capabilities of SOB (e.g., stoichiometry and efficiency) to their environment and thus gain insight into the differentiation of their niches.
2. Materials and methods
2.1. Generalized equations for aerobic sulfide oxidation
The generalized mass-balanced equations for aerobic sulfide oxidation coupled to CO2 fixation are derived by considering the energy generating and energy conserving steps separately. The energy generating step is performed with elemental sulfur (S0) and sulfate 2- (SO4 ) as two possible end products. The corresponding reactions are written as (Nelson et al., 1986; Kelly, 1999)
0 H2S + 0.5 O2 Æ S + H2O, (1)
2- + H2S + 2 O2 Æ SO4 + 2 H . (2)
When both reactions occur simultaneously, the generalized equation for the energy generating step is
0 2- + H2S + (2 – 1.5x) O2 Æ x S + (1 – x) SO4 + x H2O + (2 – 2x) H , (3)
0 2- where x and (1–x) are the parts of the total H2S pool oxidized to S and SO4 , respectively (0x1).
In SOB, H2S serves not only as the energy source but also as the electron donor for 0 the reduction of CO2. Analogously to the energy generating step, this can occur with S or 2- SO4 as two possible end products, i.e.,
0 H2S + 0.5 CO2 Æ S + 0.5 CH2O + 0.5 H2O, (4)
2- + H2S + 2 CO2 + 2 H2O Æ SO4 + 2 CH2O + 2 H . (5)
0 2- A critical assumption in our derivation is that the S :SO4 product ratio (defined by x) is the
89 3. STOICHIOMETRY AND EFFICIENCY OF SULFUR OXIDATION
same for both the energy generating and CO2 fixing steps. Consequently, the generalized equation for the CO2 fixing step is written as
H2S + (2 – 1.5x) CO2 + (2 – 2.5x) H2O Æ 0 2- + x S + (1 – x) SO4 + (2 – 1.5x) CH2O + (2 – 2x) H . (6)
To arrive at a generalized equation for the complete aerobic sulfide oxidation coupled to CO2 reduction, we assume that part y (0y1) of the total H2S pool is used for energy generation (Eq. 3) while the remaining part 1–y is used for CO2 reduction (Eq. 6). Thus, by summing Eq. 3 multiplied with y and Eq. 6 multiplied with 1–y we obtain
0 2- + H2S + ȞO2 O2 + ȞCO2 CO2 Æ ȞS0 S + ȞSO4 SO4 + ȞorgC CH2O + ȞH2O H2O + ȞH+ H , (7) where the stoichiometric coefficients of the reactants involved are ȞO2 = y(2–1.5x), ȞCO2 =
ȞorgC = (1–y)(2–1.5x), ȞS0 = x, ȞSO4 = (1–x), ȞH2O = yx–(1–y)(2–2.5x) and ȞH+ = 2(1–x). Equation 7 is the general mass-balanced equation for aerobic sulfide oxidation coupled to
CO2 fixation.
2.2. Efficiency of energy conservation: the traditional calculation approach
The thermodynamic efficiency of energy conservation, İ, is generally defined as the ratio between the Gibbs free energies of the endergonic (energy-consuming) and exergonic
(energy-generating) reactions. Specifically for aerobic sulfide oxidation, where CO2 reduction (Eq. 6 multiplied by 1 – y) and O2 reduction (Eq. 3 multiplied by y) are the endergonic and exergonic reactions, respectively, İ is written as
(1 y) 'Gr (CO 2 red) İ I , (8) y 'Gr (O 2 red) where ǻGr is expressed per mole of H2S oxidized. At non-standard conditions, the Gibbs free energy of a reaction is calculated as
0 ǻGr = ǻGr + RT ln Q, (9)
0 where ǻGr is ǻGr at standard biochemical conditions (pH=7, reactant concentrations 1 M, temperature 25 oC), R is the universal gas constant, T is temperature, and Q is the ratio of the activity coefficients of the products and educts involved, which can be approximated by the
90 II. Manuscripts ratio of the corresponding concentrations (Thauer et al., 1977). Using Eqs. 3 and 6, the respective ǻGr values in Eq. 8 are calculated as
0 0 0 2- 0 0 ǻGr(O2 red) = x ǻGf (S ) + (1 – x) ǻGf (SO4 ) + x ǻGf (H2O) – ǻGf (H2S) 0 – (2 – 1.5x) ǻGf (O2) + RT ln Q1, (10)
0 0 0 2- 0 0 ǻGr(CO2 red) = x ǻGf (S ) + (1 – x) ǻGf (SO4 ) – ǻGf (H2S) – (2 – 2.5x) ǻGf (H2O) 0 0 + (2 – 1.5x) [ǻGf (CH2O) – ǻGf (CO2)] + RT ln Q2, (11) where the Gibbs free energies of formation of the respective reactants at standard 0 biochemical conditions, ǻGf , are tabulated in (Thauer et al., 1977) and the quotients Q1 and
Q2 are given by
2 (1x) (22x) [SO 4 ] [H ] Q1 (21.5x) , (12) [H 2S] [O 2 ]
2 (1x) (22x) [SO 4 ] [H ] Q2 (21.5x) . (13) [H 2S] [CO 2 ]
Equations 8-13 summarize the traditional approach for the calculations of the energy conservation efficiency of autotrophic aerobic sulfide oxidation (e.g., Nelson and Hagen, 1995).
2.3. Factorization of the traditional energy conservation efficiency
The general aim of calculating the efficiency of energy conservation is to gain insights into how the energy generated by an exergonic reaction is converted into the biochemical currency ATP and how this ATP is further utilized to drive endergonic reactions. For the specific case of autotrophic sulfur oxidation, we divide the flow of energy into three steps: conversion of the Gibbs free energy released by sulfur oxidation into ATP, transfer of this
ATP to the site of CO2 reduction (that is, not for processes associated with “cell maintenance”), and ATP utilization for driving CO2 reduction (see Supplementary Figure S1).
The efficiency of the first step, İSO, characterizes the efficiency of the sulfur oxidation pathway. It is calculated as
'Er (O 2 red) İSO , (14) 'Gr (O 2 red)
91 3. STOICHIOMETRY AND EFFICIENCY OF SULFUR OXIDATION
where ¨Er(O2 red) describes the energy gained, in the form of ATP, from the reduction of O2.
Analogously, the efficiency of ATP utilization for the reduction of CO2, İu, is calculated as
'Gr (CO 2 red) İ u , (15) 'Er (CO 2 red) where ¨Er(CO2 red) describes the energy requirements, in the form of ATP, of CO2 reduction.
Finally, the efficiency of the energy transfer, İt, is calculated as
(1 y) 'Er (CO 2 red) İ t , (16) y 'Er (O 2 red) where, as above, y and 1 – y describe the fractions of the total H2S pool used for energy generation and CO2 reduction, respectively. Note that the values of ¨Er and ¨Gr in Eqs. 14–
16 are expressed per mole of H2S.
As follows from Eqs. 8 and 14–16, the overall thermodynamic efficiency İI can be expressed as a product of the partial efficiencies of the three steps involved in the energy flow associated with autotrophic sulfur oxidation, namely
İ I İSO İ u İ t . (17)
Because İI can be calculated from thermodynamic data based on the stoichiometry of autotrophic sulfur oxidation, this means that Eq. 17 can be used to calculate any one of the efficiencies İSO, İu or İt provided that the other two are known. Although this is not possible at the current state of knowledge, Eq. 17 makes it possible to estimate their minimal values.
Specifically, the minimal value of İSO is reached when both İu and İt are maximal (i.e., İu = İt
= 1), which gives İSO,I,min = İI. Analogously, İu,min = İt,min = İI. Thus, İI represents the minimal value of the partial efficiencies İSO, İu and İt.
2.4. Efficiency of energy conservation: the new calculation approach
The traditional approach of efficiency calculation does not consider that sulfide oxidation and + CO2 reduction in SOB are coupled via redox couples such as NAD /NADH, FAD/FADH2 and oxidized/reduced Ferredoxin (Fdox/Fdred). In our new approach we account for this coupling explicitly. Also, we distinguish between two possible CO2 fixation pathways observed in SOB: the Calvin cycle and the reverse tricarboxylic acid (rTCA) cycle. In
92 II. Manuscripts accordance with Bar-Even et al. (2010), we assume that glyceraldehyde-3-phosphate (GA3P) is the primary product of both CO2 fixation pathways.
In the SOB employing the Calvin cycle, CO2 fixation occurs according to reaction
+ 6 NADH + 3 CO2 + Pi Æ 6 NAD + GA3P + 7 H2O. (18)
This reaction requires conversion of ATP into ADP and Pi, and its energy requirement is -1 ǻGr(CO2 fix) = 69.7 kJ CO2 (Supplemental material of Bar-Even et al., 2010). The reducing equivalents NADH required for the reduction of CO2 in Eq. 18 are produced via membrane- associated reverse electron transport (RET) reactions with zero-valent sulfur (S0) or sulfate 2- (SO4 ) as two possible end products:
+ 0 + H2S + NAD Æ S + NADH + H , (19)
+ 2- + H2S + 4 NAD + 4 H2O Æ SO4 + 4 NADH + 6 H . (20)
These two endergonic reactions are driven by proton motive force, which is generated either directly by the exergonic reaction (Eq. 3) or by utilizing ATP. When both of them occur simultaneously, the generalized equation for the RET reaction is
+ H2S + (4 – 3x) NAD + (4 – 4x) H2O Æ 0 2- + x S + (1 – x) SO4 + (4 – 3x) NADH + (6 – 5x) H , (21)
0 2- where x again describes the S :SO4 product ratio, and its energy requirement is calculated as
0 0 0 2- 0 0 ǻGr(RET) = x ǻGf (S ) + (1 – x) ǻGf (SO4 ) – ǻGf (H2S) – 4(1 – x) ǻGf (H2O) 0 0 + + (4 – 3x) [ǻGf (NADH) – ǻGf (NAD )] + RT ln Q3. (22)
0 0 + -1 Here the difference ǻGf (NADH) – ǻGf (NAD ) = 60.99 kJ (mol NADH) (Alberty, 1998) and the reaction quotient Q3 is given by
2 (1x) (43x) (65x) [SO 4 ] [NADH] [H ] Q3 (43x) . (23) [H 2S] [NAD ]
In the SOB employing the rTCA cycle, CO2 fixation occurs according to reaction
+ 3 NADH + 4 Fdred + FADH2 + 3 CO2 Æ 3 NAD + 4 Fdox + FAD + GA3P + 7 H2O. (24)
-1 Energy requirement of this reaction is ǻGr(CO2 fix) = 64.3 kJ CO2 (Bar-Even et al., 2010).
The reducing equivalents NADH, FADH2 and Fdred required for the reduction of CO2 in Eq.
24 are produced by the reduction of the corresponding redox couples with H2S according to
93 3. STOICHIOMETRY AND EFFICIENCY OF SULFUR OXIDATION two possible reactions:
+ H2S + 0.5 NAD + 0.67 Fdox + 0.17 FAD Æ 0 + S + 0.5 NADH + 0.67 Fdred + 0.17 FADH2 + 1.17 H , (25)
+ H2S + 2 NAD + 2.67 Fdox + 0.67 FAD + 4 H2O Æ 2- + SO4 + 2 NADH + 2.67 Fdred + 0.67 FADH2 + 6.67 H . (26)
When both of these reactions occur simultaneously, the generalized equation for the RET reaction is
+ H2S + (2–1.5x) NAD + (2.67–2x) Fdox + (0.67–0.5x) FAD + (4–4x) H2O Æ 0 2- + x S + (1–x) SO4 + (2–1.5x) NADH + (2.67–2x) Fdred + (0.67–0.5x) FADH2 + (6.67–5.5x) H (27) and its energy requirement is calculated as
0 0 0 2- 0 0 ǻGr(RET) = x ǻGf (S ) + (1 – x) ǻGf (SO4 ) – ǻGf (H2S) – 4(1 – x) ǻGf (H2O) 0 0 + + (2 – 1.5x) [ǻGf (NADH) – ǻGf (NAD )] 0 0 + (2.67 – 2x) [ǻGf (Fdred) – ǻGf (Fdox)] 0 0 + (0.67 – 0.5x) [ǻGf (FADH2) – ǻGf (FAD)] + RT ln Q4. (28)
0 0 + -1 0 0 Here, ǻGf (NADH) – ǻGf (NAD ) = 60.99 kJ (mol NADH) , ǻGf (FADH2) – ǻGf (FAD) = -1 0 0 -1 42.65 kJ (mol FADH2) , ǻGf (Fdred) – ǻGf (Fdox) = 38.88 kJ (mol Fdox) (Alberty, 1998) and the reaction quotient Q4 is given by
2 (1x) (21.5x) (2.672x) (0.670.5x) (6.675.5x) [SO 4 ] [NADH] [Fd red ] [FADH2 ] [H ] Q4 (21.5x) (2.672x) (0.670.5x) . (29) [H 2S] [NAD ] [Fd ox ] [FAD]
Taken together, CO2 reduction in SOB comprises two steps: production of the reducing equivalents (Eq. 21 or 27) and the actual CO2 fixation (Eq. 18 or 24), as already pointed out by Kelly (1999). Thus, the total energy requirement of CO2 reduction is given by the sum of
ǻGr(RET) and ǻGr(CO2 fix). Analogously to Eq. 8, in our new approach we therefore calculate the thermodynamic energy conservation efficiency of autotrophic sulfur oxidation as
(1 y) ['Gr (RET) 'Gr (CO 2 fix)] İ II , (30) y 'Gr (O 2 red) where the corresponding ǻGr values are calculated as described above. It should be noted that
İII > İI because ǻGr(CO2 red) calculated by the traditional approach (Eq. 11) is always lower
94 II. Manuscripts than the sum ǻGr(RET) + ǻGr(CO2 fix).
2.5. Factorization of the new energy conservation efficiency
Analogously to the factorization of the traditional thermodynamic efficiency (see Eq. 17), the new thermodynamic efficiency İII defined by Eq. 30 can also be written as a product of partial efficiencies characterizing the processes involved in the autotrophic sulfur oxidation.
Specifically, because CO2 reduction in our new approach is divided into RET and CO2 fixation, expression 15 for the efficiency of ATP utilization must be written as
'Gr (RET) 'Gr (CO 2 fix) İ u,II , (31) 'Er (RET) 'Er (CO 2 fix) where ¨Er(RET) and ¨Er(CO2 fix) describe the energy requirements, in the form of ATP, of the RET and CO2 fixation reactions, respectively. This implies that İII can be factorized as
İ II İSO İ u,II İ t . (32)
To make the influence of the RET and CO2 fixation reactions more explicit, we define their corresponding efficiencies as
'Gr (RET) İ RET (33) 'Er (RET) and
'Gr (CO 2 fix) İ CO2 . (34) 'Er (CO 2 fix)
This makes it possible to write İII as a product
1 İ II İSO İ t İ CO2 İ RET [D İ RET (1D) İ CO2 ] (35) where the parameter Į is defined as
'G (CO fix) D r 2 . (36) 'Gr (RET) 'Gr (CO 2 fix)
Previous work has already characterized the realistic ATP requirements of CO2 fixation. Assuming 0% oxygenase activity of RuBisCO, ATP requirements of the Calvin
95 3. STOICHIOMETRY AND EFFICIENCY OF SULFUR OXIDATION
cycle are 3 ATP per CO2, whereas ATP requirements of the rTCA cycle are ~1.67 ATP per
CO2 (Bar-Even et al., 2010). As pointed out by Kelly (1999), these requirements can be considered constant. Assuming the ATP energy content of 41 kJ (mol ATP)-1, this translates -1 -1 into Er(CO2 fix) of 123 kJ (mol CO2) and 68.3 kJ (mol CO2) for the Calvin and rTCA cycle, respectively. Considering the ¨Gr(CO2 fix) values calculated by Bar-Even et al. (2010)
(see Section 1.3), the energy conservation efficiency of the CO2 fixation can therefore be considered as a known parameter: İCO2 = 0.57 for the Calvin cycle and İCO2 = 0.94 for the rTCA cycle (Bar-Even et al., 2010). Similarly, the parameter Į has also a known value depending on the parameter x and on the CO2 fixation pathway employed by the SOB, as follows from Eqs. 22, 28 and 36.
Taken together, this means that although the efficiencies İSO, İt and İRET are generally unknown, they are related through Eq. 35. This makes it possible to calculate any one of these efficiencies provided that the other two are known. Since this is not possible at the current state of knowledge, Eq. 35 can be used to estimate their minimal values. Of specific interest in this study is the minimal value of the efficiency of the sulfur oxidation pathway,
İSO. This value is reached when the efficiencies İRET and İt are maximal (i.e., İRET = İt = 1), which gives
D (1 y) ['Er (RET) 'Gr (CO 2 fix)] İ SO,II,min İ II , (37) İ CO2 (1D) y 'Gr (O 2 red) as follows from Eqs. 30, 35 and 36. It should be noted that İSO,II,,min > İSO,I,min. Our new approach therefore provides a more constrained range of possible efficiencies of sulfur oxidation coupled to O2 reduction in comparison to the traditional approach.
2.6. Generalized equations and efficiencies for other reduced sulfur oxidation processes
Many SOB can use alternative reduced sulfur species for the reduction of O2 and CO2, such as thiosulfate. Additionally, in anoxic environments or habitats that are characterized by - fluctuating conditions, such as hydrothermal vents, also the reduction of NO3 has to be - considered as a possible sink of electrons during reduced sulfur oxidation. NO3 can either be + reduced partially to N2 by denitrification or completely to NH4 by dissimilatory nitrate reduction to ammonia (DNRA) to generate energy for CO2 fixation. The generalized mass-
96 II. Manuscripts balanced equations for these processes are derived analogously as Eqs. 1–7 and are given in the Supplementary Table S1. Furthermore, the corresponding energy conservation efficiencies İI, İII and İSO,II,min are calculated from expressions similar to Eqs. 8, 30 and 37 with the exception that the ǻGr values in the nominator and denominator are calculated based on the stoichiometry of the corresponding energy-conserving and energy-generating reaction. We implemented these calculations in R (www.cran.r-project.org) as functions and scripts that can be freely downloaded from the internet (http://nanosims.geo.uu.nl/SOX; Supplement 1).
3. Results
3.1. Stoichiometry of autotrophic sulfur oxidation from rate measurements Here we demonstrate how the generic theoretical framework can be used to rapidly evaluate the stoichiometry of autotrophic reduced sulfur oxidation from rate measurements of the reactants involved. Additional examples are provided in Supplement 2.
The first example involves marine Beggiatoa strain MS-81-6. Nelson et al. (1986) reported that this strain performed aerobic sulfide oxidation coupled to CO2 fixation with the
O2:H2S consumption ratio of ~1.65 and the CO2:H2S consumption ratio of 0.35. Using
Table S1A, these experimental values lead to equations ȞO2/ȞH2S = y(2–1.5x) = 1.65 and
ȞCO2/ȞH2S = (1–y)(2–1.5x) = 0.35, which yield y = 0.825 and x = 0. As follows from Table S1A, this implies that (i) the aerobic oxidation of sulfide in this strain was performed according to equation
2- + H2S + 1.65 O2 + 0.35 CO2 + 0.35 H2O Æ SO4 + 0.35 CH2O + 2 H ,
(ii) sulfate was the exclusive product, and (iii) 82.5% of the sulfide pool was used in the energy gaining reaction with oxygen (Eq. 3) while the remaining 17.5% was used for CO2 fixation (Eq. 6). This is consistent with the conclusions of Nelson et al. (1986).
The second example deals with SOB that live in symbiosis with tubeworms Riftia. -1 -1 Girgius et al. (2002) reported that the sulfide consumption rate of 6.75 μmol H2S g h by the whole-worm symbiosis was accompanied by the total rates of O2 and CO2 consumption -1 -1 -1 -1 of 12.4 μmol O2 g h and 12.45 μmol CO2 g h , respectively. Assuming that host respiration constitutes 25% of the whole-worm O2 consumption (see Supplement 3), the
97 3. STOICHIOMETRY AND EFFICIENCY OF SULFUR OXIDATION
-1 -1 estimated rates of O2 and CO2 consumption by the symbionts are 9.3 μmol O2 g h and -1 -1 15.55 μmol CO2 g h , respectively. This directly translates into equations ȞO2/ȞH2S = y(2–
1.5x) = 9.3/6.75 = 1.38 and ȞCO2/ȞH2S = (1–y)(2–1.5x) = 15.55/6.75 = 2.3, which yield x = – 1.12, y = 0.375 and the corresponding equation
0 2- + H2S + 1.12 S + 1.38 O2 + 2.3 CO2 + 3.42 H2O Æ 2.12 SO4 + 2.3 CH2O + 4.24 H .
This means that during the particular experiment reported by Girgius et al. (2002) the Riftia symbionts probably oxidized not only H2S but also a relatively large amount of stored zero- valent sulfur to fix CO2, with 37.5% of this total pool of reduced sulfur used for energy generation. This result is reasonable considering that the sum of the O2 and CO2 consumption rates cannot be larger than double the sulfide consumption rate if aerobic sulfide oxidation is 2- performed with H2S and SO4 as the only reduced and oxidized sulfur species, respectively (see Table S1A).
3.2. Energy conservation efficiency from stoichiometry The energy conservation efficiency of autotrophic sulfur oxidation can be calculated once the stoichiometry of the generalized equation is known. First, we demonstrate this calculation for Beggiatoa strain MS-81-6 analysed above using the traditional calculation approach and 0 assuming standard biochemical conditions. Substitution of x = 0 and of the ǻGf values 0 -1 tabulated in Thauer et al. (1977) to Eqs. 10–11 yields ǻGr (O2 red) = -829 kJ mol and 0 -1 ǻGr (CO2 red) = 145 kJ mol . Subsequent substitution of these values together with y = 0.825 obtained above to Eq. 8 yields İI = 0.037. This is consistent with the efficiency value of 0.038 obtained by Nelson & Hagen (1995). The small discrepancy is most likely due 0 -1 to the fact that Nelson & Hagen (1995) used ǻGf (O2) = 0 kJ mol for the standard Gibbs energy of formation of O2, which corresponds to O2 gas, while we used the value of 16.4 kJ -1 mol , which corresponds to dissolved O2.
Traditionally, the efficiency value of 0.037 is interpreted such that Beggiatoa strain MS-81-6 conserves 3.7% of the energy gained by aerobic sulfide oxidation into biomass. Based on our analysis above (Eq. 17), a more accurate interpretation is that 0.037 is the product of the partial efficiencies of (i) ATP generation by O2 reduction (İSO), (ii) ATP utilization for CO2 fixation (İu) and (iii) the transfer of ATP gained by O2 reduction for CO2
98 II. Manuscripts fixation and thus not for cellular maintenance (İt). For example, an assumption that
Beggiatoa strain MS-81-6 uses ATP exclusively for CO2 fixation and not for any other biochemical reaction (İt = 1), and that its ATP utilization for CO2 fixation occurs without loss of energy (İu = 1), would imply that the efficiency of ATP generation by O2 reduction in this strain is İSO = İI/(İtİu) = 0.037.
In the second example we analyse the difference between the efficiency values calculated according to the traditional (İI; Eq. 8) and our new approach (İII; Eq. 30). We do this for aerobic thiosulfate oxidizers Sulfurimonas denitrificans and Thioalkalivibrio versutus, which fix CO2 using the rTCA cycle (Hügler et al., 2005) and the Calvin cycle, respectively. Based on the substrate consumption rates reported by Hoor (1981) and Sorokin et al. (2001) we calculated x = 0 and y = 0.833 for S. denitrificans and x = 0 and y = 0.844 for T. versutus (see Supplement 4). Using these values in the traditional approach, we obtained the efficiency of İI = 0.0314 for S. denitrificans and İI = 0.0289 for T. versutus, whereas our new approach gave values İII = 0.0799 for S. denitrificans and İII = 0.0796 for T. versutus.
We see that our new approach yields substantially larger efficiency values than the traditional approach. This is generally because the new approach accounts for the fact that
CO2 reduction by SOB is performed in two separate steps (i.e., the reduction of an electron carrier via reverse electron transport followed by the reduction of CO2 coupled to the electron carrier oxidation in the respective CO2 fixation pathway) and because the theoretical energy requirements of these two steps (ǻGr(RET) + ǻGr(CO2 fix); see Eq. 30) are substantially larger than the theoretical energy requirement of the net CO2 reduction reaction (ǻGr(CO2 red); see Eq. 8) considered in the traditional approach. Additionally we see that the relative difference between the efficiency values of the compared SOB depends on the choice of the approach, particularly if the SOB employ different CO2 fixation pathways. In this specific case the efficiency İI of S. denitrificans is by about 8% larger than that of T. versutus, whereas their İII values are practically identical. This is related to the fact that the ǻGr values of the RET and CO2 fixation reactions (Eq. 30) depend on the type of the electron carriers involved in the CO2 fixation pathways, which differ between the rTCA and Calvin cycle (compare Eqs. 18 and 21 vs. Eqs. 24 and 27).
As a last point, we illustrate an additional insight that can be gained from the factorized form of İII (see Eq. 35). The fact that S. denitrificans and T. versutus have practically
99 3. STOICHIOMETRY AND EFFICIENCY OF SULFUR OXIDATION
identical values of the overall efficiency İII might be misinterpreted by concluding that their corresponding partial efficiencies İSO, İRET, İCO2 and İt are also the same. However, we know that the efficiency İCO2 of the rTCA cycle used by S. denitrificans to fix CO2 is almost two- fold larger than that of the Calvin cycle employed by T. versutus. This implies that at least one of the efficiencies İSO, İRET and İt must differ between the two SOB. By calculating the minimum values of these efficiencies using Eqs. 35–37 we found that all of them are higher for T. versutus (İSO,II,min = İt,II,min = 0.1030 and İRET,II,min = 0.052) than for S. denitrificans
(İSO,II,min = İt,II,min = 0.0818 and İRET,II,min = 0.051). This suggests that T. versutus has a more efficient sulfur oxidation pathway (larger İSO or İRET) or lower cellular maintenance requirements (larger İt) than S. denitrificans. It should be noted that this conclusion can only be drawn based on our new calculation approach but not based on the traditional approach.
Therefore, our new approach, and particularly the value of İSO,II,min, is better suited to identify differences between sulfur oxidation pathways among SOB, particularly if the compared
SOB employ different CO2 fixation pathways.
3.3. Sensitivity of the calculated efficiency towards reactant concentrations Until now our calculations assumed standard biochemical conditions. However, as pointed out by Nelson and Hagen (1995) and Kelly (1999), the actual experimental conditions should be used when calculating the ǻGr values for the energy gaining and energy conserving reactions. In the literature on SOB activity these concentrations are in most cases not reported or not well constrained. Thus, it is important to evaluate possible errors introduced by this uncertainty in the calculation of the energy conservation efficiency. This is achieved through sensitivity analysis.
To perform this analysis, we varied the concentration of one of the reactants between 1 M and 1 nM, or the pH between 2 and 12, while keeping the other reactant concentrations as well as the stoichiometry of the reaction mediated by an SOB unchanged. In our formalism the variations of the reactant concentrations and pH are captured in the variation of the quotients Q (see Eqs. 12, 13, 23 and 29). As examples, we used the aerobic sulfide oxidizing bacteria Thermithiobacillus tepidarius and Halothiobacillus neapolitanus, which employ the Calvin cycle for CO2 fixation, and the denitrifying thiosulfate oxidizing bacteria Sulfurimonas hongkongienses and Sulfurimonas denitrificans, which employ the rTCA cycle
100 II. Manuscripts for CO2 fixation.
This analysis revealed that the calculated efficiency values are most sensitive to the + concentrations of CO2 (İI) and of the electron carriers such as NAD /NADH (İSO,II,min), whereas the sensitivity towards the concentration of the terminal electron acceptor (TEA) is 2- lowest (Fig. S2). Sensitivities towards pH, the reduced sulfur compound (e.g., S2O3 or H2S) 2- and SO4 (Fig. S2) as well as towards temperature between 0–30 °C (data not shown) are intermediate but also rather small. For instance, for the aerobic sulfide oxidizing bacteria T. tepidarius and H. neapolitanus a change in the concentration of NAD+ by three orders of magnitude would change the calculated value of İSO,II,min by ~14 %, while the same change in the concentration of CO2 would change İI by ~24 %. In contrast the calculated values of
İSO,II,min and İI would only change by ~4% if the concentration of O2 changed by three orders of magnitude. The errors introduced by uncertainties in the reactant concentrations are similar for the denitrifying thiosulfate oxidizing bacteria S. hongkongienses and S. denitrificans (Fig. S2).
As the aim of this study is to compare efficiencies among SOB, it is important to evaluate how the errors introduced by the uncertainties in the reactant concentrations would affect this comparison. To do this we considered two SOB whose efficiencies calculated at standard conditions differ, and determined the change in the reactant concentrations required to make the efficiencies equal. We illustrate the result on the same pairs of aerobic and anaerobic SOB as above. When using the traditional approach, the efficiency İI calculated for T. tepidarius would become equal to that of H. neapolitanus if the former were calculated 6 2- with a 10 -fold lower SO4 concentration than the standard concentration of 1M or at pH=10 2- instead of pH=7. In contrast, the difference in the SO4 concentration or pH would need to be 20 2- considerably larger (~10 -fold for SO4 , or at a pH§11) to achieve the same effect for the efficiency İSO,II,min calculated by our new approach (Fig. S2A). Similar conclusion can be drawn for S. hongkongienses and S. denitrificans (Fig. S2B) as well as for SOB that perform other types of autotrophic sulfur oxidation (data not shown). We therefore conclude that, although the sensitivities towards reactant concentrations are similar for both approaches, our new approach is more robust to resolve differences between SOB with respect to their energy conservation efficiency if the calculations are affected by an uncertainty in the concentrations of the reactants involved.
101 3. STOICHIOMETRY AND EFFICIENCY OF SULFUR OXIDATION
3.4. Efficiency of autotrophic sulfur oxidation in different strains of SOB The reason for different efficiencies of autotrophic sulfur oxidation is generally assumed to lie in the different biochemical pathways that the SOB employ for the oxidation of the reduced sulfur compound (e.g., Hagen and Nelson, 1997; Kelly, 1999). To gain more insights into this possible relationship, we compiled literature data on cultivated strains of SOB, calculated their energy conservation efficiencies, and listed them together with the available information on the identified sulfur oxidation and carbon fixation pathways and selected specific enzymes. To follow the outcomes of the above analyses, we used İSO,II,min to compare the SOB. Specifically, we first calculated the complete stoichiometry of sulfur oxidation coupled to TEA and CO2 reduction based on rates reported in the literature. As for most SOB these rates were obtained under not well-constrained experimental conditions, we could not include the concentration dependency of the ǻGr values (in the form of the reaction quotient Q) in our efficiency calculations. Thus, to still make a comparison possible, we chose to calculate the efficiency values at standard biochemical conditions.
The results of this compilation are summarized in Table 1 and Fig. 2. First, they show that the efficiencies cover a wide range with no apparent clustering according to the SOB phylogeny. Furthermore, they suggest possible links between the efficiency, the sulfur oxidation pathway and/or the type of TEA reductase. This is quite astonishing considering that the efficiency values were calculated from rate measurements performed under different experimental settings (e.g., batch reactor studies, continuous cultivation) and that coherent data on both stoichiometry and biochemical pathways employed by SOB are rather limited.
With regard to aerobic thiosulfate oxidizing bacteria, for which the available literature data is most abundant, our analysis shows that autotrophic thiosulfate oxidation measured under aerobic conditions appears to be performed with the highest efficiency if the SOB are facultatively anaerobic (indicated by asterisk in Fig. 2), a trend noticed already, for instance, by Kelly (1982). Concerning the pathways of thiosulfate oxidation, SOB equipped with the enzyme reverse dissimilatory sulfite reductase (rDSR), which is involved in the branched pathway (red arrows in Fig. 1), tend to have the highest efficiencies among the aerobic thiosulfate oxidizers, whereas those relying on the Sox pathway (green arrows in Fig. 1) appear to have the lowest efficiencies. In contrast, no clear patterns are apparent for the SOB employing the SI4 pathway (blue arrows in Fig. 1), whose efficiencies span from low to high
102 II. Manuscripts values. The same is true for the APS pathway for sulfite oxidation, which can be part of both the branched and SI4 pathways (Fig. 1).
Aerobic sulfide oxidizing bacteria employing pathways that involve oxidation of zero-valent sulfur (black arrows in Fig. 1) to sulfite via rDSR (red arrows in Fig. 1) have the highest efficiency (Table 1). This is similar to the pattern identified for the aerobic thiosulfate oxidizing bacteria. Most strikingly, extremely high efficiencies of energy conservation are found among aerobic sulfide oxidizers that live in symbiotic association with invertebrates such as the Riftia tubeworm. Note that these high values are not the consequence of the fact that they were calculated at standard biochemical conditions. Specifically, the concentrations of the substrates and products involved would have to be in the kilomolar and picomolar range, respectively, to obtain efficiency values comparable to those of the other, less efficient SOB. Since it is unlikely that the host regulates the substrates and products in such an extreme range of concentrations, the high efficiency values of the sybmiotic SOB are realistic.
With respect to SOB that couple thiosulfate oxidation to denitrification, the data available in the literature is very limited. Nevertheless, the present data indicate that SOB equipped with a membrane-bound nitrate reductase system (Nar) have in general high efficiencies whereas those relying exclusively on a periplasmic nitrate reductase system (Nap), namely S. denitrificans, have a very low energy conservation efficiency (Table 1).
103 3. STOICHIOMETRY AND EFFICIENCY OF SULFUR OXIDATION
Figure 2: Energy conservation efficiencies of SOB performing aerobic thiosulfate oxidation (A), thiosulfate oxidation coupled to denitrification (B) and aerobic sulfide oxidation (C). Values were calculated using experimental data in the literature (see Table 1) based on the traditional approach (İI) and our new approach (İSO,II,min). Asterisks in panel A indicate that the SOB are facultatively anaerobic.
104 Table 1: Characteristics of autotrophic sulfur oxidizing bacteria (SOB) derived from literature data and calculated in this study (I, II, SO,II,min, CO2:TEA). C-fixation Sulfur oxidation pathways Selected enzymes 1 2 3 4 5 pathway 12 x y I (%) II (%) SO,II,min(%) CO2:TEA PB Ref
6 7 8 9 10 11 Calvin rTCA Branched Sox SI4 APS rDSR SDO Nar Nap
Thiobacillus denitrificans 0 0.710 6.39 17.59 22.76 0.41 ! X13 X X X X T
0 0.740 5.50 15.13 19.58 0.35
Thiothrix CT3 0 0.738 5.55 15.29 19.78 0.36 " X
Beggiatoa str. D-402 0 0.761 4.91 13.52 17.50 0.31 " X X X X 8
Thermothrix thiopara 0 0.783 4.34 11.93 15.44 0.28 ! X 9, 10
Thermithiobacillus tepidarius 0 0.804 3.81 10.50 13.58 0.24 " X X 5, 11, 12
Thioalkalivibrio versutus 0 0.844 2.90 7.96 10.30 0.18 " X X 13, 14
Acidithiobacillus ferrooxidans 0 0.848 2.80 7.72 9.99 0.18 " X X X 3, 16, 17, 18
Paracoccus versutus 0 0.853 2.70 7.42 9.60 0.17 X X 10, 15
Halothiobacillus neapolitanus 0 0.859 2.57 7.07 9.15 0.16 " X X X X 3, 19, 20
Thiomicrospira thioparus 0 0.863 2.48 6.84 8.85 0.16 ! X X X 20, 21
Sulfurimonas denitrificans 0 0.833 3.13 7.99 8.18 0.20 X X 6, 39, 40
Acidithiobacillus thiooxidans 0 0.874 2.26 6.21 8.03 0.14 " X X X X 10, 22–24
Halothiobacillus halophilus 0 0.875 2.23 6.15 7.96 0.14 " X X 25–27 Aerobic thiosulfateoxidizers Thiothrix ramosa 0 0.875 2.23 6.15 7.96 0.14 " X X X 5, 28
Thioalkalispira microaerophila 0 0.884 2.05 5.65 7.31 0.13 " X 14, 29
Thioalkalimicrobium aerophilum 0 0.891 1.91 5.27 6.82 0.12 " X X 13
Thiomicrospira halophila 0 0.891 1.91 5.27 6.82 0.12 " X X? X 5, 30
Thioalkalibacter halophilus 0 0.891 1.91 5.27 6.82 0.12 " X X? 5, 31
Thiobacillus thioparus 0 0.898 1.78 4.89 6.33 0.11 ! X X X X X X 5, 17, 32–36
Thioclava pacifica 0 0.913 1.49 4.10 5.31 0.10 X X? 37
Thioalkalimicrobium sibericum 0 0.916 1.43 3.95 5.11 0.09 " X X? 13
Thiomicrospira sp. Strain L-12 0.2 0.921 1.29 3.91 4.99 0.09 " X X X 38, 39 105 106 Thiobacillus denitrificans 0 0.775 5.15 14.18 18.35 0.36 ! X X X X X 1, 4, 6,
0 0.812 4.11 11.31 14.63 0.29 40
Sulfurimonas hongkongensis 0 0.852 3.08 7.85 8.04 0.22 X X 41
Sulfurimonas gotlandica 0 0.875 2.53 6.45 6.61 0.18 X X X X X 42, 43
Sulfurimonas gotlandica-like 0 Epsilonproteobacterium 0.887 2.26 5.76 5.89 0.16 X X 43, 44 (environmental sample)
Denitrifying thiosulfate oxidizers thiosulfate Denitrifying Sulfurimonas denitrificans 0 0.902 1.93 4.91 5.03 0.14 X X X 6, 45
Riftia pachyptila symbionts -1.12 0.375 30.52 77.27 98.86 1.67 " X 5 5
Solemya reidi symbionts 0.17 0.429 22.83 59.80 76.87 1.33 " X X X X X 49, 50
Riftia pachyptila symbionts 0.33 0.500 16.80 44.50 57.29 1 " X X X X X 5, 46, 48, 51
Beggiatoa str. MS-81-1c 0.05 0.641 9.72 25.30 32.50 0.56 " X X 52
Ridgeia piscesae symbionts 0 0.714 6.98 18.13 23.29 0.40 " X X 53
Thermithiobacillus tepidarius 0 0.808 4.14 10.76 13.81 0.24 " X X 12, 54 Aerobic sulfideoxidizers Beggiatoa str. MS-81-6 0 0.825 3.70 9.60 12.33 0.21 " X X 52
Halothiobacillus neapolitanus 0 0.859 2.86 7.43 9.54 0.16 " X X X X 20, 53, 51
1 I is the efficiency of energy conservation calculated at biochemical standard conditions, i.e., 1 M concentration of all reactants, 25°C, pH 7, according to the traditional approach. 2 II is the efficiency of energy conservation calculated at biochemical standard conditions according to our new approach. 3 SO,II,min is the minimum efficiency of the sulfur oxidation coupled to TEA reduction calculated at biochemical standard conditions based on our new approach. 4 - TEA is the terminal electron acceptor in the energy generating reaction (O2 in aerobic processes or NO3 in denitrification). The complete equations for the corresponding autotrophic sulfur oxidation reactions are given in Table S2. 5PB refers to the proteobacterial class. 6Calvin refers to the Calvin-Benson-Bassham cycle. 7TCA refers to the reverse tricarboxilic acid cycle. 8rDSR refers to the reverse dissimilatory sulfite reductase. 9SO refers to sulfur dioxygenase. 10Nar refers to the membrane-bound nitrate reductase system. 11Nap refers to the periplasmic nitrate reductase system. 12References: 1: (Justin and Kelly, 1978); 2: (Aminuddin, 1980); 3: (Kelly, 1999); 4: (Beller et al., 2006); 5: (Meyer et al., 2007); 6: (Hoor, 1981); 7:(Rossetti, 2003); 8: (Grabovich et al., 2001); 9: (Brannan and Caldwell, 1983); 10: (Mason et al., 1987): 11: (Lu and Kelly, 1988); 12: (Wood and Kelly, 1986); 13: (Sorokin et al., 2001) ; 14: (Sorokin et al., 2013) ; 15: (Friedrich et al., 2001); 16: (Eccleston and Kelly, 1978); 17: (Rohwerder and Sand, 2003); 18: (Pronk et al., 1990); 19: (Hempfling and Vishniac, 1967); 20: (Kelly, 1982); 21: (Lengeler et al., 2009); 22: (Masau et al., 2001); 23: (Kamimura et al., 2005); 24: (Nakamura et al., 2014); 25: (Wood and Kelly, 1991); 26: (Kelly and Wood, 2000); 27: (Shi et al., 2011); 28: (Odintsova et al., 1993); 29: (Sorokin, 2002); 30: (Sorokin et al., 2006); 31: (Banciu et al., 2008); 32: (Smith and Kelly, 1988); 33: (Peck, 1960); 34: (Suzuki, 1974) ; 35: (Trüper, 1982); 36: (Loy et al., 2009); 37: (Sorokin et al., 2005); 38: (Ruby and Jannasch, 1982); 39: (Sievert et al., 2007); 40: (Schedel et al., 1975); 41: (Cai et al., 2014); 42: (Bruckner et al., 2013); 43: (Grote et al., 2012); 44: (Brettar et al., 2006); 45: (Sievert et al., 2008); 46: (Nelson and Hagen, 1995); 47: (Girguis et al., 2002); 48: (Markert et al., 2007); 49: (Anderson et al., 1987); 50: (Stewart et al., 2011); 51: (Girguis and Childress, 2006); 52:(Hagen and Nelson, 1997); 53: (Nyholm et al., 2008); 54:(Kelly et al., 1993) ; 55:(Veith et al., 2012); 56:(Kelly et al., 1997) 12 ‘X’ indicates the presence of a pathway or enzyme; ‘X?’ indicates that the presence of a pathway or enzyme is likely but not conclusively confirmed. 13 II and SO,II,min were calculated based on the assumption that the symbiotic SOB employ the Calvin cycle for CO2 fixation. 3. STOICHIOMETRY AND EFFICIENCY OF SULFUR OXIDATION
3.5. Stoichiometry from the efficiency of energy conservation and limited rate measurements Most stoichiometries and efficiency values listed in Tables 1 and S2 were calculated based on rate measurements during which the studied SOB oxidized the sulfur compound completely to sulfate. However, for many SOB main products of sulfur oxidation may include also other sulfur species, such as zero-valent sulfur (e.g. Ghosh and Dam, 2009). To calculate the 2- 0 stoichiometry of autotrophic sulfur oxidation at an arbitrary SO4 :S production ratio, one generally needs to provide gross rates of three reactants involved. However, accurate quantification of three gross rates is often experimentally difficult. Here we demonstrate that the stoichiometry of autotrophic sulfur oxidation can be estimated if the consumption/production rates of only two reactants involved are measured, provided that certain additional conditions characterizing the activity of an SOB are known.
Our framework shows that the stoichiometry is completely determined by the values of the parameters x and y. To find these values, two constraints are needed. The first constraint is obtained by measuring the rates of two reactants involved, whereby their ratio directly provides a relationship between x and y, as follows from the generalized mass balanced equations in Table S1. The second constraint is obtained from the knowledge of how the CO2:TEA consumption ratio (essentially the parameters y; see Table S1) or the 2- 0 energy conservation efficiency (e.g., İII), varies depending on the SO4 :S production ratio, i.e., on the parameter x. Unfortunately, such information is presently not available because, to the best of our knowledge, the complete stoichiometry of autotrophic sulfur oxidation has never been measured over variable x in an isolated SOB. Therefore, to illustrate the general approach based on our generic framework, we first assume that either the CO2:TEA ratio or the efficiency İII is independent of x and equal to the value calculated from the known stoichiometry at x = 0 (see Table 1). We use the aerobic sulfide oxidizing Beggiatoa str. MS- 81-6 as an example.
Based on the experimental data reported by Nelson et al. (1986) we showed above 2- that this SOB performs aerobic sulfide oxidation to SO4 (i.e., x = 0) with the CO2:O2 consumption ratio of 0.21, O2:H2S ratio of 1.65 and, when accounting for the energy requirements of the CO2 fixation pathway, with the efficiency of İII = 9.60 % (Table 1 and
S2). Let us now suppose that in a separate experiment the O2:H2S consumption ratio by this
107 3. STOICHIOMETRY AND EFFICIENCY OF SULFUR OXIDATION
strain would be 0.5. What would be the corresponding stoichiometry?
As follows from Table S1A, the generalized stoichiometric coefficients must be related as ȞO2/Ȟ+6 = O2:H2S = 0.5, which yields the first relationship between x and y, namely y(2–1.5x) = 0.5. If we additionally assume that the energy conservation efficiency İII is independent of x and equal to the value at x = 0 (i.e., 9.60 %; Table 1), the second relationship between x and y is provided by Eq. 30. By solving these two equations numerically (implemented in a script written in R available on the internet at http://nanosims.geo.uu.nl/SOX; Supplement 1) we obtained x = 0.920 and y = 0.807 and thus the stoichiometry
2- 0 + H2S + 0.5 O2 + 0.12 CO2 Æ 0.08 SO4 + 0.92 S + 0.12 CH2O + 0.8 H2O + 0.16 H .
As mentioned above and applied in the work of Nelson et al. (1986), the alternative approach assumes that the growth yield per mole of TEA utilized is independent of x and equal to the value determined at x = 0 (i.e., CO2:O2 = 0.21). Using the generalized stoichiometric coefficients (see Table S1), this implies ȞCO2/Ȟ2 = (1–y)/y = 0.21 and thus the value of y = 0.826. Combination of this value with the first constraint (i.e., y(2–1.5x) = 0.5; see above) then yields x = 0.929 and thus the stoichiometry
2- 0 + H2S + 0.5 O2 + 0.11 CO2 Æ 0.07 SO4 + 0.93 S + 0.11 CH2O + 0.82 H2O + 0.14 H .
We see that the efficiency-based and CO2:TEA-based approach give a very similar stoichiometry of the overall sulfide oxidation reaction, suggesting that they are almost equivalent. To verify this, we performed the same calculations over all possible values of x as well as for different aerobic and anaerobic SOB. If the two approaches were equivalent, the calculated CO2:TEA consumption ratios would not change with x. As shown in Fig. 3A-B, this is approximately the case, since the calculated CO2:TEA ratios vary by less than ~20% over the complete interval of x. Additionally, this variability is decreased if the reactant concentrations are decreased from values at standard biochemical conditions to a more environmentally relevant range (compare solid and dashed lines in Fig. 3A-B). This means that the possible error made when determining the stoichiometric coefficients (for an arbitrary value of x) based on the CO2:TEA-based approach or the efficiency-based approach will be relatively small (<20%).
108 II. Manuscripts
2- 0 Figure 3: CO2:TEA ratios in selected SOB calculated over the complete range of SO4 :S production ratios (represented by 0 The assumption that either the CO2:TEA ratio or the efficiency İII is independent of x is likely not generally applicable to all SOB. Indeed, Høgslund et al. (2009) showed that 0 sulfide oxidation to S in bundles of Thioploca spp. is completely uncoupled from CO2 fixation (CO2:TEA = 0), implying y = 1 and İII = 0 at x = 1. It is therefore likely that in some SOB the variability of the CO2:TEA ratio or of the efficiency İII with the parameter x is better approximated by a linear function that reaches a maximum at x = 0 and some other value, such as zero, at x = 1. By assuming that this is the case for the efficiency İII, we found that the calculated CO2:TEA ratio also very closely follows a linear trend (Fig. 3C-D). This means that, again, the possible error made when determining the stoichiometric coefficients (for an arbitrary value of x) based on the CO2:TEA-based approach or the efficiency-based approach will be very small. 109 3. STOICHIOMETRY AND EFFICIENCY OF SULFUR OXIDATION Overall, these examples demonstrate how our framework can be applied to calculate the stoichiometry of autotrophic sulfur oxidation reactions from two experimentally determined rates (e.g., the consumption rates of the oxidant and of the reduced sulfur species) 2- 0 at an arbitrary SO4 :S production ratio. Prerequisite for such calculations is the knowledge of a function that describes the dependency of the CO2:TEA yield or of the energy 2- 0 conservation efficiency on the SO4 :S production ratios parameterized by x. When, due to the lack of experimental data, these functions need to be approximated based on experiments performed at only one or two values of x, the results obtained from the CO2:TEA-based approach and the efficiency-based approach will be very similar. 4. Discussion 4.1. Calculation of energy conservation efficiency in SOB The aim of calculating the energy conservation efficiency is to gain insights into the relationship between the ATP requirements and ATP yields of microbially mediated processes (Hoor, 1981; Baas-Becking and Parks, 1927). For chemolithoautotrophic sulfur oxidation the efficiency is calculated as the ratio between the Gibbs free energy required to fix CO2 and the Gibbs free energy gained from the oxidation of the reduced inorganic compound (Kelly, 1982, 1990). However, this ratio is generally difficult to interpret because of the complexity of the ATP-consuming and ATP-producing pathways involved. Specifically, ATP is gained during the transfer of electrons from the reduced sulfur compound to the TEA (e.g., O2 or - NO3 ) in the diverse sulfur oxidation pathways (Fig. 1) and additionally during substrate-level phosphorylation (e.g., in the APS pathway, Fig. 1). In contrast, ATP is required during the reverse electron transport from the sulfur compound to the electron carrier such as NAD+ (Elbehti et al., 2000), the reduction of CO2 by the electron carrier in the carbon fixation pathway, and for all other biochemical pathways associated with cell maintenance. Since each of the enzymes involved in the diverse steps of sulfur compound oxidation has a specific electron acceptor in the electron transport chain, the amount of proton-motive-force, and thus ATP, generated per electron transferred depends on the level at which the electron enters the transport chain and the type of TEA reductase. Similarly, the ATP requirement of the electron carrier reduction depends on the level at which electrons enter the transport chain. Last but not least, ATP requirements of the CO2 fixation also depend on the specific carbon fixation pathway used. 110 II. Manuscripts The traditional calculation approach of the energy conservation efficiency is based on the Gibbs free energy of the net reactions involved in energy consumption (Eq. 6) and energy generation (Eq. 3) (Nelson and Hagen, 1995; Kelly, 1999). Since this approach does not consider the above-mentioned complexity of the pathways involved in the autotrophic sulfur oxidation, especially the influence of the CO2 fixation pathway and reverse electron transport reactions, it cannot identify which biochemical pathway or reaction has the highest impact on the calculated efficiency. In contrast, our new approach makes this differentiation at least partially possible. Specifically, the calculated value of İII represents the overall energy conservation efficiency of sulfur oxidation that accounts for the efficiency of the CO2 fixation pathway employed by the SOB. Additionally, by formulating this overall efficiency as a product of partial efficiencies of sulfur oxidation coupled to TEA reduction (İSO), energy utilization for reverse electron transfer and CO2 fixation (İRET and İSO) and the “transfer efficiency” (İt) (see Eq. 35 and Fig. S1B), our new approach makes it possible to constrain the range of values that these partial efficiencies can have. Of particular utility is the minimum efficiency of sulfur oxidation coupled to TEA reduction (İSO,II,min), which allows the comparison of SOB with respect to the efficiency of their sulfur oxidation pathways independent of the type and efficiency of their CO2 fixation pathway. 4.2. Variability of energy conservation efficiencies amongst SOB Possible links between the energy conservation efficiency and pathways of sulfur oxidation have been extensively discussed before (Nelson and Hagen, 1995; Hagen and Nelson, 1997; Kelly, 1982, 1999, 2003). By combining the efficiency values calculated by our new approach with additional information derived from diverse research approaches (e.g., physiological studies, enzyme assays and genome sequencing), our analysis provides new insights in this discussion. Specifically, it identifies the importance of rDSR and of the TEA reduction step and questions the role of the APS pathway. 4.2.1. Role of rDSR As noticed by Kelly (1982), facultative anaerobic SOB appear to generally have a higher energy conservation efficiency than obligate aerobic SOB, a trend clearly supported also by 111 3. STOICHIOMETRY AND EFFICIENCY OF SULFUR OXIDATION our results (Fig. 2). This pattern can be understood by noticing that most facultative anaerobic thiosulfate oxidizing SOB are equipped with the “branched” pathway for thiosulfate oxidation (red arrows in Fig. 1; Table 1), which is characterized by a truncated Sox system. In SOB that possess a Sox system that is not truncated (green arrows in Fig. 1) a thiosulfate-oxidizing multi-enzyme (TOMES) complex catalyses the complete oxidation of thiosulfate to sulfate, whereby the oxidation of the intermediate zero-valent sulfur is catalyzed by the soxCD enzymes of this complex (e.g. Ghosh and Dam, 2009). However, these enzymes are missing in the branched pathway and, instead, the transiently stored zero- valent sulfur is oxidized, after its reactivation to glutathione persulfide (GSSH), by other enzymes such as rDSR or the oxygen-dependent sulfur dioxygenase (SDO) (Rohwerder and Sand, 2003). While the complete TOMES complex directly channels all electrons derived from thiosulfate oxidation into the electron transport chain at the level of cyt c (Kelly et al., 1997), rDSR catalyses the cytoplasmic oxidation of GSSH to sulfite and donates the 6 electrons into the electron transport chain eventually at the level of quinone (Holkenbrink et al., 2011). This rDSR-mediated mechanism might therefore provide more ATP and thus eventually lead to a higher energy conservation efficiency as compared to the Sox pathway. rDSR is suspected to be the most ancient enzyme equipment of SOB and was probably employed by anaerobic anoxygenic phototrophs already in the early Proterozoic era (Meyer and Kuever, 2007). The fact that rDSR is only conserved in extant facultative anaerobic SOB (and phototrophs) might therefore explain the apparent link between the high energy conservation efficiency and the capability of an anaerobic life style. 4.2.2. Role of the terminal electron acceptor reduction step The importance of the type of TEA reductase for the energy conservation efficiency is suggested by our data compiled for the denitrifying SOB (Table 1). Namely, Sulfurimonas denitrificans exclusively relies on a periplasmically oriented nitrate reductase system (Nap) and does not possess a membrane-bound cytoplasmically oriented nitrate reductase system (Nar) (Sievert et al., 2008). Nitrate oxidation via Nap was shown not to contribute to the generation of proton-motive-force and thus ATP (Stewart et al., 2002), which is consistent with the rather poor energy conservation efficiency of this Epsilonproteobacterium. Most other known important denitrifying SOB, such as the large marine Beggiatoa and Thioploca, 112 II. Manuscripts are equipped with a membrane-bound nitrate reductase system (Nar) (e.g. Crossman, 2007), which oxidizes nitrate on the cytoplasmic side of the membrane and thus contributes to the generation of proton motive force (Simon et al., 2008; Simon and Klotz, 2013). This is consistent with their generally higher energy conservation efficiency (Table 1). Thus, in - denitrifying SOB the enzymes directly involved in TEA (NO3 ) reduction appear to have at least a similar impact on the energy conservation efficiency as the sulfur oxidation pathway. 4.2.3. Role of the APS pathway The role of the pathway of sulfite oxidation to sulfate, where the sulfite is derived, e.g., from sulfide oxidation by rDSR (Fig. 1), has been extensively discussed in the past (e.g. Kelly, 2003). This oxidation step can be mediated either by sulfite:cyt c:oxidoreductase (SOAR) or via the APS pathway (Fig. 1). Depending on the pathway of sulfur oxidation, SOARs are either associated with the TOMES complex or are complex-independent (Fig. 1). While SOARs have been identified in all sulfite oxidizing free-living SOB, the APS pathway appears to be an “extra” pathway that is not crucial for the operation of complete sulfur oxidation (Kappler and Dahl, 2001). It is known that the APS pathway allows additional ATP gain via substrate level phosphorylation before the electrons enter the electron transport chain (Aminuddin, 1980), and its use should thus lead to a higher overall energy conservation efficiency. However, we did not identify a clear link between the efficiency and the presence of the APS pathway in our present data (Table 1), suggesting that the electron transport processes are more important than this substrate level phosphorylation step. Nevertheless, the presence of the APS pathway might still be the cause of variations in efficiency among closely related species, such as Beggiatoa spp., as suggested by Nelson and Hagen (1995) and Hagen and Nelson (1997). Also, it has to be considered that the APS pathway in the anoxygenic phototroph Allochromatium vinosum only contributes to energy generation at high irradiances (Sánchez et al., 2001), which correspond to saturating availability of TEA and sulfur compound in chemolithotrophic SOB when sulfite oxidation via SOR might become the rate limiting step. Thus, also in SOB the APS pathway might operate only under certain growth conditions. It was not shown for all SOB listed in Table 1 that the APS pathway was active during the rate measurements, which might disguise a possible link between the calculated efficiency and the APS pathway. 113 3. STOICHIOMETRY AND EFFICIENCY OF SULFUR OXIDATION 4.3. Sulfur oxidation at a variable sulfate-to-elemental-sulfur product ratio An interesting outcome of our analysis was the way to calculate the stoichiometry of 2- 0 autotrophic sulfur oxidation at a variable SO4 :S production ratio from the rates of only two reactants involved (see Sec. 3.5). This calculation relied on an additional information about 2- 0 the activity of the SOB, namely on the relationship between the SO4 :S production ratio (parameter x) and the energy conservation efficiency or the CO2:TEA consumption ratio. Ideally, this relationship should be determined experimentally; however, since such data are presently not available for an isolated SOB, our calculation assumed two specific forms of this relationship: a constant or a linearly decreasing function of x. In the following we clarify the biochemical interpretation and limitations of these assumptions. The assumption of a constant CO2:TEA consumption ratio is equivalent to the assumption that the amount of ATP generated per electron transported to the TEA and the amount of ATP consumed per electron transported to the electron carrier (e.g., NAD+) are independent of the origin of the electrons, i.e., whether the electrons are derived from the first 0 0 2- (S production) or second (S to SO4 ) oxidation step. This is because the rate of TEA reduction linearly correlates with the rate of electrons transported (e.g., 4 electrons per O2). In contrast, the assumption of a constant energy conservation efficiency implicitly assumes 0 0 2- that both the first (reduced sulfur compound to S ) and second (S to SO4 ) oxidation steps have the same efficiency, i.e., the amount of ATP gained from TEA reduction and required for the electron carrier reduction per kJ of the calculated Gibbs free energy is the same for both oxidation steps. The assumption of a linear dependency of the CO2:TEA consumption ratio on x considers that the amount of ATP generated during electron transport to the TEA and the amount of ATP required during electron transport to the electron carriers depends on the origin of the electrons. Since the transported electrons derived from each oxidation reaction will have a specific constant potential to generate ATP, a linear dependency of the CO2:TEA ratio on x will result from the linear “mixing” of the electrons originating from the first and second oxidation step. Similarly, the assumption of a linear dependency of the efficiency on x is a consequence of the linear “mixing” of the efficiencies specific for the complete and incomplete sulfur oxidation reactions. It should be noted that a major shortcoming of the CO2:TEA-based approach is that it 114 II. Manuscripts does not consider the experimental conditions, such as reactant concentrations or the effect of the oxygen concentration on the efficiency of the CO2 fixation pathway (İCO2). These conditions are expected to affect the ATP yield and ATP consumption per transported electron. In contrast, the efficiency-based approach does allow to account for this. Specifically, if the conditions were well-constrained during the experimental determination of the relation between the efficiency and x, and during the independent measurement of two reactant rates, our framework would make it possible, at least theoretically, to precisely predict the complete stoichiometry of autotrophic sulfur oxidation, and thus primary productivity, at any experimental condition. 4.4. Interpretation of reactant fluxes in the environment In mixed microbial communities different processes that utilize the same reactants often spatially overlap, which hampers our ability to interpret the measured net consumption/production rates of the reactants involved. To illustrate how our framework can aid such interpretation, we consider microbial mats inhabited by one dominant SOB in addition to other bacteria. In such mats, it is often observed that, in the zone where sulfide is oxidized with oxygen, the ratio between the steady-state O2 and H2S consumption rates is 2 (e.g. Jørgensen et al., 2010). What does this ratio suggest about the S, O and C cycling in the system? 2- 0 Knowing that under steady-state conditions SOB exclusively produce SO4 and no S (Nelson et al., 1986), the O2:H2S ratio of 2 suggests complete aerobic sulfide oxidation to sulfate (e.g. Jørgensen et al., 2010). Although this interpretation is correct, it ignores carbon cycling within the sulfide oxidation zone. If carbon cycling is taken into account, as it should be because the SOB that mediate the aerobic sulfide oxidation must also grow, this interpretation must be reevaluated. Our framework implies that the general stoichiometry of aerobic sulfide oxidation coupled to CO2 fixation under steady state is (Table S1A) 2- + H2S + 2y O2 + 2(1 – y) CO2 + 2(1 – y) H2O Æ SO4 + 2(1 – y) CH2O + 2 H . Thus, the O2:H2S ratio due to the SOB activity is 2y, which is generally lower than 2 because 0 115 3. STOICHIOMETRY AND EFFICIENCY OF SULFUR OXIDATION consumption must be due to another kind of activity. A plausible candidate is aerobic respiration according to the stoichiometry 2(1 – y) O2 + 2(1 – y) CH2O Æ 2(1 – y) CO2 + 2(1 – y) H2O. This shows that when the two processes occur simultaneously the O2:H2S ratio of 2 can be achieved only if the stoichiometric coefficient for CO2 in the SOB-driven reaction is the same as that in the aerobic respiration reaction (i.e., 2(1–y)). Thus, assuming that the steady-state conditions are met, the O2:H2S ratio of 2 suggests that the biomass built up by the dominant SOB through aerobic sulfide oxidation is completely recycled within the sulfide oxidation zone via aerobic respiration. Note that this argument is independent of y and thus of the energy conservation efficiency of the dominant SOB. The efficiency only determines the fraction of the O2 flux that is used for organic carbon recycling. This analysis can be extended to cases when the O2:H2S consumption ratio in the sulfide oxidation zone is different from 2 while the stoichiometry of the SOB activity remains unchanged. Specifically, O2:H2S<2 would suggest export of the SOB-generated biomass out of the sulfide oxidation zone (e.g., for degradation by anaerobic processes), whereas O2:H2S>2 would indicate import of an external reductant (e.g., organic carbon) into, and its aerobic oxidation within, the zone. 4.5. Implications for niche differentiation among SOB Past and present research on SOB revealed that (i) sulfur oxidation pathways are highly diverse and characterized by a degree of redundancy (Fig. 1), (ii) the product of sulfide 0 2- oxidation can vary substantially (between S and SO4 ) even in one SOB dependent on growth conditions and phase, (iii) the energy conservation efficiency of SOB varies widely (Table 1), and (iv) SOB possessing one of the most ancestral enzymes involved in sulfur oxidation (rDSR) appear to be also the most efficient (Table 1). In the following we attempt a synthesis of these findings in the context of niche differentiation among SOB. To understand the apparent diversification towards lower energy conservation efficiency, it needs to be realized that efficiency is generally not the only measure of success in a given environment. Generally, the success is determined by the growth rate, which is a product of the substrate uptake rate and the growth yield (i.e., the amount of organic carbon 116 II. Manuscripts synthesized per unit of substrate utilized). Since the growth yield and efficiency are directly related (as follows from Table S1 and Eqs. 8 and 30), being efficient is only one of the possibilities for being successful. If organisms compensate their inefficient energy conservation by speed with which they utilize the substrate, they can be equally or even more successful than the efficient ones (e.g. Sorokin and Kuenen, 2005). These strategies are, however, likely dominant only in environments with unlimited substrates. The situation is different in environments where the substrates are limited, e.g., because of a limited external supply. In the context of SOB this can occur if the flux of the reduced sulfur compound (Sred) or of the TEA into the sulfur oxidizing zone is capped, e.g., by the rate of diffusion or by the substrate concentration in the seeping water delivering the substrate through advection. Under such circumstances, the flux of the external substrate supply will define the maximum substrate uptake rate. Ignoring the role of other phenotypes such as motility, this implies that the success of a specific SOB in the sulfur oxidizing zone will depend only on its energy utilization efficiency. A possible strategy for an SOB to become successful under such circumstances is to match its substrate uptake stoichiometry to the TEA:Sred ratio fixed by the environment. Such an SOB will utilize the available substrates completely, leaving nothing for potential competitors. Table S1 and equation 30 show that this can be achieved essentially by "adjusting" two parameters: y, and thus the energy conservation efficiency, and x, i.e., the 0 2- S :SO4 product ratio. Which of the parameters is "adjusted" depends on the time-scale at which the environmental setting (the TEA:Sred ratio) varies. Specifically, a long-term stable environment with a low TEA:Sred ratio should select for an SOB with a high efficiency, and vice versa. This could, for example, provide a possible explanation why SOB living in symbiotic association with invertebrates are highly efficient (Table 1). Specifically, symbiotic SOB need to compete with the host for O2 and thus likely face O2 limitation, which requires them to have evolved highly efficient substrate utilization. On the other hand, they can "afford" to be highly efficient because they do not have to retain flexibility in the presumably stable environment regulated by the host. In contrast, environments with short-term TEA:Sred fluctuations should select for SOB that possess mechanisms for rapid optimization of the substrate utilization stoichiometry. 117 3. STOICHIOMETRY AND EFFICIENCY OF SULFUR OXIDATION Phenotypic traits operating on short time-scales, such as motility coupled to chemotaxis or the ability to store TEA (nitrate), will likely play a role. However, another important 0 2- mechanism could be the ability to perform sulfur oxidation at a variable S :SO4 ratio. Our framework formulates this ability through the parameter x while the overall stoichiometry of the autotrophic sulfur oxidation reaction is additionally dependent on the parameter y.This makes it possible to predict the optimal range of sulfur oxidation efficiencies that an SOB should have to be successful in a given fluctuating environment. Specifically, this range should be such that the corresponding TEA:Sred utilization ratios during the incomplete (at x = 1) and complete (at x = 0) oxidation of Sred would match the range of the TEA:Sred fluctuations imposed by the environment. To illustrate this, we assume that the O2:H2S flux ratio available in the environment rapidly fluctuates between 0.45 and 1.7. To optimally utilize the available O2 and total sulphide (H2S) pool at all times, the SOB should vary the product of its sulphide oxidation 0 2- between two extremes: S when the O2:H2S ratio reaches 0.45, and SO4 when the O2:H2S ratio reaches 1.7. As follows from Table S1, the O2:H2S ratio is equal to y(2-1.5x), which yields y = 0.9 to satisfy the condition O2:H2S = 0.45 at x = 1 (incomplete sulfide oxidation) and y = 0.85 to satisfy the condition O2:H2S = 1.7 at x = 0 (complete sulfide oxidation). Assuming standard conditions and Calvin cycle, the efficiency of the most successful SOB should therefore vary between İII = 4.3% and İII = 8% for the incomplete and complete aerobic sulfide oxidation, respectively, with the corresponding stoichiometries varying between 0 H2S + 0.45 O2 + 0.05 CO2 Æ S + 0.05 CH2O + 0.95 H2O and 2- + H2S + 1.7 O2 + 0.3 CO2 + 0.3 H2O ĺ SO4 + 0.3 CH2O + 2 H . There are possibly other strategies that SOB living in fluctuating environments employ to become successful, including minimization of S0 production to prevent cell bursting (if S0 is stored intra-cellularly) or maximization of the growth yield. Although they can be explored within our theoretical framework, their thorough theoretical analysis would go beyond the scope of this study. Nevertheless, the examples and analyses discussed above demonstrate that the concepts and equations put forward in this study provide a generic 118 II. 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Thiomicrospira halophila sp. nov., a moderately halophilic, obligately chemolithoautotrophic, sulfur-oxidizing bacterium from hypersaline lakes. Int. J. Syst. Evol. Microbiol. 56, 2375–80. doi:10.1099/ijs.0.64445-0. Sorokin, D. Y., Tourova, T. P., Spiridonova, E. M., Rainey, F. A., and Muyzer, G. (2005). Thioclava pacifica gen. nov., sp. nov., a novel facultatively autotrophic, marine, sulfur-oxidizing bacterium from a near-shore sulfidic hydrothermal area. Int. J. Syst. Evol. Microbiol. 55, 1069–75. doi:10.1099/ijs.0.63415-0. Stewart, F. J., Dmytrenko, O., Delong, E. F., and Cavanaugh, C. M. (2011). Metatranscriptomic analysis of sulfur oxidation genes in the endosymbiont of Solemya velum. Front. Microbiol. 2, 134. doi:10.3389/fmicb.2011.00134. Stewart, V., Lu, Y., and Darwin, A. J. (2002). Periplasmic nitrate reductase (NapABC enzyme) supports anaerobic respiration by Escherichia coli K-12. J. Bacteriol. 184, 1314–23. Suzuki, I. (1974). Mechanisms of inorganic oxidation and energy coupling. Annu. Rev. Microbiol. 28, 85–101. doi:10.1146/annurev.mi.28.100174.000505. Thauer, R. K., Jungermann, K., and Decker, K. (1977). Energy conservation in chemotrophic anaerobic bacteria. Bacteriol. Rev. 41, 100–180. Trüper, H. G. (1982). “Microbial Processes in the Sulfur Cycle Through Time,” in Mineral Deposits and the Evolution of the Biosphere, eds. H. D. Holland and M. Schidlowski (Berlin, Heidelberg: Springer Berlin Heidelberg), 5–30. doi:10.1007/978-3-642-68463-0. Veith, A., Botelho, H. M., Kindinger, F., Gomes, C. M., and Kletzin, A. (2012). The sulfur oxygenase reductase from the mesophilic bacterium Halothiobacillus neapolitanus is a highly active thermozyme. J. Bacteriol. 194, 677–85. doi:10.1128/JB.06531-11. Wood, A. P., and Kelly, D. P. (1986). Chemolithotrophic metabolism of the newly-isolated moderately thermophilic, obligately autotrophic Thiobacillus tepidarius. Arch. Microbiol. 144, 71–77. doi:10.1007/BF00454959. Wood, A. P., and Kelly, D. P. (1991). Isolation and characterisation of Thiobacillus halophilus sp. nov., a sulphur-oxidising autotrophic eubacterium from a Western Australian hypersaline lake. Arch. Microbiol. 156, 277–280. doi:10.1007/BF00262998. 125 3. STOICHIOMETRY AND EFFICIENCY OF SULFUR OXIDATION ASSESSMENT OF THE STOICHIOMETRY AND EFFICIENCY OF CO2 FIXATION COUPLED TO REDUCED SULFUR OXIDATION Klatt, Judith M., Polerecky, Lubos Supplementary Information Content 1. Implementation of the efficiency calculation in R 2. Calculation of the stoichiometry of autotrophic sulfur oxidation – other examples 3. Net and gross fluxes of O2 and CO2 in Riftia pachyptila tubeworms 4. Details of efficiency calculations for Sulfurimonas denitrificans and Thioalkalivibrio versutus 5. Supplementary Figures 5.1. Figure S1: Schematic representation of the biochemical transformations involved in energy conservation by SOB. 5.2. Figure S2: Sensitivity analysis of the calculated energy conservation efficiency of sulfur oxidation coupled to CO2 fixation. 6. Supplementary Table S1 6.1. Table S1A: Generalized mass-balanced equations for aerobic oxidation of reduced sulfur species coupled to CO2 fixation. 6.2. Table S1B: Generalized mass-balanced equations for anaerobic oxidation of reduced sulfur species coupled to DNRA and CO2 fixation 6.3. Table S1C: Generalized mass-balanced equations for anaerobic oxidation of reduced sulfur species coupled to denitrification and CO2 fixation 6.4. Table S1D: Generalized mass-balanced equations for anaerobic oxidation of reduced sulfur species coupled to nitrite reduction and CO2 fixation. 6.5. Table S1E: Generalized mass-balanced equation for anaerobic oxidation of reduced sulfur species coupled to N2O reduction and CO2 fixation. 7. Supplementary Table S2: Stoichiometries of autotrophic sulfur oxidation calculated for specific SOB strains based on experimental data available in the literature. 8. Supplementary References 126 II. Manuscripts 1. Implementation of the efficiency calculation in R R-scripts for the calculation of the energy conservation efficiency of autotrophic sulfur oxidation processes can be downloaded from http://nanosims.geo.uu.nl/SOX. The required functions are loaded in R using command source('SOX_functions.R') To calculate the efficiency for a given stoichiometry, the values of x and y need to be provided as input parameters. In the following we assume that these values are equal to x = 0 and y = 0.825. The efficiency of aerobic sulfide oxidation at standard biochemical conditions is calculated with the traditional approach using command aerob_efficiency(x=0,y=0.825,Sred="H2S",approach=1)$eff To calculate the efficiency using our new approach one needs to additionally specify the carbon fixation pathway, e.g., aerob_efficiency(x=0,y=0.825,Sred="H2S",approach=2,Cfix="Calvin")$ef f To calculate the efficiency of aerobic thiosulfate oxidation, one needs to specify thiosulfate as the reduced sulfur compound, i.e., aerob_efficiency(x=0,y=0.825,Sred="S2O3",approach=2,Cfix="Calvin")$e ff By default, the efficiency is calculated at standard biochemical conditions. To calculate it at different conditions, one needs to specify the concentrations of the reactants involved. For 2- example, for the concentrations of total sulfide [H2S] = 30 μM, [SO4 ] = 20 mM, + [O2] = 100 μM, total inorganic carbon [CO2] = 2.5 mM, [NAD ] = [NADH] = 1 mM, pH of 6 and temperature of 15 °C, the efficiency is calculated using command aerob_efficiency(x=0,y=0.825,Sred="H2S",approach=2,Cfix="Calvin",S=3 0e-6, sulfate=20e-3,O2=100e-6,CO2=2.5e-3,NAD=1e-3,NADH=1e-3,pH=6, Temp=15)$eff The efficiency of anaerobic sulfur oxidation coupled to denitrification is calculated using the function denit_efficiency. The input parameters are the same as for the aerob_efficiency function. For example, to calculate the efficiency for an SOB that 127 3. STOICHIOMETRY AND EFFICIENCY OF SULFUR OXIDATION performs thiosulfate oxidation coupled to denitrification and employs the rTCA cycle as the CO2 fixation pathway using our new approach, one needs to use command denit_efficiency(x=0,y=0.825,Sred="S2O3",approach=2,Cfix="rTCA")$eff By default, the efficiency is calculated at standard biochemical conditions. To calculate the efficiency at different conditions, one needs to specify the concentrations of the reactants 2- 2- involved. For example, for the concentrations [S2O3 ] = 30 μM, [SO4 ] = 20 mM, - + [NO3 ] = 100 μM, total inorganic carbon [CO2] = 2.5 mM, [N2] = 10 mM, [NAD ] = [NADH] = [Fdox] = [Fdred] = [FAD] = [FADH2] = 1 mM, pH of 6 and temperature of 15 °C, the efficiency is calculated using command denit_efficiency(x=0,y=0.825,Sred="S2O3",approach=2,Cfix="rTCA",S=30 e-6, sulfate=20e-3,NO3=100e-6,N2=10e-3,CO2=2.5e-3,NAD=1e-3,NADH=1e- 3,Fdox=1e-3, Fdred=1e-3,FAD=1e-3,FADH2=1e-3,pH=6, Temp=15)$eff If $eff in the above function calls is omitted, the function call will additionally display the values of the Gibbs free energy for the energy generating and energy conserving reactions and of the quotients Q. The values of the partial efficiencies (İSO,II,min,, İCO2 and İRET,II,min) are displayed only for our new calculation approach (i.e., when approach=2) The website also contains additional examples illustrating (i) the calculation of sulfur oxidation stoichiometry from a given efficiency and the ratio between the reduced sulfur compound and the terminal electron acceptor (TEA) (file stoichiometry_examples.R), (ii) the sensitivity analysis of the efficiency calculation (file sensitivity_analysis.R; examples shown in Fig. S2), and (iii) the dependence of the CO2:TEA ratio on the parameter x (file CO2_TEA_x.R; examples shown in Fig. 3). 2. Calculation of the stoichiometry of autotrophic sulfur oxidation – other examples 2.1. Beggiatoa strain MS-81-1c Hagen & Nelson (1997) found that CO2 fixation in the Beggiatoa strain MS-81-1c occurred at a CO2:O2 consumption ratio of 0.596 and was accompanied by a considerable production 0 0 of S (S :O2 ratio of 0.04). According to Table S1A, this can be transformed into equations ȞCO2/ȞO2 = (1–y)/y = 0.596 and ȞS0/ȞO2 = x/[y(2-1.5x)] = 0.04, which yield y = 0.64 and x = 0.05. Thus, in this strain, aerobic oxidation of sulfide is performed according to equation 2- 0 + H2S + 1.23 O2 + 0.69 CO2 + 0.64 H2O Æ 0.95 SO4 + 0.05 S + 0.69 CH2O + 1.9 H 128 II. Manuscripts with 64% of the sulfide pool used in the energy gaining reaction with oxygen, 5% of the sulfide pool converted to zero-valent sulfur and the remaining 95% completely oxidized to 2- SO4 . 2.2. Thioploca bundles Otte et al. (1999) found that intact bundles of Thioploca sp. fixed CO2 in the absence of externally supplied sulfide by oxidizing their intra-cellularly stored zero-valent sulfur to sulfate accompanied by nitrate reduction to ammonium. When corrected for the rates in + 2- disrupted control samples, the production of NH4 and SO4 occurred at a ratio of ȞNH4/ȞSO4 = 0.5. This ratio is equal to 0.75y, which yields y = 0.67. Thus, zero-valent sulfur oxidation in the Thioploca bundles coupled to ammonification and CO2 fixation is expected to occur according to equation 0 - 2- + + S + 0.5 NO3 + 0.5 CO2 + 2 H2O Æ SO4 + 0.5 NH4 + 0.5 CH2O + 1 H , with 67% of the zero-valent sulfur pool used in the energy gaining reaction with nitrate. 0 - Based on this stoichiometry, a growth yield of 0.5 mole of CO2 per mole of S and a NO3 :CO2 consumption ratio of 1 are expected. However, in another experiment with Thioploca - bundles Otte et al. (1999) measured a NO3 :CO2 consumption ratio of about 0.55. If this ratio is used as a starting value, Table S1B implies that ȞNO3/ȞCO2 = 0.75y/[1.5(1–y)] = 0.55, which yields y = 0.8 and thus a considerably different stoichiometry of the sulfur oxidation reaction: 0 - 2- + + S + 0.6 NO3 + 0.3 CO2 + 1.9 H2O Æ SO4 + 0.6 NH4 + 0.3 CH2O + 0.8 H . This apparent inconsistency suggests that during the different experiments reported by Otte et al. (1999) the studied Thioploca bundles (i) had a different physiological status, (ii) that an unidentified additional electron donor or acceptor was involved, or (iii) that significant sulfur cycling within the bundles occurred. The latter possibility would be consistent with the finding of filamentous sulfate reducers co-inhabiting the Thioploca sheaths (Fukui et al., 1999; Teske et al., 2009). 129 3. STOICHIOMETRY AND EFFICIENCY OF SULFUR OXIDATION 3. Net and gross fluxes of O2 and CO2 in Riftia pachyptila tubeworms Girguis et al. (2002) performed several high-pressure respirometry experiments to determine net fluxes of sulfide, oxygen and CO2 for the host-symbiont system of tubeworms Riftia pachyptila. Specifically, for the untreated tubeworms they observed a net O2 consumption -1 -1 rate of ~10 μmol O2 g h , while after the addition of amiloride the O2 consumption rate -1 -1 decreased to ~2.5 μmol O2 g h (both values estimated from Figure 5 in Girguis et al., 2002). Additionally, the treatment with amiloride resulted in a net production of CO2, suggesting that the symbiont cells became inactive. Assuming that the amiloride treatment inhibited completely the activity of the symbionts but had no effect on the host, these values suggest that host respiration contributes an estimated 25% to the net O2 consumption of the whole-worm symbiosis while the symbionts-mediated autotrophic sulfide oxidation activity contributes the remaining 75%. To calculate the gross rate of CO2 fixation by the symbionts from the net rate of CO2 consumption by the whole-worm symbiosis, it is necessary to know the rate of CO2 production due to the respiration by the host. We estimated this rate by assuming that the host consumes O2 and produces CO2 at a ratio of 1:1. Thus, using the values reported by Girguis et al. (2002) for an experiment where they simultaneously measured the net rates of O2 -1 -1 -1 -1 consumption (12.4 μmol O2 g h ), CO2 consumption (12.45 μmol CO2 g h ) and sulfide -1 -1 consumption (6.75 μmol H2S g h ) for the whole-worm symbiosis, we estimated the gross -1 -1 O2 consumption rate by the symbionts to be 0.75 × 12.4 = 9.3 μmol O2 g h and the -1 -1 corresponding gross rate of CO2 fixation to be 12.45 + 0.25 × 12.4 = 15.55 μmol CO2 g h . 4. Details of efficiency calculations for Sulfurimonas denitrificans and Thioalkalivibrio versutus Hoor (1981) reported that Sulfurimonas denitrificans performs aerobic thiosulfate oxidation coupled to CO2 fixation with a growth yield of 0.333 mole of carbon per mole of thiosulfate, with thiosulfate completely oxidised to sulfate. Using Table S1A, this information leads to equations ȞCO2/ȞS2O3 = (1–y) (2–1.5x) = 0.333 and x = 0, which yield y = 0.833 and the corresponding equation 130 II. Manuscripts 2- 2- + S2O3 + 1.67 O2 + 0.33 CO2 + 1.33 H2OÆ2 SO4 + 0.33 CH2O + 2 H . 0 Using these stoichiometric coefficients and the ǻGf values tabulated by Thauer (1977), and 0 considering that Sulfurimonas denitrificans employs the rTCA cycle, we obtained ǻGr (O2 -1 0 -1 0 0 red) = -842.29 kJ mol , ǻGr (CO2 red) = 131.77 kJ mol , [ǻGr (CO2 fix) + ǻGr (RET)] = -1 0 -1 335.52 kJ mol and [ǻEr(CO2 fix) + ǻGr (RET)] = 343.52 kJ mol . Assuming standard biochemical conditions, substitution of these values together with the value of y to Eqs. 8, 30 and 37 gave the efficiencies İI = 0.0314, İII = 0.0799 and İSO,II,min = 0.0818. Sorokin et al. (2001) reported that Thioalkalivibrio versutus performs aerobic thiosulfate oxidation coupled to CO2 fixation with a growth yield of 5 g protein per mole of thiosulfate, with thiosulfate completely oxidised to sulfate. Assuming a carbon:protein ratio of 0.75 (Hoor, 1981; Simon and Azam, 1989), this leads to equations ȞCO2/ȞS2O3 = (1–y) (2–1.5x) = 0.3125 and x=0, which yield y=0.859 and the corresponding equation 2- 2- + S2O3 + 1.69 O2 + 0.31 CO2 + 1.31 H2OÆ 2 SO4 + 0.31 CH2O + 2 H . 0 Using these stoichiometric coefficients and the ǻGf values tabulated by Thauer (1977), and 0 considering that Thioalkalivibrio versutus employs the Calvin cycle, we obtained ǻGr (O2 -1 0 -1 0 0 red) = -842.29 kJ mol , ǻGr (CO2 red) = 131.77 kJ mol , [ǻGr (CO2 fix) + ǻGr (RET)] = -1 0 -1 362.70 kJ mol and [ǻEr(CO2 fix) + ǻGr (RET)] = 469.36 kJ mol . At standard biochemical conditions this yields the efficiency values of İI = 0.0289, İII = 0.0796 and İSO,II,min = 0.1030. 131 3. STOICHIOMETRY AND EFFICIENCY OF SULFUR OXIDATION 5. Supplementary Figures A Sulfuroxidation Exergonic reaction: y̩Gr (TEAred) İSO ̩Er(SO):EnergygainedbySO red)] red) t 2 ATP ATP ATP ATP ATP ATP ATP ATP ATP İ u İ (CO (TEA r SO G İ ̩E (CO red) :Energyutilized G r 2 ̩ = [ İ ̩ I t forCO reduction ) y 2 İ y Ͳ ATP ATP ATP ATP ATP ATP ATP ATP ATP – (1 = I “maintenance” İ İu Endergonic reaction: CO2 reduction (1Ͳy) ̩Gr (CO2 red) B Sulfuroxidation Exergonic reaction: y̩Gr (TEAred) 1 Ͳ ] İSO ̩Er(SO):EnergygainedbySO CO2 İ ) (RET)] Į ATP ATP ATP ATP ATP ATP ATP ATP ATP Ͳ G (1 ȴ red) + + İt RET İ fix) (TEA 2 r Į [ ATP ATP ATP ATP ATP ATP ATP ATP ATP G t ̩ (CO İ G RET –y ̩ “maintenance” ̩E (CO fix):Energyutilized ̩E (RET):Energyutilized ɸ r 2 r [ Ͳ forCO fixation forreversee transport ) 2 y CO2 Ͳ İ ATP ATP ATP ATP ATP ATP ATP ATP (1 SO = İ II = İ II İ İ CO2Ͳfix İRET Endergonic reaction: CO fixation ReverseeͲ transport 2 (1Ͳy)(̩G (CO fix)+ȴG(RET)) r 2 Figure S1: Schematic representation of the biochemical transformations involved in energy conservation by SOB. Panels A and B correspond to the traditional and our new approach for calculating the energy conservation efficiency, respectively. In both approaches energy available from sulfur oxidation (SO) with the terminal electron acceptor (TEA) is transformed into the biochemical currency ATP with an efficiency İSO. A fraction İt of this ATP is utilized for CO2 reduction, while the remaining fraction (1 – İt) is used for cellular maintenance reactions. In the traditional approach the ATP is utilized for CO2 reduction with an efficiency İu. In contrast, our new approach considers that ATP is actually driving two processes: reverse electron transport to reduce electron carriers (e.g., + NAD , FAD, Ferredoxin), and the CO2 reduction reactions in the respective CO2 fixation pathway. The corresponding efficiencies of these processes are İRET and İCO2. ǻEr and ǻGr correspond to the energy of the reaction in the form of ATP and the Gibbs free energy of the reaction, respectively. 132 II. Manuscripts SO 2í 1 M, pH 7 4 A H2S 100 mM O2 T. tepidarius 10 mM CO2 1 mM pH pH 2 100 μM NAD+ HSO,II,min 10 μM NADH H. neapolitanus 1 μM H 100 nM 10 nM HSO,II,min 1 nM T. tepidarius pH 12 HI 0.05 0.10 0.15 0.20 HI H. neapolitanus 1eí34 1eí26 1eí18 1eí10 1eí02 Q SO 2í 4 B H2S í NO3 S. hongkongienses CO2 pH Fdox HSO,II,min Fdred pH 2 H S. denitrificans HSO,II,min HI pH 12 HI S. denitrificans 0.00 0.02 0.04 0.06 0.08 0.10 0.12 1eí28 1eí20 1eí12 1eí04 1e+04 Q Figure S2: Sensitivity analysis of the calculated energy conservation efficiency of sulfur oxidation coupled to CO2 fixation. Shown are values of İI and İSO,II,min calculated for (A) aerobic sulfide oxidizing bacteria Thermithiobacillus tepidarius and Halothiobacillus neapolitanus and for (B) denitrifying thiosulfate oxidizing bacteria Sulfurimonas hongkongienses and Sulfurimonas denitrificans.. In all calculations we separately varied the concentrations of each reactant from 1M to 1 nM, or the pH from 2 to 12, while keeping the other reactant concentrations constant at 1 M or pH 7. For the denitrifying SOB, which employ the rTCA cycle for CO2 fixation, the slope of the variation + of İSO,II,min with the concentration of the oxidized/reduced electron carriers NAD /NADH and FAD/FADH2 is the same as that for the oxidized/reduced Ferredoxin (Fdox/Fdred). The sensitivity of + İSO,II,min to changes in the concentrations of NAD /NADH and FAD/FADH2 is lower than the + sensitivity to changes in the concentrations of Fdox/Fdred. Data-points corresponding to NAD /NADH and FAD/FADH2 are, however, not displayed to maintain graph clarity. In all graphs the large gray- filled squares represent the efficiency calculated at standard biochemical conditions (1M concentrations of all reactants, pH=7, temperature 25 °C). 133 134 Table S1A: Generalized mass-balanced equations for aerobic oxidation of reduced sulfur species coupled to CO2 fixation. 2- 5H2S oxidation S2O3 oxidation 0 2- 0 2- Energy-generating reactions H2S + 0.5 O2 b S + H2O S2O3 + 0.5 O2 b S + SO4 2- + 2- 2- + H2S + 2 O2 b SO4 + 2 H S2O3 + 2 O2 + H2O b 2 SO4 + 2 H 0 2- + 2- 0 2- + H2S + (2–1.5x) O2 b x S + (1–x) SO4 + x H2O + (2–2x)H S2O3 + (2–1.5x) O2 + (1–x) H2O bx S + (2–x) SO4 + (2–2x)H 0 2- 0 2- Energy-conserving reactions H2S + 0.5 CO2 b S + 0.5 CH2O + 0.5 H2O S2O3 + 0.5 CO2 + 0.5 H2O §b S + SO4 + 0.5 CH2O (traditional approach) 2- + 2- 2- + H2S + 2 CO2 + 2 H2O b SO4 + 2 CH2O + 2 H S2O3 + 2 CO2 + 3 H2O b 2 SO4 + 2 CH2O + 2 H 0 2- + 2- 0 2- + H2S + (2–1.5x) CO2 + (2–2.5x) H2O bx S + (1–x) SO4 + (2–1.5x) CH2O + (2–2x) H S2O3 + (2–1.5x) CO2 + (3–2.5x) H2O bx S + (2–x) SO4 + (2–1.5x) CH2O + (2–2x) H + 0 + 2- + + 0 2- Energy-conserving reactions H2S + NAD b S + NADH + H S2O3 + NAD + H2O + 2 H b S + SO4 + 4 NADH (Calvin cycle, new approach)1 + 2- + 2- + 2- + H2S + 4 NAD + 4 H2O b SO4 + 4 NADH + 6 H S2O3 + 4 NAD + 5 H2O b 2 SO4 + 4 NADH + 6 H + 0 2- + 2- + 0 2- + H2S + (4–3x) NAD + (4–4x) H2O bx S + (1–x) SO4 + (4–3x) NADH + (6–5x) H S2O3 + (4–3x) NAD + (5–4x) H2O bx S + (2–x) SO4 + (4–3x) NADH + (6–8x) H + 2- + Energy-conserving reactions H2S + 0.5 NAD + 0.67 Fdox + 0.17 FAD b S2O3 + 0.5 NAD + 0.67 Fdox + 0.17 FAD + H2O b 1 0 + 0 2- + (rTCA cycle, new approach) S + 0.5 NADH + 0.67 Fdred + 0.17 FADH2 + 1.17 H S + SO4 + 0.5 NADH + 0.67 Fdred + 0.17 FADH2 +1.17 H + 2- + H2S + 2 NAD + 2.67 Fdox + 0.67 FAD + 4 H2O b S2O3 + 2 NAD + 2.67 Fdox + 0.67 FAD + 5 H2O b 2- + 2- + SO4 + 2 NADH + 2.67 Fdred + 0.67 FADH2 + 6.67 H 2 SO4 + 2 NADH + 2.67 Fdred + 0.67 FADH2 + 6.67 H + 2- + + H2S + (2 – 1.5x) NAD + (2.67 – 2x) Fdox + (0.67 – 0.5x) FAD + (4 – 4x) H2O b S2O3 + (2 – 1.5x) NAD + (2.67 – 2x) Fdox + (0.67 – 0.5x) FAD + (5 – 4x) H2O b 0 2- + 0 2- + x S + (1 – x) SO4 + (2 – 1.5x) NADH + (2.67 – 2x) Fdred + (0.67 – 0.5x) FADH2 + (6.67 – 5.5x) H x S + (1 – x) SO4 + (2 – 1.5x) NADH + (2.67 – 2x) Fdred + (0.67 – 0.5x) FADH + (6.67 – 5.5x) H 0 2- + 2- 0 2- + Generalized coupled reaction H2S + *O2 O2 + *CO2 CO2 + *H2O H2O b*S0 S + *SO4 SO4 + *orgC CH2O + *H+ H S2O3 + *O2 O2 + *CO2 CO2 + *H2O H2O b*S0 S + *SO4 SO4 + *orgC CH2O + *H+ H *O2 = y (2–1.5x) *O2 = y (2–1.5x) *CO2 = (1–y) (2–1.5x) *CO2 = (1–y) (2–1.5x) *H2O = (1–y) (2–2.5x) – yx *H2O =y (1–x) + (1–y) (3–2.5x) *S0 = x *S0 = x *SO4 = (1–x) *SO4 = (2–x) *orgC = (1–y) (2–1.5x) *orgC = (1–y) (2–1.5x) *H+ = (2–2x) *H+ = (2–2x) 1 Equation for CO2 fixation coupled to electron carrier oxidation (Eqs. 18 and 24) is the same for all sulfur oxidation processes8 Table S1B: Generalized mass-balanced equations for anaerobic oxidation of reduced sulfur species coupled to DNRA and CO2 fixation. 2- 5H2S oxidation S2O3 oxidation 1 - + 0 + 2- - + 0 2- + Energy-generating reactions H2S + 0.25 NO3 + 0.5 H b S + 0.25 NH4 + 0.75 H2O S2O3 + 0.25 NO3 + 1.5 H b S + SO4 + 0.25 NH4 + 0.25 H2O - 2- + 2- - 2- + H2S + NO3 + H2O b SO4 + NH4 S2O3 + NO3 + 2 H2O b 2 SO4 + NH4 - + 0 2- + 2- - + 0 2- + H2S + (1–0.75x) NO3 + (1–1.75x) H2O + 0.5x H b x S + (1–x) SO4 + (1–0.75x) NH4 S2O3 + (1–0.75x) NO3 + (2–2.25 x) H2O + 1.5x H bx S + (2–x) SO4 + (1–0.75x) NH4 - 0 2- + + 2- - 0 2- + + Generalized coupled reaction H2S + *NO3 NO3 + *CO2 CO2 + *H2O H2O b *S0 S + *SO4 SO4 + *NH4+NH4 + *orgC CH2O + *H+ H S2O3 + *NO3 NO3 + *CO2 CO2 + *H2O H2O b *S0 S + *SO4 SO4 + *NH4+ NH4 + *orgC CH2O + *H+ H *NO3 = y (1–0.75x) *NO3 = y (1–0.75x) *CO2 = (1–y) (2–1.5x) *CO2 = (1–y) (2–1.5x) *H2O =y (1–1.75x) + (1–y) (2–2.5x) *H2O =y (2–2.25x) + (1–y) (3–2.5x) *S0 = x *S0 = x *SO4 = (1–x) *SO4 =(2–x) *NH4 = y (1–0.75x) *NH4 = y (1–0.75x) *orgC = (1–y) (2–1.5x) *orgC = (1–y) (2–1.5x) *H+ = (1–y) (2–2x) – y 0.5x *H+ = (1–y) (2–2x) – y 1.5x 1 The generalized energy conserving reactions are the same as in Table S1A. 135 136 Table S1C: Generalized mass-balanced equations for anaerobic oxidation of reduced sulfur species coupled to denitrification and CO2 fixation. 2- 5H2S oxidation S2O3 oxidation 1 - 0 + 2- - + 0 2- Energy-generating reactions H2S + 0.4 NO3 b S + 0.2 N2 + 1.2 H2O + 0.6 H S2O3 + 0.4 NO3 + 0.4 H b S + SO4 + 0.2 N2 + 0.2 H2O - 2- + 2- - 2- + H2S + 1.6 NO3 b SO4 + 0.8 N2 + 0.8 H2O + 0.4 H S2O3 + 1.6 NO3 + 0.2 H2O b 2 SO4 + 0.8 N2 + 0.4 H - 0 2- + 2- - 0 2- + H2S + (1.6–1.2x) NO3 + (- 0.8– 0.4x) H2O bx S + (1–x) SO4 + (0.8–0.6x) N2 + (0.4 + 0.2x)H S2O3 + (1.6–1.2x) NO3 + (0.2– 0.4x) H2O bx S + (2–x) SO4 + (0.8–0.6x) N2 + (0.4–0.8x)H - 0 2- + 2- - 0 2- + Generalized coupled reaction H2S + *NO3 NO3 + *CO2 CO2 + *H2O H2O b*S0 S + *SO4 SO4 + *N2 N2 + *orgC CH2O + *H+ H S2O3 + *NO3 NO3 + *CO2 CO2 + *H2O H2O b*S0 S + *SO4 SO4 + *N2 N2 + *orgC CH2O + *H+ H *NO3 = y (1.6–1.2x) *NO3 = y (1.6–1.2x) *CO2 = (1–y) (2–1.5x) *CO2 = (1–y) (2–1.5x) *H2O =y (–0.8–0.4x) + (1–y) (2–2.5x) *H2O =y (0.2– 0.4x)+ (1–y) (3–2.5x) *S0 = x *S0 = x *SO4 =(1–x) *SO4 = (2–x) *N2 = y (0.8–0.6x) *N2 = y (0.8–0.6x) *orgC = (1–y) (2–1.5x) *orgC = (1–y) (2–1.5x) *H+ = y (0.4+0.2x) + (1–y) (2–2x) *H+ = y (0.4–0.8x)+ (1–y) (2–2x) 1 The generalized energy conserving reactions are the same as in Table S1A. Table S1D: Generalized mass-balanced equations for anaerobic oxidation of reduced sulfur species coupled to nitrite reduction and CO2 fixation. 2- 5H2S oxidation S2O3 oxidation 1 - + 0 2- - + 0 2- Energy-generating reactions H2S + 0.67 NO2 + 0.67 H b S + 0.33 N2 + 1.33 H2O S2O3 + 0.67 NO2 + 0.67 H b S + SO4 + 0.33 N2 + 0.33 H2O - + 2- 2- - + 2- H2S + 2.67 NO2 + 0.67 H b SO4 + 1.33 N2 + 1.33 H2O S2O3 + 2.67 NO2 + 0.67 H b 2 SO4 + 1.33 N2 + 0.33 H2O - + 0 2- 2- - 0 2- + H2S + (2.67–2x) NO2 + 0.67 H bx S + (1–x) SO4 + (1.33–1x) N2 + 1.33 H2O S2O3 + (0.67 + 2x) NO2 + (–0.33) H2O bx S + (2–x) SO4 + (0.33–x) N2 + (–0.67) H - 0 2- + 2- - 0 2- + Generalized coupled reaction H2S + *NO2 NO2 + *CO2 CO2 + *H2O H2O b*S0 S + *SO4 SO4 + *N2 N2 + *orgC CH2O + *H+ H S2O3 + *NO2 NO2 + *CO2 CO2 + *H2O H2O b*S0 S + *SO4 SO4 + *N2 N2 + *orgC CH2O + *H+ H *NO2 = y (2.67–2x) *NO2 = y (0.67+2x) *CO2 = (1–y) (2–1.5x) *CO2 = (1–y) (2–1.5x) *H2O = (1–y) (2–2.5x) – 1.33 y *H2O = –0.33 y + (1–y) (3–2.5x) *S0 = x *S0 = x *SO4 =(1–x) *SO4 =(1–x) *N2 = y (1.33–1x) *N2 = y (0.33–x) *orgC = (1–y) (2–1.5x) *orgC = (1–y) (2–1.5x) *H+ = (1–y) (2–2x) – 0.67 y *H+ = –0.67y + (1–y) (2–2x) 1 The generalized energy conserving reactions are the same as in Table S1A. 137 138 Table S1E: Generalized mass-balanced equations for anaerobic oxidation of reduced sulfur species coupled to N2O reduction and CO2 fixation. 2- 5H2S oxidation S2O3 oxidation 1 0 2- 0 2- Energy-generating reactions H2S + N2O b S + N2 + H2O S2O3 + N2O b S + SO4 + N2 2- + 2- 2- + H2S + 4 N2O b SO4 + 4 N2 + 2 H S2O3 + 4 N2O + H2O b 2 SO4 + 4 N2 + 2 H 0 2- + 2- 0 2- + H2S + (4–3x) N2O bx S + (1–x) SO4 + (4–3x) N2 + x H2O + (2–2 x) H S2O3 + (4–3x) N2O + (1–x) H2O bx S + (2–x) SO4 + (4–3x) N2 + (2–2x) H 0 2- + 2- 0 2- + Generalized coupled reaction H2S + *N2O N2O + *CO2 CO2 + *H2O H2O b*S0 S + *SO4 SO4 + *N2 N2 + *orgC CH2O + *H+ H S2O3 + *N2O N2O + *CO2 CO2 + *H2O H2O b*S0 S + *SO4 SO4 + *N2 N2 + *orgC CH2O + *H+ H *N2O = y (4–3x) *N2O = y (4–3x) *CO2 = (1–y) (2–1.5x) *CO2 = (1–y) (2–1.5x) *H2O = (1–y) (2–2.5x) – yx *H2O =y (1–x) + (1–y) (3–2.5x) *S0 = x *S0 = x *SO4 =(1–x) *SO4 =(2–x) *N2 = y (4–3x) *N2 = y (3–2x) *orgC = (1–y) (2–1.5x) *orgC = (1–y) (2–1.5x) *H+ = (2–2x) *H+ = (2–2x) 1The generalized energy conserving reactions are the same as in Table S1A. Table S2: Stoichiometries of autotrophic sulfur oxidation calculated for specific SOB strains based on experimental data available in the literature. x y Reference(s) PART A: Aerobic thiosulfate oxidation 2- 2- + Thiobacillus denitrificans S2O3 + 1.42 O2 + 0.58 CO2 + 1.58 H2O b 2 SO4 + 0.58 CH2O + 2 H 0 0.710 (Justin and Kelly, 1978) 2- 2- + Thiobacillus denitrificans S2O3 + 1.48 O2 + 0.52 CO2 + 1.52 H2O b 2 SO4 + 0.52 CH2O + 2 H 0 0.740 (Hoor, 1981) 2- 2- + Thiothrix CT3 S2O3 + 1.48 O2 + 0.52 CO2 + 1.52 H2O b 2 SO4 + 0.52 CH2O + 2 H 0 0.738 (Rossetti, 2003) 2- 2- + Beggiatoa str. D-402 S2O3 + 1.52 O2 + 0.48 CO2 + 1.48 H2O b 2 SO4 + 0.48 CH2O + 2 H 0 0.761 (Grabovich et al., 2001) 2- 2- + Thermothrix thiopara S2O3 + 1.57 O2 + 0.44 CO2 + 1.44 H2O b 2 SO4 + 0.44 CH2O + 2 H 0 0.783 (Mason et al., 1987) 2- 2- + Thermithiobacillus tepidarius S2O3 + 1.61 O2 + 0.39 CO2 + 1.39 H2O b 2 SO4 + 0.39 CH2O + 2 H 0 0.804 (Wood and Kelly, 1986) 2- 2- + Thioalkalivibrio versutus S2O3 + 1.69 O2 + 0.31 CO2 + 1.31 H2O b 2 SO4 + 0.31 CH2O + 2 H 0 0.844 (Sorokin et al., 2001) 2- 2- + Acidithiobacillus ferrooxidans S2O3 + 1.70 O2 + 0.30 CO2 + 1.30 H2O b 2 SO4 + 0.30 CH2O + 2 H 0 0.848 (Eccleston and Kelly, 1978) 2- 2- + Paracoccus versutus S2O3 + 1.71 O2 + 0.29 CO2 + 1.29 H2O b 2 SO4 + 0.29 CH2O + 2 H 0 0.853 (Mason et al., 1987) (Hempfling and Vishniac, 1967; Halothiobacillus neapolitanus S O 2- + 1.72 O + 0.28 CO + 1.28 H O b 2 SO 2- + 0.28 CH O + 2 H+ 0 0.859 2 3 2 2 2 4 2 Kelly, 1982) 2- 2- + Thiomicrospira thioparus S2O3 + 1.73 O2 + 0.27 CO2 + 1.27 H2O b 2 SO4 + 0.27 CH2O + 2 H 0 0.863 (Kelly, 1982) 2- 2- + Sulfurimonas denitrificans S2O3 + 1.67 O2 + 0.33 CO2 + 1.33 H2O b 2 SO4 + 0.33 CH2O + 2 H 0 0.833 (Hoor, 1981) 2- 2- + Acidithiobacillus thiooxidans S2O3 + 1.75 O2 + 0.25 CO2 + 1.25 H2O b 2 SO4 + 0.25 CH2O + 2 H 0 0.874 (Mason et al., 1987) 2- 2- + Halothiobacillus halophilus S2O3 + 1.75 O2 + 0.25 CO2 + 1.25 H2O b 2 SO4 + 0.25 CH2O + 2 H 0 0.875 (Wood and Kelly, 1991) 2- 2- + Thiothrix ramosa S2O3 + 1.75 O2 + 0.25 CO2 + 1.25 H2O b 2 SO4 + 0.25 CH2O + 2 H 0 0.875 (Odintsova et al., 1993) 2- 2- + Thioalkalispira microaerophila S2O3 + 1.77 O2 + 0.23 CO2 + 1.23 H2O b 2 SO4 + 0.23 CH2O + 2 H 0 0.884 (Sorokin, 2002) 2- 2- + Thioalkalimicrobium aerophilum S2O3 + 1.78 O2 + 0.22 CO2 + 1.22 H2O b 2 SO4 + 0.22 CH2O + 2 H 0 0.891 (Sorokin et al., 2001) 2- 2- + Thiomicrospira halophila S2O3 + 1.78 O2 + 0.22 CO2 + 1.22 H2O b 2 SO4 + 0.22 CH2O + 2 H 0 0.891 (Sorokin et al., 2006) 2- 2- + Thioalkalibacter halophilus S2O3 + 1.78 O2 + 0.22 CO2 + 1.22 H2O b 2 SO4 + 0.22 CH2O + 2 H 0 0.891 (Banciu et al., 2008) 2- 2- + Thiobacillus thioparus E6 S2O3 + 1.80 O2 + 0.20 CO2 + 1.20 H2O b 2 SO4 + 0.20 CH2O + 2 H 0 0.898 (Smith and Kelly, 1988) 2- 2- + Thioclava pacifica S2O3 + 1.83 O2 + 0.17 CO2 + 1.17 H2O b 2 SO4 + 0.17 CH2O + 2 H 0 0.913 (Sorokin et al., 2005) 2- 2- + Thioalkalimicrobium sibericum S2O3 + 1.83 O2 + 0.17 CO2 + 1.17 H2O b 2 SO4 + 0.17 CH2O + 2 H 0 0.916 (Sorokin et al., 2001) 139 2- 2- 0 + Thiomicrospira sp. Strain L-12 S2O3 + 1.57 O2 + 0.13 CO2 + 0.93 H2O b 1.8 SO4 + 0.2 S + 0.13 CH2O + 1.6 H 0.2 0.921 (Ruby and Jannasch, 1982) 140 PART B: Thiosulfate oxidation coupled to denitrification 2- - 2- + Thiobacillus denitrificans S2O3 + 1.24 NO3 + 0.45 CO2 + 0.83 H2O b 2 SO4 + 0.62 N2 + 0.45 CH2O + 0.76 H 0 0.775 (Justin and Kelly, 1978) 2- - 2- + Thiobacillus denitrificans S2O3 + 1.30 NO3 + 0.38 CO2 + 0.73 H2O b 2 SO4 + 0.65 N2 + 0.38 CH2O + 0.70 H 0 0.812 (Hoor, 1981) 2- - 2- + Sulfurimonas hongkongensis S2O3 + 1.36 NO3 + 0.30 CO2 + 0.61 H2O b 2 SO4 + 0.68 N2 + 0.30 CH2O + 0.64 H 0 0.852 (Cai et al., 2014) 2- - 2- + Sulfurimonas gotlandica S2O3 + 1.40 NO3 + 0.25 CO2 + 0.55 H2O b 2 SO4 + 0.70 N2 + 0.25 CH2O + 0.60 H 0 0.875 (Bruckner et al., 2013) Sulfurimonas gotlandica-like 2- - 2- + Epsilonproteobacterium S2O3 + 1.42 NO3 + 0.23 CO2 + 0.52 H2O b 2 SO4 + 0.71 N2 + 0.23 CH2O + 0.58 H 0 0.887 (Brettar et al., 2006) (environmental sample) 2- - 2- + Sulfurimonas denitrificans S2O3 + 1.44 NO3 + 0.20 CO2 + 0.47 H2O b 2 SO4 + 0.72 N2 + 0.20 CH2O + 0.56 H 0 0.902 (Hoor, 1981) PART C: Aerobic sulfide oxidation 0 2- + Riftia pachyptila symbionts H2S + 1.12 S + 0.75 O2 + 2.3 CO2 + 3.42 H2O b 2.12 SO4 + 2.3 CH2O + 4.24 H -1.12 0.375 (Girguis et al., 2002) 2- 0 + Solemya reidi symbionts H2S + 0.75 O2 + 1 CO2 + 0.83 H2O b 0.83 SO4 + 0.17 S + 1 CH2O + 1.67 H 0.17 0.429 (Anderson et al., 1987) 2- 0 + Riftia pachyptila symbionts H2S + 0.75 O2 + 0.75 CO2 + 0.42 H2O b 0.67 SO4 + 0.33 S + 0.75 CH2O + 1.33 H 0.33 0.500 (Girguis and Childress, 2006) 2 0- + Beggiatoa str. MS-81-1c H2S + 1.23 O2 + 0.69 CO2 + 0.64 H2O b 0.95 SO4 + 0.05 S + 0.69 CH2O + 1.9 H 0.05 0.641 (Hagen and Nelson, 1997) 2- + Ridgeia piscesae symbionts H2S + 1.43 O2 + 0.57 CO2 + 0.57 H2O b SO4 + 0.57 CH2O + 2 H 0 0.714 (Nyholm et al., 2008) 2- + Thermithiobacillus tepidarius H2S + 1.62 O2 + 0.38 CO2 + 0.38 H2O bSO4 + 0.38 CH2O + 2 H 0 0.808 (Wood and Kelly, 1986) 2- + Beggiatoa str. MS-81-6 H2S + 1.65 O2 + 0.35 CO2 + 0.35 H2O b SO4 + 0.35 CH2O + 2 H 0 0.825 (Hagen and Nelson, 1997) 2- + Halothiobacillus neapolitanus H2S + 1.72 O2 + 0.28 CO2 + 0.28 H2O b SO4 + 0.28 CH2O + 2 H 0 0.859 (Kelly, 1982) II. Manuscripts 8. Supplementary References Anderson, A. E., Childress, J. J., and Favuzzi, J. A. (1987). Net uptake of CO2 Driven by Sulphide and Thiosulphate Oxidation in the Bacterial Symbiont-Containing Clam Solemya Reidi. J. Exp. Biol. 133, 1–31. Banciu, H. L., Sorokin, D. Y., Tourova, T. P., Galinski, E. A., Muntyan, M. S., Kuenen, J. G., and Muyzer, G. (2008). 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Chemolithotrophic metabolism of the newly-isolated moderately thermophilic, obligately autotrophic Thiobacillus tepidarius. Arch. Microbiol. 144, 71–77. doi:10.1007/BF00454959. Wood, A. P., and Kelly, D. P. (1991). Isolation and characterisation of Thiobacillus halophilus sp. nov., a sulphur-oxidising autotrophic eubacterium from a Western Australian hypersaline lake. Arch. Microbiol. 156, 277–280. doi:10.1007/BF00262998. 143 144 II. Manuscripts 4. H2SCAN ENHANCE AND INHIBIT OXYGENIC PHOTOSYNTHESIS IN A CYANOBACTERIUM FROM SULPHIDIC SPRINGS Klatt, Judith M.1*; Haas, Sebastian1; Yilmaz, Pelin1; de Beer, Dirk1; Polerecky, Lubos1,2 1Max Planck Institute for Marine Microbiology, Celsiusstr.1, 28359 Bremen, Germany 2Department of Earth Sciences – Geochemistry, Faculty of Geosciences, Utrecht University, Princetonplein 9, 3584 CC, Utrecht, The Netherlands Published in Environmental Microbiology (2015) *corresponding author: 145 4. REGULATION OF CYANOBACTERIAL PHOTOSYNTHESIS BY H2S Summary We used microsensors to investigate the combinatory effect of H2S and light on oxygenic photosynthesis in biofilms formed by a cyanobacterium from sulphidic springs. We found that photosynthesis was both positively and negatively affected by H2S. (i) H2S accelerated the recovery of photosynthesis after prolonged exposure to darkness and anoxia. We suggest that this is possibly due to regulatory effects of H2S on photosystem I components and/or on the Calvin cycle. (ii) H2S concentrations of up to 210 μM temporarily enhanced the photosynthetic rates at low irradiance. Modelling showed that this enhancement is plausibly based on changes in the light-harvesting efficiency. (iii) Above a certain light-dependent concentration threshold H2S also acted as an inhibitor. Intriguingly, this inhibition was not instant but occurred only after a specific time interval that decreased with increasing light intensity. That photosynthesis is most sensitive to inhibition at high light intensities suggests that H2S inactivates an intermediate of the oxygen evolving complex that accumulates with increasing light intensity. We discuss the implications of these three effects of H2S in the context of cyanobacterial photosynthesis under conditions with diurnally fluctuating light and H2S concentrations, such as those occurring in microbial mats and biofilms. Introduction Over the last centuries scientific interest in H2S was focused on its toxicity. This effect of H2S is mainly based on the inhibition of cytochrome c oxidase, the enzyme catalyzing the last step of oxidative phosphorylation in respiration (Beauchamp et al., 1984; Cooper and Brown, 2008). H2S can therefore negatively affect a multiplicity of organisms including bacteria, animals (Beauchamp et al., 1984; Dorman, 2002) and plants (Lamers et al., 2013). In the last two decades, however, rising attention has been given to H2S as a crucial metabolic regulator in higher organisms. In both animals and plants H2S serves as a gasotransmitter with diverse physiological functions (Hancock et al., 2011; Wang, 2012). In plants, externally supplied H2S was successfully tested as a growth-supporting agent, wherein the species-specific balance between the positive and toxic effects has to be considered (Lisjak et al., 2013). The specific positive effects of both endogenously produced and externally provided H2S include increased resistance against pathogens (Bloem et al., 2004; Rausch and Wachter, 2005), improved freezing and heat tolerance (Stuiver et al., 1992; Li et al., 2012), protection against heavy metal toxicity (Zhang et al., 2008; Zhang et al., 2010) and drought resistance (Jin et al., 2011). Besides increasing the 146 II. Manuscripts stress tolerance, H2S was also shown to be involved in the regulation of plant development (Rennenberg and Filner, 1983; Zhang et al., 2008) and other complex processes such as stomatal closure (García-Mata and Lamattina, 2010; Lisjak et al., 2010; Shen et al., 2013). In addition to these complex mechanisms, H2S also impacts the basic constituents of oxygenic photosynthesis in plants. For instance, H2S up-regulates the expression of photosynthetic genes such as those coding for RuBisCO and ferredoxin (Chen et al., 2011), promotes the activity of superoxide dismutase (Zhang et al., 2009), induces an increase in the chlorophyll content (Zhang et al., 2009; Chen et al., 2011) and possibly affects photosystem stoichiometry (PSI:PSII ratio) (Dooley et al., 2013). These components of photosynthesis had probably already evolved in the ancestor of all oxygenic phototrophs, most probably in a cyanobacterium (Mulkidjanian et al., 2006). This raises the question whether similar stimulatory functions of H2S on photosynthesis can also be found in these bacterial oxygenic phototrophs. This would indicate that the positive regulatory effects represent evolutionarily old regulatory mechanisms thereby setting the basis for the development of further functions of H2S in photosynthetic eukaryotes after the endosymbiotic event (McFadden, 2001). Cyanobacterial oxygenic photosynthesis evolved in microbial mat-like structures (stromatolites) about 3.8–2.4 Ga ago (Buick, 2008). Possibly, these systems were characterized by intense sulphur cycling and abundant H2S in the microenvironment of the early cyanobacteria (Nisbet and Fowler, 1999; Buick, 2008). Even after the first success of oxygenic photosynthesis in oxygenation of the atmosphere during the great oxygenation event, cyanobacteria still had to thrive in partially sulphidic oceans (Canfield, 1998; Johnston et al., 2009). Exposure of oxygenic phototrophs to H2S therefore runs like a thread through the history of their evolution. Still, the oxygen evolving complex (OEC) of all contemporary cyanobacteria studied so far is inhibited to a species-specific degree by H2S – even in species that are successful in sulphidic environments (Miller and Bebout, 2004). This suggests that the inhibitory effect of H2S is evolutionary deep-rooted and that plausibly already the first OEC was vulnerable to H2S toxicity. Consequently, cyanobacteria must have rapidly evolved strategies to cope with this toxicity and/or to even make use of H2S. One strategy might have been the active depletion of H2S by anoxygenic photosynthesis (Jørgensen et al., 1986). This capability to perform both oxygenic and anoxygenic photosynthesis is conserved in several specialized contemporary cyanobacterial species from sulphidic habitats (Cohen et al., 1975; Cohen 147 4. REGULATION OF CYANOBACTERIAL PHOTOSYNTHESIS BY H2S et al., 1986; Garcia-Pichel and Castenholz, 1990) (Klatt et al., 2015). The advantage of this strategy is the coupling of depletion of the toxic H2S to energy conservation. In contrast, the strategies of obligate oxygenic phototrophic cyanobacteria that cannot photosynthetically deplete H2S are comparatively poorly understood. So far, only Cohen et al. (1986) have shown for two cyanobacterial species that H2S can act as a stimulator of photosynthetic rates under specific light conditions and H2S concentrations. However, the parameter range, as well as the time frame over which this enhancement occurs, remain unclear. Typical contemporary habitats of sulphide-adapted cyanobacteria are microbial mats and biofilms (Cohen et al., 1986; Jørgensen et al., 1986; Voorhies et al., 2012) that are regularly used as analogues for stromatolites (Seckbach and Oren, 2010). Due to the activity of their inhabitants and mass transfer limitations, these systems are generally characterized by steep vertical gradients in the concentrations of sulphide and oxygen, pH and light (Seckbach and Oren, 2010). Moreover, due to the diurnally fluctuating input of light energy, the conditions in these systems are substantially changing also in time (Revsbech et al., 1983; Jørgensen et al., 1986). We hypothesized that all cyanobacteria that are competitive in such environments have special adaptations to H2S and light dynamics, even if they are not able to actively deplete sulphide by anoxygenic photosynthesis. We therefore aimed to identify and elucidate possible adaptation mechanisms of obligate oxygenic phototrophic cyanobacteria to the fluctuating exposure to H2S and light in such spatially heterogeneous and temporally dynamic systems. To fulfil this aim, we obtained a non-axenic mono-algal cyanobacterial culture from a thin microbial mat growing in a cold sulphidic spring and investigated its potential for anoxygenic photosynthesis and the inhibitory and stimulatory effects of H2S on its oxygenic photosynthesis. The measurements were performed in biofilms formed by the cyanobacterium using microsensors in a setup that allows creating artificial sulphide gradients across the biofilms (Fig. S1 in Supporting Information). We monitored the long- term effects of H2S on photosynthesis and simulated diurnal light and H2S fluctuations of natural microbial mats and biofilms. Results Studied strain The studied cyanobacterium strain FS34 originates from a thin microbial mat sampled in the Frasassi sulphidic springs (Galdenzi et al., 2008). The cyanobacterium is filamentous 148 II. Manuscripts (diameter of ~4 μm, length of up to ~100 μm; Fig. S2 in Supporting Information) and motile. The results of the tree reconstruction placed the sequence FS34 with other sequences of Planktothrix spp. (Fig. S2), and the sequence identity calculations returned values above 95%. Therefore, we propose that the isolate is a member of the genus Planktothrix and refer to it as Planktothrix str. FS34. Planktothrix str. FS34 does not perform sulphide-driven anoxygenic photosynthesis To determine whether Planktothrix str. FS34 can perform oxygenic and anoxygenic photosynthesis, the first experiments were conducted in a stirred batch reactor (Klatt et al., 2015). These experiments showed that Planktothrix str. FS34 can perform oxygenic photosynthesis in the presence of hydrogen sulphide (Fig. S3 in Supporting Information). However, as total sulphide was not consumed, we conclude that Planktothrix str. FS34 does not perform anoxygenic photosynthesis with sulphide as an electron donor within the studied range of irradiance (20–500 μmol photons m-2 s-1). During these batch reactor experiments we observed the formation of biofilms and aggregates. Thus, the bulk measurements did not accurately reflect the conditions that the cyanobacteria were actually exposed to in their microenvironment, because oxygen, pH and sulphide gradients typically develop within photosynthetically active biofilms (Wieland and Kühl, 2000). Although this does not invalidate the conclusion that they do not perform anoxygenic photosynthesis, further experiments were therefore performed in the biofilms with microsensors to accurately link the rates of photosynthesis to the local H2S concentration. Recovery of oxygenic photosynthesis after long-term exposure to darkness and anoxia To study the effects of H2S on the recovery of photosynthesis in Planktothrix str. FS34, we incubated biofilms of Planktothrix str. FS34 for ~10 h in the dark and anoxia and, after switching on the light, monitored the rates of gross photosynthesis (GP) together with the concentrations of O2 and H2S and pH inside the biofilm. After the first dark- anoxic incubation, which was done without H2S, GP increased slowly after the onset of illumination and the biofilm had to be illuminated for more than 6 h before GP reached an apparent steady state (Fig. 1A, triangles). After the next dark-anoxic incubation, which was done in a stable H2S gradient across the biofilm achieved by the addition of Na2S stock solution to the reservoir of the flow cell below the biofilm (Fig. S1), switching on the light led to a remarkably faster recovery of GP than in the previous H2S-free 149 4. REGULATION OF CYANOBACTERIAL PHOTOSYNTHESIS BY H2S incubation. The concentration of H2S gradually decreased within the biofilm because of the equilibrium shift in the sulphide pool due to an increase in pH (Fig. 1B–1D). When the H2S concentration in the biofilm was still between 10 μM (surface) and 20 μM (1 mm depth) the GP rates at all depths reached a transient maximum that was between 1.5- and 2-fold higher than the apparent steady state level reached after ~6 h of incubation (Fig. 1A, rectangles). Intriguingly, the acceleration of the GP recovery as well as the level of the transient GP maximum were higher in deeper layers where H2S concentrations were higher and light intensities plausibly lower (Fig. 1). When the same biofilm was again incubated in the absence of light, O2 and H2S, the recovery of GP after the light was switched on was again slow and gradual (data not shown). A long-term exposure of Planktothrix str. FS34 to H2S in the dark and anoxia has therefore two effects on its photosynthesis: it accelerates the rate of photosynthesis recovery and induces a transiently higher rate than in a steady state reached several hours after the light becomes available. Dynamics of the inhibition of photosynthesis by H2S To study the inhibitory effects of H2S on photosynthesis in Planktothrix str. FS34, we incubated biofilms for ~6 h at different levels of incident irradiance and H2S concentration and monitored O2, H2S, pH and the rate of GP in the uppermost layer of the biofilm. Before each measurement, we incubated the biofilm for 1-3 h in the dark under stable H2S and O2 gradients. In contrast to the results obtained after the long-term dark- anoxic incubations (see above), O2 concentrations in the biofilm increased immediately when the light was switched on after this short-term dark incubation (data not shown). Additionally, H2S concentrations concomitantly decreased due to a pH increase. When the H2S concentration in the dark had been adjusted to 230 μM, the subsequent exposure to a low light (irradiance of 41 μmol photons m-2 s-1) led to a rapid (within ~30 min) establishment of an apparent steady state (Fig. 2A). In this state, which we hereafter refer to as the “active state,” the cyanobacteria performed oxygenic photosynthesis at a constant rate while exposed to the H2S concentration of 180 μM (Fig. 2A). However, this active state was stable only for ~3 h, after which GP rapidly decreased to zero (Fig. 2A). The cyanobacteria remained in this inactive state as long as H2S was present. However, GP completely recovered within ~30 min after the H2S was removed from the biofilm (data not shown). The inhibitory effect of H2S was therefore reversible. In contrast to low light levels, a stable active state of photosynthesis in the presence of H2S was not observed at higher light levels. For example at irradiance of 130 μmol photons m-2 s-1 the 150 II. Manuscripts GP rate shortly increased after the onset of light, but then rapidly collapsed (Fig. 2B). At even higher light intensities, this peak of activity disappeared completely and the cells were photosynthetically inactive immediately after exposure to light (data not shown). Thus, oxygenic photosynthesis in Planktothrix str. FS34 exhibits a biphasic pattern: it is not inhibited during a short-term exposure to H2S, while a long term H2S exposure leads to an inhibition, which occurs faster at higher light intensities. dark light 125 200 ) -1 A s − H S ; depth 0. 1 mm 100 2 150 -1 − H S ; depth 0. 8 mm L 2 2 75 S (μM) + H S ; depth 0. 1 mm 100 2 2 + H S ; depth 0. 8 mm H 50 2 50 25 GP (nmol O (nmol GP 0 0 -10 -2 0 2 4 6 8 time (h) H S (μM) H S (μM) H S (μM) 2 2 2 0 20 40 60 80 100120140 0 20 40 60 80 100120140 0 20 40 60 80 100120140 S (mM) S (mM) S (mM) tot tot tot 0.0 0.5 1.0 1.5 2.0 2.5 3.0 0.0 0.5 1.0 1.5 2.0 2.5 3.0 0.0 0.5 1.0 1.5 2.0 2.5 3.0 −2 . 0 −2 . 0 −2 . 0 −1 . 5 B −1 . 5 C −1 . 5 D −1 . 0 −1 . 0 −1 . 0 −0 . 5 −0 . 5 −0 . 5 0.0 0.0 0.0 0.5 0.5 0.5 1.0 1.0 1.0 Depth (mm)Depth (mm)Depth (mm)Depth 1.5 1.5 1.5 2.0 2.0 2.0 2.5 2.5 2.5 0 10203040 0 10203040 0 10203040 O (μM) O (μM) O (μM) 2 2 2 78910 78910 78910 pH pH pH Figure 1: Recovery of oxygenic photosynthesis in the Planktothrix str. FS34 biofilm after the long-term exposure to darkness and anoxia in the presence (+H2S) and absence (–H2S) of H2S. (A): Volumetric rates of gross photosynthesis at two depths in the biofilm as a function of time, -2 -1 determined at incident irradiance of 69 μmol photons m s . The corresponding H2S concentration measured at the same depths (dashed line: 0.1 mm, solid line: 0.8 mm) are also shown. (B) – (D): Depth profiles of O2 and H2S concentrations and pH measured in the biofilm at time points 0 h, 3 h and 6 h during the exposure to H2S (+H2S-treatment). Stot concentrations were calculated from the local H2S concentrations and pH. 151 4. REGULATION OF CYANOBACTERIAL PHOTOSYNTHESIS BY H2S -2 dark 41 μmol photons m s-1 60 A 200 ) -1 40 150 s -1 100 L 20 2 50 0 0 -2 -1 dark 130 μmol photons m s S (μM) 2 60 B 200 H 40 150 GP (nmol O (nmol GP 100 20 50 0 0 0123456 time (h) Figure 2: Transition from the photosynthetically active state to an inactive state in Planktothrix str. FS34 biofilms. Gross photosynthesis rates (GP) were determined in the uppermost photosynthetically active layer of the biofilm at incident irradiances of 41 μmol photons m-2 s-1 -2 -1 (A) and 130 μmol photons m s (B). The corresponding H2S concentrations were measured in parallel in the same layer of the biofilm. Temporary enhancement of oxygenic photosynthesis by H2S at low light intensities To study the stimulating effects of H2S on photosynthesis in Planktothrix str. FS34, we measured the rates of GP in the uppermost layer of the biofilm as a function of irradiance (PI-curves) and H2S concentrations. Before each measurement the biofilm was incubated in the dark with the O2 concentration in the water column adjusted to ~20 % air saturation and the H2S concentration in the biofilm adjusted to a specific value by injecting the Na2S stock solution into the bottom part of the flow cell. After the establishment of a stable H2S gradient in the biofilm, the light was switched on, which led to the dynamic increase and decrease of GP described above (see Fig. 2). When all measured parameters (GP, H2S, pH) stabilized for more than 2 min, i.e., when a stable active state established, volumetric rates of GP and the H2S concentration were recorded. Afterwards, the light was switched off to allow for an increase in the H2S concentration in the measured point of the biofilm to the original value and then GP measurements were repeated at higher light intensities. Subsequently, the concentration of H2S was further increased by injection of Na2S stock into the bottom part of the flow cell and the entire procedure of GP measurements at different light intensities was repeated until partial PI-curves at various H2S concentrations were obtained. In the absence of H2S, the rate of GP exhibited a typical dependence on irradiance, increasing linearly at low light levels, reaching a maximum at some intermediate level and decreasing due to photoinhibition at high light levels (Fig. 3A). In contrast, the presence of sulphide induced complex patterns, wherein the interplay between both light intensity and H2S concentration determined the photosynthetic rates (Fig. 3B). At H2S 152 II. Manuscripts concentrations below 150 μM photosynthesis was enhanced at all light intensities and no inhibition was observed within the time frame of the measurement. Interestingly, the increase of the GP rate with irradiance was quadratic in the irradiance range of 0-40 μmol -2 -1 photons m s , as opposed to an approximately linear increase in the absence of H2S. -2 -1 Above 40 μmol photons m s the GP rates approached the values measured at zero H2S concentration and continued to increase with a slope comparable to the slope at zero H2S. However, there was a clear off-set towards higher rates. At H2S concentrations above 150 μM, the PI curve was characterized by a sharp decline above a certain threshold light intensity that decreased with increasing H2S concentrations (Fig. 3B). This decline corresponded to the rapid collapse (within <60 min) of the active state described above (see Fig. 2B). These clear patterns were only observed when plotted dependent on H2S concentrations but not as a function of Stot concentrations (data not shown). 0.7 40 H S (μM) AB2 0.6 35 0 ) -1 1 ) s -1 30 90 -1 0.5 s 150 L -1 2 25 170 L 0.4 2 180 20 200 230 μmol O 0.3 15 0.2 GP 10 GP (nmol O (nmol GP 0.1 5 0.0 0 0 100 200 300 400 0 50 100 150 Light intensity (μmol photons m-2 s-1) Light intensity (μmol photons m-2 s-1) Figure 3: Volumetric rates of gross photosynthesis as a function of incident irradiance (PI curves) measured in the uppermost layer of the Planktothrix str. FS34 biofilm. (A) Complete PI curve in the absence of H2S. The data points were fitted by the model of (Eilers and Peeters, 1988) (B) Partial PI curves measured at different H2S concentrations in another biofilm sample. Discussion Oxygenic photosynthesis is the light-driven process of H2O oxidation to O2 coupled to the reduction of CO2 to organic carbon. This coupling is facilitated by various redox reactions in the membrane (the electron transport chain) and in the cytosol during the Calvin cycle. Previous work has demonstrated that H2S can affect a variety of these intermediate steps in plants and some cyanobacteria. For instance, H2S inhibits the photosynthetic electron transport already during its first reaction, the H2O splitting in the OEC (Miller and Bebout, 2004). The activity of photosystem I (PSI), which couples electron flow from PSII via the cytochrome b6f complex to the last steps in the linear transport chain that serve NADP+ reduction, appears to be up-regulated (Dooley et al., 2013). Also H2S 153 4. REGULATION OF CYANOBACTERIAL PHOTOSYNTHESIS BY H2S increases the expression of the gene encoding for ferredoxin, one of the electron carriers between PSI and NADP+ (Chen et al., 2011). Even the activity of enzymes of the Calvin cycle is promoted by H2S (Chen et al., 2011). Also our study demonstrates that the effects of H2S can be multifaceted in a single organism. Specifically, H2S appears to have three main effects on oxygenic photosynthesis in Planktothrix str. FS34: (1) it can accelerate the recovery of photosynthesis after exposure to darkness and anoxia, (2) it can act as an inhibitor, and, before inhibition occurs, (3) it can temporarily enhance photosynthesis during a time interval whose duration decreases with light intensity. We propose that these effects are caused by separate mechanisms that are discussed in the following. Acceleration of the recovery of photosynthesis after exposure to darkness and anoxia We observed a very slow recovery of the photosynthetic activity in Planktothrix str. FS34 after extended exposure to anoxia and darkness. Intriguingly, the recovery time was substantially shortened by H2S (Fig. 1). A delay in the onset of photosynthesis after darkness and anoxia is a well documented phenomenon in oxygenic phototrophs (Schreiber et al., 2002) and is explained by the fact that the Calvin cycle cannot be instantaneously activated. After long periods of darkness ATP is depleted while NADPH is abundant. The latter cannot be oxidized to NADP+ due to the limitation in ATP. Also the electron transport chain cannot become oxidized due to the limitation in the NADP+ or other, external, electron acceptors such as O2 (McDonald et al., 2011). In other words, without a terminal electron acceptor the non-cyclic oxygenic photosynthetic electron transport cannot be active. Consequently, photosynthetic ATP generation, which is crucial to generate the electron acceptor NADP+ via the activity of RuBisCO, is inhibited. To overcome the problem of ATP limitation and over-reduction of the cellular components, oxygenic phototrophs generally use cyclic photophosphorylation in PSI (Schreiber et al., 2002). This process serves the generation of ATP in the absence of an electron acceptor for the non-cyclic oxygenic photosynthetic electron transport (Buchanan, 1991; Ghysels et al., 2013). H2S might therefore increase the rate of cyclic photophosphorylation in PSI. This could, for instance, be accomplished by increasing the number of PSI reaction centres or associated pigments, as seen in Zostera marina seedlings (Dooley et al., 2013). Alternatively, it could also be achieved through an increased activity of ferredoxin, which is also involved in the cyclic electron transport, as observed in Spinacia oleracea seedlings (Chen et al., 2011). 154 II. Manuscripts However, because in the absence of H2S the recovery time of photosynthesis after darkness and anoxia was extraordinarily long (up to 6 h, compared to minutes measured by Ghysels et al., 2013 or Schreiber et al. 2002) and the recovery was also delayed after prolonged darkness under aerobic conditions (up to 4 h, data not shown), it is more likely that H2S has yet another crucial regulatory role. Enzymes involved in the Calvin cycle are generally activated indirectly by light. The mechanism behind this activation is the reduction of disulphide-bonds, which increases the catalytic capability of the respective enzyme. The reductant needed for this reaction is thioredoxin, which in turn receives electrons from ferredoxin at a light-dependent rate (Schürmann and Buchanan, 2008; Michelet et al., 2013). Therefore, we hypothesize that the function of H2S in Planktothrix str. FS34 is similar to that of thioredoxin, i.e., it directly activates enzymes involved in the carbon fixation pathway by disulphide bond reduction. Further research is required to directly identify the mechanisms involved in the acceleration of the recovery of photosynthesis in Planktothrix str. FS34 after prolonged darkness. Light-dependent inhibition of photosystem II activity by H2S We found that H2S enhances photosynthesis in Planktothrix str. FS34 temporarily, i.e., during an active state. This active state can last for several hours at low light intensities but is substantially shortened or even disappears at higher light intensities (Fig. 2). The conspicuous light-dependency of the inhibition in Planktothrix str. FS34 suggests that H2S binds to an intermediate that is formed at a rate that is determined by the light intensity. We propose that this intermediate is generated during the light-driven water oxidation process in the OEC, because H2S was shown to inhibit PSII activity at its donor site (Miller and Bebout, 2004). We have incorporated this concept into a basic model of the electron transport processes in PSII (Fig. 4). Importantly, in our model the reactions that actually convert the light energy into chemical energy, i.e., the excitation of the catalytic Chl a dimer in PSII (kE1), the reduction of the primary electron acceptor QA (kQ) and the generation of the intermediate in the OEC (kC), are not directly affected by H2S. They only indirectly depend on H2S through an inactivation of an intermediate formed in the OEC. Because our data do not allow us to identify this intermediate, we refer to it generally as OECox. Nevertheless, based on the present knowledge, this intermediate could be anything formed at the donor site of the catalytic Chl a – starting from the oxidized catalytic tyrosine-residue that mediates electron transport between the reduced 155 4. REGULATION OF CYANOBACTERIAL PHOTOSYNTHESIS BY H2S Manganese centre of OEC to the oxidized P680 to any intermediate state formed in the S- cycle of OEC described in (Jablonsky and Lazar, 2008). PSI k E3 E P680* PQ k ox E1 PSI* k Q2 E k Q1Q1 ENPQ PQred k E2 k E4 PSIox P680 P680ox k C NADP+ H2S k S1 OECox:H2S OECox OEC k S2 CO k 2 H2S O H2O O2 Figure 4: Proposed model for the kinetic control of the reactions in the photosystem II of * Planktothrix str. FS34 by H2S. E is the photon flux; P680 and P680 is the Chl a dimer in the ground and excited state, respectively; P680ox is the oxidized form of the Chl a dimer that receives electrons from the oxygen evolving complex (OEC). OECox is the intermediate formed during H2O oxidation that is inhibited by H2S. OECox:H2S is the bound (and thus inhibited) form of this intermediate; PSI is the reaction centre of PSI in the ground state; PSI* is the excited catalytic Chl a in the PSI reaction centre; PSIox is the oxidized form of the PSI reaction centre that can receive electrons from reduced plastoquinone (PQred). Definitions of the process rates (ki) are given in Table 1. Model implementation was done in R (www.cran.r-project.org) using the deSolve package (Soetaert et al., 2010). Numerical implementation of this model based on the rate laws described in Table 1 indeed enabled us to simulate the biphasic pattern of photosynthetic oxygen production observed in Planktothrix str. FS34 at low light intensities and the fast establishment of the inactive state at high light intensities (compare Fig. 2 with Fig. 5A and 5C). Specifically, the long-term stable active state at low light intensities occurs because the OEC is only slowly removed from the reactive pool through binding to H2S (compare the evolution of OEC and OECox:H2S in Fig. 5B). This makes it still available for the interaction with the oxidized catalytic Chl a dimer in PSII (P680ox), the generation rate of which is not directly affected by H2S and continues dependent on the light intensity. Only upon inhibition of the complete OEC pool by H2S does the photosynthetic rate rapidly decrease to zero. In contrast, a stable active state does not occur at high light intensities because the intermediate (OECox) is generated at a higher rate, which leads to a more rapid accumulation of OECox:H2S and consequently to a faster depletion of OEC available for 156 II. Manuscripts the oxidation by P680ox (Fig. 5D). We therefore conclude that the likely mechanism of the inhibition of oxygenic photosynthesis in Planktothrix str. FS34 is based on binding of H2S to an oxidized intermediate of the oxygen evolving complex whose formation is driven by light. -2 -1 -2 -1 10 μmol photons m s 20 10 μmol photons m s 100 15 A B 80 ) -1 60 10 OE C s -1 OE C 40 ox L 5 OE C :H S 2 ox 2 20 0 0 -2 -1 -2 -1 100 μmol photons m s 100 μmol photons m s 200 100 150 C D 80 GP (nmol O (nmol GP 60 100 40 % of total OEC pool 50 20 0 0 0123456 0123456 time (h) time (h) Figure 5: Simulation of the transition from an active state to an inactive state in Planktothrix str. FS34. Shown are temporal dynamics of the volumetric rates of gross photosynthesis (GP) at low (A) and high (C) irradiance together with the corresponding partitioning of the OEC pool (B and D). In both cases H2S concentration of 200 μM was used. Note that the area under the two curves in (A) and (C), i.e., the total amount of O2 produced, is the same. Thus, at a certain H2S concentration a fixed amount of O2 can be produced before complete inhibition occurs and only the time that it takes to generate this specific amount depends on light. In our model this total amount of O2 produced is defined by the ratio between the rates kO and kS1 (kO:kS1=İ/(ț [H2S]); Table 1). Enhancement of the energy conservation efficiency in PSII and PSI by H2S Our data showed that oxygenic photosynthesis in Planktothrix str. FS34 is enhanced by H2S at low light intensities (Fig. 1A, 3B). Intriguingly, H2S appears to additionally transform the normally linear increase of photosynthesis with irradiance into a quadratic one (Fig. 3B). The initial slope of the PI-curve is mainly determined by the energy conversion efficiency of PSII, while the decrease of the slope at higher light intensities and the resulting saturation of photosynthesis (Fig. 3A) are determined by the rate limiting steps downstream of PSII, which can either be plastoquinone diffusion time or carbon fixation rate in the Calvin cycle (Sukenik et al., 1987; Cardol et al., 2011). In Spinacia oleracea seedlings, H2S increased the maximal photosynthetic rate, but did not affect the initial slope of the PI-curve (Chen et al., 2011). This increase was accompanied by an increased activity of RuBisCO and other photosynthetic components mediating the electron transfer between PSI and CO2 (Chen et al., 2011). Also in Planktothrix str. FS34 H2S appears to affect PSI-associated electron transport, which accelerates recovery of photosynthetic rates after exposure to darkness (see above). The enhancement of the rates, however, cannot exclusively be based on this effect of H2S. This is because the 157 4. REGULATION OF CYANOBACTERIAL PHOTOSYNTHESIS BY H2S enhancement in Planktothrix str. FS34 occurred already in the range where the PI curve is normally linear and not in the range where light saturation occurs. Thus, the effect of H2S cannot be limited only to the acceleration of electron transport downstream of PSII. We propose that H2S also has an effect on the energy conversion efficiency in PSII. Energy conservation efficiency can be increased by increasing the rate of excitation of the catalytic Chl a dimer, P680 (kE1 in Fig.4), or by decreasing the rate of excitation energy loss into heat and fluorescence (kE2). The rate kE1 can either be increased by increasing the pool of P680, i.e., the number of photosystems, or by increasing the absorption cross-section of PSII-associated pigments (factor Į in Table 1). In our model we assumed the latter. Specifically, we used the Michaelis-Menten kinetics with a low half-saturation constant (~2 μM) in combination with a linear function to describe the increase in the absorbance cross-section with H2S and light, respectively. Although we presently lack a mechanistic explanation for this increase, the most probable mechanism is a light-driven chemical transformation in PSII that leads to conformational changes and eventually to an increase in the absorbance cross-section. This would be consistent with the fact that the increase in photosynthesis is rapidly achieved (Fig. 2). Assuming that only PSII activity is enhanced (kE1 in Fig. 4), rate limitation in PSI (kE3 in Fig. 4) is expected when kE1 approaches a certain threshold. This is because the electron transport components between PSII and PSI, e.g., plastoquinone (PQ), would become rapidly fully reduced and could not receive electrons from PSII anymore, which would slow down the complete electron transport process. The “bump” on the PI-curve at low irradiances indeed suggests that rate limitation in PSI plays a role above an H2S- dependent light threshold. However, our data show that after this “bump” the rates of photosynthesis are higher than those measured in the absence of H2S. This implies that the potential PSI-associated electron transport rates must also increase in the presence of H2S. A possible mechanism could be an increase in the maximum rate of CO2 fixation in the Calvin cycle. However, since this positive effect on the Calvin cycle would mainly increase the slope of the PI curve above the “bump”, which would be inconsistent with our data, we deem this mechanism as unlikely. Another mechanism could be an increase in the absorbance cross-section in PSI (parameter ș in Table 1), plausibly due to the redistribution of excitation energy between the photosystems (state transition (Allen et al., 1989)) or an increase of the number of PSIs in the thylakoid membrane. We implemented this by assuming that the parameter ș increases with H2S according to the Michaelis- Menten kinetics but is light-independent. 158 II. Manuscripts Table 1: Definition of the rate laws governing the redox reactions shown in Figure 4, and values of the constants used for the simulations in Figures 5 and 6. Expression* Description Constants‡ The rate of generation of an excited catalytic Chl a dimer (P680*) in PSII. It depends on irradiance (E in μmol photons m-2 s-1), the availability of the -1 kE1 = Į E [P680] ground state Chl a ([P680] in nmol L ) and the -3 2 -1 Į0 = 1 x 10 m μmol absorbance cross-section factor Į that describes the -6 4 -2 [H 2S] Ș = 30 x 10 m s μmol Į = Į0 + Ș efficiency of conversion of the externally available -1 KS = 2 μmol L K S +[H 2S] photon flux into a volumetric rate of pigment excitation. The absorbance cross-section is assumed to increase with the H2S concentration and irradiance. The rate of reformation of the ground state P680 from the excited state P680*. This reformation yields fluorescence and heat. Together with Į, the ȕ = 1 x 10-3 s-1 kE2 = ȕ [P680*] parameter ȕ determines the relation between the external photon flux and the potential electron transport rate. [PQ ] The rate of P680* oxidation coupled to the ox -1 kQ1 = Ȗ [P680*] reduction of oxidized plastoquinone (PQox). This Ȗ = 1 s K g +[PQ ox ] -1 process results in the formation of a highly reactive Kg = 50 nmol L oxidized Chl a dimer (P680ox). The rate of reduced plastoquinone (PQ ) oxidation [PQ ] red red coupled to the reduction of oxidized photosystem I -1 kQ2 = ȡ [PSI ox ] ȡ = 100 s K +[PQ ] (PSIox). This rate controls the availability of PQox -1 r red Kr = 5 μmol L for the reduction by PSII and PSI in the ground state. The rate of generation of an intermediate in the -1 -1 kC = į [P680ox] [OEC] oxygen evolving complex (OECox), driven by į = 0.1 L nmol s P680ox. The rate of H2O oxidation to O2 coupled to the reduction of OECox. Division of kO in nmol -1 -1 _1 İ = 1 s kO = İ [OECox] electrons L s by 4 gives the rate of oxygenic -1 -1 photosynthesis in nmol O2 L s . The rate of binding of H2S to OECox. This binding -6 -1 -1 kS1 = ț [OECox] [H2S] ț = 20 x 10 L μmol s leading to the formation of OECox:H2S. The rate of the reverse reaction that results in the -6 -1 kS2 = ı [OECox:H2S] ı = 0.2 x 10 s release of OECox from OECox:H2S. The rate of generation of an excited reaction centre kE3 = ș E [PSI] in PSI. It depends on irradiance (E), the availability -3 2 -1 ș0 = 1 x 10 m μmol of the ground state PSI and the absorbance cross- -6 2 -1 [H 2S] ȗ = 2 x 10 m μmol ș = ș0 + ȗ section factor ș. In contrast to factor Į, factor ș is -1 K +[H S] KH = 500 μmol L H 2 assumed to increase only with the H2S concentration but not with irradiance. The rate of photosystem I (PSI*) oxidation. PSI* oxidation is coupled to NADP+ reduction and -1 -1 [PSI*] finally to CO2 fixation in the Calvin cycle. Thus, Ȟ = 900 nmol L s k E4 =Q -1 K E [PSI*] we implemented the rate limitation in the Calvin KE = 4 μmol L cycle indirectly as a limitation of the PSI oxidation rate, which is given by the maximum rate Ȟ. *Rates are in nmol L-1 s-1, where L refers to the volume of the biofilm. ‡The values correspond to the assumed concentrations of 1000 nmol L-1 for the total P680 pool (P680 + P680* + P680ox), the total OEC pool (OEC + OECox + OECox:H2S) and the total PQ pool (PQox + PQred), -1 * and of 1500 nmol L for the total PSI pool (PSI + PSI + PSIox) in the biofilm. 159 4. REGULATION OF CYANOBACTERIAL PHOTOSYNTHESIS BY H2S Numerical implementation of these mechanisms and empirically-derived rate laws gave a reasonably good agreement between the experimentally determined (Fig. 3) and modelled (Fig. 6) PI curves. Therefore, we conclude that the stimulation of photosynthetic rates in Planktothrix str. FS34 by H2S is likely based on (1) a rapidly saturated increase of the absorbance cross-section in PSII and (2) a simultaneous, but less pronounced, increase of the light harvesting capabilities in PSI. 40 H S (μM) 2 35 0 1 ) -1 30 100 s 150 -1 25 200 L 2 250 20 15 10 GP (nmol O (nmol GP 5 0 0 50 100 150 -2 -1 irradiance (μmol photons m s ) Figure 6: Simulation of the PI curves in Planktothrix str. FS34 at different H2S concentrations. The rates of gross photosynthesis (GP) are non-zero only if the active state was stable for more than 60 min. Implications for the ecophysiology of Planktothrix str. FS34 In microbial mats and biofilms anoxygenic photosynthesis and aerobic sulphide oxidation (driven by photosynthetically produced O2) are often responsible for the depletion of sulphide in the photic zone (Jørgensen et al., 1986). These processes are, however, restricted to the times of daylight. During night biotic and abiotic sulphide removal is - limited by the supply of oxidant (e.g., O2, NO3 ) from the overlying water column and consequently, sulphide concentrations in the photic zone generally increase (Jørgensen et al., 1986). Our measurements reveal that during early mornings such high H2S concentrations are not necessarily disadvantageous for obligate oxygenic phototrophic cyanobacteria such as Planktothrix str. FS34. Specifically, H2S is even crucial to shorten the recovery time of photosynthesis after darkness during night and, especially under low light intensities, H2S can temporarily increase the photosynthetic rate. However, our data indicate that their niche within sulphidic environments is rather narrow, as their activity strongly depends on defined temporal dynamics of H2S and light, and their interplay. Specifically, the H2S concentrations must be <230 μM, otherwise 160 II. Manuscripts inhibition would occur instantaneously also at low light intensities, whereas at about 180 -2 -1 μM H2S concentration and 40 μmol photons m s , photosynthesis is stable at an enhanced rate for more than 3.5 hours. If comparable conditions occur in the microenvironment of Planktothrix str. FS34 during the early morning, it will rapidly recover from previous dark anoxic conditions and perform photosynthesis at an enhanced rate. Its oxygenic photosynthesis will then indirectly modulate the local H2S concentration, even though it does not perform anoxygenic photosynthesis. Namely, the CO2 fixation driven by oxygenic photosynthesis will lead to an increase in pH and thereby a decrease in H2S in its microenvironment. Additionally, the H2S levels will also decrease due to the uptake of sulphide by anoxygenic phototrophs and chemotrophic sulphide oxidizers. This means that the inactivation by H2S (Fig. 2) will not occur in natural biofilms, as the sulphide species H2S is quickly removed due to the strong photosynthesis-driven pH increase, similarly to the dynamics in Fig. 1. But even without efficient H2S removal, it is expected that Planktothrix str. FS34 can remain in an active state for a complete day, given that the light intensity is very low. Also, Planktothrix str. FS34 is able to migrate inside a biofilm or a microbial mat and thus these cyanobacteria can plausibly escape from disadvantageous microenvironmental conditions. For instance, -2 -1 at 190 μM H2S and 130 μmol photons m s photosynthesis is completely inhibited after ~30 min. Migrating either deeper into the biofilm to reduce the light intensity or into a less sulphidic layer can, however, prevent the inhibition. Therefore, cyanobacteria like Planktothrix str. FS34 are probably effective competitors to obligate anoxygenic phototrophs and to cyanobacteria that can perform both anoxygenic and oxygenic photosynthesis. Implications for current models of ancient ocean chemistry The regulatory functions of H2S on oxygenic photosynthesis identified in this study for the cyanobacterium Planktothrix str. FS34 might have played a globally important role before the complete oxygenation of Earth’s atmosphere and oceans (Johnston et al., 2009) and eventually also during the subsequent sulphidic events that are thought to have caused global mass extinctions (e.g. in the end of Permian period, (Wang, 2012)). However, it is important to stress in this context that H2S is advantageous for photosynthesis in cyanobacteria like Planktothrix str. FS34 only if the exposure time to H2S is limited, or if the light intensity or H2S concentration is low. This means that such cyanobacteria will be competitive as free-living planktonic organisms in a sulphidic water column where H2S 161 4. REGULATION OF CYANOBACTERIAL PHOTOSYNTHESIS BY H2S concentrations are not substantially fluctuating only if both light intensity and H2S concentration are low. The latter implies electron donor limitation for obligate anoxygenic phototrophs. Thus, oxygenic phototrophs like Planktothrix str. FS34, which are never limited in electron donor, might have had a selective advantage. Such cyanobacteria would also be successful in environments where H2S concentrations were fluctuating – either by their own activity (i.e., through the modification of pH) or by the interaction with phototrophic or chemotrophic sulphide oxidizers. In microbial mats that have dominated shallow coastal regions for billions of years and in aggregates formed in the water column, the regulatory functions of H2S might have provided cyanobacteria like Planktothrix str. FS34 a selective advantage. The competitiveness of such obligate oxygenic phototrophs with anoxygenic phototrophs (both obligate and versatile cyanobacteria) in a sulphidic environment was plausibly crucial in the oxygenation process of our planet. This is because the competitiveness implies that oxygenic photosynthesis might have dominated under certain conditions despite abundant electron donor (H2S) for anoxygenic photosynthesis. The introduction of O2 to a sulphidic environment would have opened the stage for other competitors to anoxygenic photosynthesis, namely the light-independent aerobic sulphide oxidizers. Therefore, even if their habitat was limited to temporarily fluctuating environments, sulphide-adapted cyanobacteria like Planktothrix str. FS34 might have shifted the global balance between anoxygenic and oxygenic photosynthesis, which represents a key aspect of the oxygenation of Earth. Experimental procedures Isolation and cultivation The cyanobacterial strain FS34 was isolated from a biofilm growing in a sulphidic spring emerging from the Frasassi Cave system (Galdenzi et al., 2008). Specifically, the culture was obtained by dilution series in sulphidic and non-sulphidic medium and by picking of individual filaments from solid medium as previously described (Klatt et al., 2015).. The mono-algal culture was not axenic; however, microscopy revealed that the ratio between the cyanobacterial and other cells was >250. The contaminant cells were most likely not aerobic sulphide oxidizers, as we did not observe any biological sulphide consumption even in the presence of high oxygen concentrations (Fig. 1, Fig. S3). Also their contribution to oxygen removal was <2% and therefore negligible (data not shown). During the whole isolation procedure, the cultures were incubated at 15 °C under 162 II. Manuscripts irradiance of 200 μmol photons m-2 s-1 provided by fluorescent lamps (Envirolight, USA). For maintenance, cultures were transferred every 3–6 weeks alternating to BG-11 and sulphidic Pfennig's medium I. For the batch reactor experiments strain FS34 was grown in stirred liquid BG-11 medium until the exponential phase and then transferred into the reactor as previously described (Klatt et al., 2015). For the experiments in the flow cell, strain FS34 was also grown in BG-11 medium on a rotary shaker, which led to the formation of floating rigid biofilms that were transferred to and fixed in the flow cell. DNA extraction, polymerase chain reaction, sequencing and phylogenetic analysis DNA was extracted as previously described (Ionescu et al., 2010; Klatt et al., 2015). The partial 16S rRNA gene sequence was amplified using the cyanobacteria-specific primers CYA106F and CYA781R and the parameters for the polymerase chain reaction (PCR) as described by (Nübel et al., 1997). The PCR product of the partial 16SrRNA sequence was purified using the QIAquick PCR purification kit (Qiagen) and Sanger sequenced as previously described (Klatt et al., 2015). The partial 16S rRNA gene sequence is deposited in the European Nucleotide Archive under the accession number LN609763. For phylogenetic analysis, the 16S rRNA sequence was aligned with SINA (Pruesse et al., 2012), and was merged with the SILVA SSURef_NR99_115 dataset (Quast et al., 2013) for further processing using software package ARB (Ludwig et al., 2004). Along with the strain FS34 16S rRNA gene sequence, 20 full-length16S rRNA gene sequences from other cultivated strains belonging to the Oscillatoriales order were selected for phylogenetic tree reconstruction. Several treeing algorithms (neighbour joining, maximum parsimony, and maximum likelihood) with and without alignment position and base conservation filters were applied, and each attempt returned almost identical tree topologies. The final tree shown is calculated using RAxML version 7.0.4, with a GTRGAMMA model, a 10% cyanobacterial base conservation filter, and with 1000 bootstraps. The FS34 sequence was added later to this tree using the ARB parsimony insertion tool, again applying the 10% base conservation filter. Stirred batch reactor experiment To test whether strain FS34 is capable of anoxygenic photosynthesis using sulphide as an electron donor, culture harvested in the exponential growth phase was incubated in a stirred batch reactor without headspace (Klatt et al., 2015). The setup allows online 163 4. REGULATION OF CYANOBACTERIAL PHOTOSYNTHESIS BY H2S measurement of O2 concentration and pH and the assessment of the total sulphide 2- - concentrations (Stot = [S ]+[HS ]+[H2S]) in sub-samples taken during an incubation. In this experiment, we exposed the culture to Stot concentrations of up to 400 μM ([H2S] = 60 μM) for 5 min – 20 h in the dark, switched on the light (irradiance 20–500 μmol -2 -1 photons m s ) and measured O2 production and Stot consumption rates for at least 6 h. Flow-through chamber with a sulphidic bottom reservoir The experimental setup used for the measurements of photosynthesis in the biofilms formed by the studied cyanobacterial strain consisted of a top part that was constructed like a flow-through chamber (height: 10 cm, length: 25 cm, depth: 10 cm) and placed on top of a stirred reservoir (volume 2.4 L) (Fig. S1). The reservoir was filled with BG-11 medium to which Na2S stock solution (pH ~7.5, 200 mM) could be added by injection through a butyl rubber stopper. The chamber was separated from the reservoir by a polyester fibrous web on which the biofilm was fixed with pins. The flow chamber was fed from a thermostated recycle via gas-tight tubing. Oxygen concentration in the recycle and consequentially in the water column was adjusted by bubbling with N2 gas. To reduce exchange with air, paraffin oil was added on the water surface of the flow chamber and the thermostated recycle was covered with foil. Sulphide gradients across the biofilms were generated by the addition of Na2S from a stock solution to the reservoir and in some cases also to the recycle. Illumination of the biofilms in the flow chamber was provided by a halogen lamp (Schott KL2500). The incident irradiance at the surface of the biofilm was determined with a submerged cosine-corrected quantum sensor connected to a LI-250A light meter (both LI-COR Biosciences GmbH, Germany). Microsensors O2, pH and H2S microsensors with a tip diameter of 10–30 μm and response time of <1 s were built, calibrated and used as previously described (Revsbech, 1989; Jeroschewski et al., 1996; de Beer et al., 1997). Calibration of the H2S microsensors was performed in an acidified BG-11 medium (pH<2) to which Na2S was added stepwise. The Stot concentrations in the calibration solution was determined according to (Cline, 1969). 164 II. Manuscripts Microsensor measurements During simultaneous O2, pH and H2S measurements, the tips of the microsensors were always separated by less than 50 μm. Depth profiles were corrected for the azimuthal angle at which they were measured (Berg et al., 1998; Polerecky et al., 2007). Calculation of Stot from the local H2S concentrations and pH values measured with microsensors was done according to (Millero, 1986), using a pK value of 7.14 (Wieland and Kühl, 2000). Volumetric rates of gross photosynthesis (GP) were determined using the O2 microsensor based light-dark shift method (Revsbech and Jørgensen, 1983). Acknowledgements We thank Georg Herz and Jürgen Kohl from the mechanical workshop for their help with the construction of the experimental setup, the technicians of the microsensor group for the construction of microsensors, J. Macalady and D. S. Jones for their support during field campaigns, A. Montanari and P. Metallo for their hospitality during the sampling campaign at the Osservatorio Geologico di Coldigioco, and S. Meyer for her support during isolation and cultivation. This work was financially supported by the Max-Planck- Society. Conflicts of interest statement The authors declare no conflicts of interest. References Allen, J.F., Mullineaux, C.W., Sanders, C.E., and Melis, A. (1989) State transitions, photosystem stoichiometry adjustment and non-photochemical quenching in cyanobacterial cells acclimated to light absorbed by photosystem I or photosystem II. Photosynthesis research 22: 157-166. Beauchamp, R.O., Bus, J.S., Popp, J.A., Boreiko, C.J., and Andjelkovich, D.A. 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Biofilms (a) were fixed on a polyester fibrous web (b) that separated the flowing water column of the upper part from the reservoir in the bottom part. NaS2 stock solution was added to the reservoir through a butyl rubber stopper (c). Mixing in the bottom part was achieved by a magnetic glass stir bar (d). 170 II. Manuscripts Figure S2: Light microscope image of Planktothrix str. FS34 (scale bar is 5 μm) and a dendrogram showing the position of its 16S rRNA sequence with respect to sequences from other cultivated members of Oscillatoriales. The tree reconstruction was performed using Maximum Likelihood algorithm as implemented in RAxML. The two Gloeobacter violaceus sequences serve as the out-group. Bootstrap values are only shown when they were above 60%; the bar indicates 0.02 substitutions per nucleotide position. 171 4. REGULATION OF CYANOBACTERIAL PHOTOSYNTHESIS BY H2S dark light 500 140 120 400 100 (μM) (μM) 300 tot 2 80 S O 60 200 40 100 20 0 0 0 2 4 6 8 10 12 time (h) Figure S3: Oxygen and total sulphide concentration dynamics measured in a stirred batch reactor filled with Planktothrix str. FS34 culture. NaS2 stock solution was added to the culture in the dark to establish a final Stot concentration of 400 μM ([H2S] = 95 μM) in the reactor. Scalar irradiance -2 -1 in the reactor was 100 μmol photons m s . The decrease in the Stot concentration in the presence of photosynthetically produced oxygen was due to abiotic sulphide oxidation as confirmed by separate incubations of medium without culture. 172 II. Manuscripts 5. ANOXYGENIC PHOTOSYNTHESIS AND 2-METHYL HOPANOID PRODUCTION: A MODEL CYANOBACTERIUM ISOLATED FROM A PROTEROZOIC OCEAN ANALOG Trinity L. Hamilton1*, Judith Klatt2, Laurence M. Bird3, Katherine H. Freeman3, Dirk de Beer2, and Jennifer L. Macalady3* 1 Department of Biological Sciences, University of Cincinnati, Cincinnati, OH 45221, USA 2 Max-Planck Institute for Marine Microbiology, Bremen, Germany 3 Department of Geosciences and the Penn State Astrobiology Research Center (PSARC), Pennsylvania State University, University Park, PA 16802, USA In preparation * corresponding authors 173 5. A MODEL CYANOBACTERIUM FROM A PROTEROCOIC OCEAN ANALOG Abstract Understanding the evolution of Earth’s surface chemistry is one of the most exciting challenges in modern geoscience. The atmosphere and oceans of early Earth were oxygen-poor and, even after oxygen began to accumulate ~2.5 Ga, conditions of euxinia—a moderately oxic surface layer underlain by sulfidic waters at depth—were probably widespread throughout much of Earth’s middle age. Oxygen did not rise to present day levels until recently, ~0.5 billion years ago. Thus, the 1-2 billion year delay in the rise of oxygen at the end of the Proterozoic represents an important gap in our understanding of ancient biogeochemical cycling. Primary production fueled by sulfide- dependent anoxygenic photosynthesis, including the activity of metabolically versatile cyanobacteria, in euxinic regions of the oceans has been invoked as a mechanism for sustaining low atmospheric O2 throughout much of the Proterozoic yet we understand very little about cyanobacteria that inhabit analog environments present today and the biosignatures they leave behind. Here we report both anoxygenic photosynthesis and 2- methyl hopanoid production in a cyanobacterium isolated from Little Salt Spring—a karst sinkhole in Florida with low levels of dissolved oxygen and sulfide. The isolate requires no induction time following the addition of sulfide to perform anoxygenic photosynthesis. Under the optimal light conditions for oxygenic photosynthesis, rates of anoxygenic photosynthesis are nearly twice as high. This characterization represents a significant step toward unraveling the role of biology in planetary redox evolution. 174 II. Manuscripts Introduction Early Earth was likely oxygen-poor until ~2.5 billion years ago when a Great Oxidation Event (GOE) occurred raising the concentration of oxygen in the atmosphere from < 0.001 % of the present atmospheric level (PAL) to within 1-10 % PAL (Kump, 2008). Oxygen did not accumulate to levels observed today until ~0.5 Byr ago. Competition between oxygenic and anoxygenic phototrophs in the photic zone of early Earth oceans has been proposed as a positive feedback mechanism that delayed the oxygenation of the Earth's oceans and atmosphere during this time by stabilizing widespread ocean euxinia (Johnston et al., 2009). In this hypothesis, the outcome of ecological interactions among phototrophs in Proterozoic oceans may have had profound impacts on net oxygen accumulation in the atmosphere. This hypothesis, which invokes significant contributions to primary productivity from anoxygenic photosynthesis, including metabolically versatile cyanobacteria, has not been explicitly tested in a modern analog of Proterozoic ocean conditions and examples of cyanobacteria capable of capable of both oxygenic and anoxygenic photosynthesis are rare. Cyanobacteria are the only prokaryotes that carry out oxygenic photosynthesis and therefore played a crucial role in the evolution of Earth’s atmosphere as well as the emergence and diversification of complex life. Cyanobacteria are metabolically diverse including species that can perform anoxygenic photosynthesis in environments where sulfide is present in the photic zone (Cohen et al., 1975a; 1975b; Jørgensen et al., 1986; Garcia-Pichel and Castenholz, 1990), use hydrogen as an electron donor (Cohen et al., 1986), perform sulfide-dependent nitrogen fixation (Belkin and Padan, 1978), and grow photoheterotrophically (Kenyon et al., 1972). This phenotypic diversity enables cyanobacteria to tolerate a variety of environmental extremes, resulting in their ability occupy most environments where light is available and making them highly relevant to primary production in many ecosystems. For example, benthic cyanobacterial mats in Solar Lake, Sinai, a hypersaline pond, undergo drastic yearly changes in temperature, salinity, oxygen, light and H2S. The mats in Solar Lake are dominated by metabolically diverse cyanobacteria such as Geitlerinema sp. PCC 9228 (formerly Oscillatoria limentica) capable of anoxygenic photosynthesis and phototaxis (Cohen et al., 1975a; 1975b; Krumbien et al. 1977). Phormidium spp. survive freezing and desiccation in Antarctica (Taton et al., 2003). Phormidium spp. and Oscillatoria-like spp. survive in persistent low O2 conditions in the Middle Island Sinkhole of Lake Huron (Voorhies et al., 2012) and Little Salt Spring, a sinkhole in Florida (Hamilton et al., submitted). 175 5. A MODEL CYANOBACTERIUM FROM A PROTEROCOIC OCEAN ANALOG Today, the co-occurrence of cyanobacteria and anoxygenic phototrophs is common in euxinic lakes, cyanobacterial mats, hot springs, and hypersaline lagoons where sufficient fluxes of reduced compounds are available to support anoxygenic photosynthesis. The ability to harvest light and tolerance to oxygen are often cited as the key factors governing occurrence of phototrophs in ecological niches along stratified water columns and mats. For instance, green and purple sulfur bacteria (GSB and PSB) are found in most sunlit, sulfidic environments where PSBs are generally found at more shallow depths of the water column or mats (Meyer et al., 2011; Overmann and Garcia- Pichel, 2006). GSB have lower light requirements and are less tolerant to oxygen. In contrast, a combination of sulfide and temperature appear to inhibit photosynthesis in alkaline hot springs above ~70 ºC (Cox et al., 2011; Boyd et al., 2012). As such, the physical, geochemical and biological controls on the occurrence of phototrophs, the co- existence of phototrophs and the competition among co-occurring phototrophs are poorly understood. As oxygenic phototrophs, cyanobacteria have had profound effects on Earth’s environment and atmosphere for billions of years but these microorganisms leave poor fossil records. The most robust microfossil record exists throughout the Proterozoic where carbonate stromatolites indicative of cyanobacteria and organic biomarkers, including 2- methyl hopanes, and have been found (Des Marais, 2000). Bacteriohopanepolyols (BHPs) are bacterial pentacyclic triterpenoids produced by a number of bacteria and are preserved in the rock record as hopanes. The utility of hopanoids methylated at the C-2 position— originally thought to be diagnostic of cyanobacteria (Summons et al., 1999)—is currently unknown largely due to the fact that other organisms, including anaerobes, also produce them. These structures have been recovered from modern environments as well as ancient marine and sedimentary rocks. The distribution of hopanes in the geologic record suggest a higher abundance of the C-2 methylated forms in rocks of the Proterozoic and Archean (Summons et al., 1999; Knoll et al., 2007) and increased abundance associated with oceanic anoxic events during the Phanerozoic (Talbot et al., 2008). Our current interpretation of the biogeochemical significance of these biomarkers in ancient rocks is hindered by the taxonomic and metabolic breadth of bacteria that produce them. A better understanding of the ecological and biogeochemical constraints affecting the oxygen budget in Proterozoic ecosystems coupled to investigations of model organisms that produce geologically relevant biomarkers is paramount to unraveling the Earth’s redox evolution. Little Salt Spring, a karst sinkhole in Florida with Proterozoic- 176 II. Manuscripts like water chemistry, hosts a seasonal bloom of red pinnacle mats comprised largely of cyanobacteria and GSB (Hamilton et al., submitted). The mat contains hopanoids, including those methylated at the C-2 position. Here, we report the isolation and draft genome of a metabolically versatile Oscillatoriales-like cyanobacterium from Little Salt Spring. The isolate is capable of anoxygenic photosynthesis and also produces C-2 methyl hopanoids. A gene encoding a sulfide:quinone oxidoreductase, the only additional enzyme implicated in anoxygenic photosynthesis in cyanobacteria, is present in the draft genome. The genetic machinery necessary for methylating hopanoids at the C-2 position (hpnP). The isolate is phylogenetically distinct from other cyanobacteria capable of anoxygenic photosynthesis. Anoxygenic photosynthetic activity was observed immediately upon addition of sulfide and light, requiring no induction time and rates of anoxygenic photosynthesis were higher than oxygenic photosynthesis under optimal light conditions. This isolate represents an important model organism for examining metabolic diversity in cyanobacteria and the synthesis of 2-methyl hopanoids. These have important implications in interpreting the role of cyanobacteria in planetary redox evolution and for interpreting biosignatures in the rock record. Materials and Methods Sample collection Red pinnacle mat samples were collected by divers in June of 2012 from Little Salt Spring, a 78 meter diameter sinkhole lake located in Sarasota County, FL (lat. 27°04'30"N, long. 82°14'00"W). The geology and hydrology of the sinkhole have been described previously (Zarikian et al., 2005) as well as the spring geochemistry and microbiology of the red pinnacle mat (Hamilton et al., submitted). Diver-collected pinnacle mats were sampled from the water-sediment interface at ~14 m. Mat samples for lipid, pigment, and nucleic acid analyses were preserved in RNALater (Ambion, 3 parts per sample volume) and stored at -20 ºC until further processed. Samples of biofilm for enrichment were transferred to sterile 50-mL conical tubes, overlaid with spring water collected from the same location as the mat sample, and stored in the dark at 4 ºC prior to inoculation. 177 5. A MODEL CYANOBACTERIUM FROM A PROTEROCOIC OCEAN ANALOG Enrichment and isolation Samples of red pinnacle mat were homogenized and a small aliquot (~ 500 uL) was added to BG11 media (Rippka et al., 1979) supplemented with 25 mM HEPES (B-HEPES) and adjusted to pH 7.2. Enrichment cultures were maintained in 60-mL of liquid media in 125-mL conical flasks at 100 RPM at 28 ºC under either a day-night cycle or continuously illuminated with 100 ȝmol photons m-2 s-1 under cool white fluorescent lamps. An axenic culture was achieved using a dilution series in liquid media where the highest dilution which showed growth was taken as inoculum for the next dilution. Light microscopy was performed periodically to visually examine enrichments for purity. The dilution to extinction strategy was continued until light microscopy indicated the presence of a single morphotype and sequencing of the 16S rRNA gene returned a single phylotype. Growth of heterotrophic contaminants was tested on agar plates incubated in the dark. Growth of the isolate was monitored with chlorophyll a concentration determined spectrophotometrically using the absorption at 665 nm of a methanol extract and an extinction coefficient of 0.075 ml ȝgí1 (made from a filtered 2-ml culture subsample) (De Marsac and Houmard, 1988) or protein concentration using the Bradford assay (Bradford, 1976) with bovine serum albumin (Sigma-Aldrich, St. Louis, MO) as the standard. Nucleic acid analyses Samples of biofilm (~ 1.5 mL) were harvested by centrifugation, the excess media removed by decanting, and cell pellets frozen immediately (-20 °C) or subjected to nucleic acid extraction. Genomic DNA was extracted as previously described (Boyd et al., 2007). For amplification of near full-length 16S small subunit RNA genes, bacterial domain primers 27F and 1492R were used (Lane, 1991) with the following thermal cycling conditions: 5 min at 94 °C for initial denaturation followed by 30 cycles of PCR consisting of 60 s at 94 °C, 30 s at 52 °C and 90 s at 72 °C, followed by 20 min at 72 °C for final elongation. Reactions were performed in triplicate, purified using a QIAquick PCR Purification Kit (Qiagen, Valencia, CA) and sequenced at the Genomics Core Facility of the Huck Institutes of the Life Sciences at Penn State University. Sequences were assembled and manually checked using using Bio-Edit (v7.2.5), and checked for chimeras using CHIMERA_CHECK (Cole et al., 2003). Putative chimeras were excluded from subsequent analyses. 178 II. Manuscripts Pigment analysis Pigment extraction and HPLC analyses were performed as previously described (Frigaard et al., 2004). Briefly, samples of pure culture were concentrated by centrifugation (12 000 g), and pigments were extracted from 200 ȝL pellets by sonication in acetone/methanol (7:2, v/v). Extracts were centrifuged (12 000 g) to remove debris and filtered through a 0.22-ȝm Teflon syringe filter. Pigments were separated by HPLC (Agilent Model 1100; Agilent Technologies, Palo Alto, CA, USA) equipped with a diode array detector (model G1315B) on a 25 cm by 4.6 mm analytical discovery 5-ȝm C18 column (Supelco, Bellefonte, PA, USA); the system was controlled by Agilent ChemStation software (Agilent Technologies). Genomic sequencing, assembly, and completeness A draft genome of the LSS cyanobacterium was generated from genomic DNA extracted as described above. Purified DNA was sequenced with on an Ion Torrent Personal Genome Machine according to the Ion Torrent protocol at the Penn State University sequencing facility. Briefly, double-stranded gDNA was quantified by Qubit (Life Technologies, Carlsbad, CA) and 100 ngs of DNA was fragmented for 7 minutes at 37 ºC using the Ion Shear Reagent according to the Ion Xpress™ Plus gDNA and Amplicon Library Preparation Users Guide (Life Technologies, Carlsbad, CA). Fragments were purified using AMPure XP Reagent (Beckman Coulter, Brea, CA) and checked for size on a Bioanalyzer DNA High Sensitivity chip (Agilent Technologies, Santa Clara, CA ). Adaptors were ligated to the fragments and DNA was nicked, repaired and then purified. A 315 bp library was size-selected on a Pippin (Sage Science, Beverly MA) using the "tight setting". Following purification, the library was checked again on the Bioanalyzer and then amplified on beads in an emulsion PCR using the Ion Xpress Template Kit and 200 base sequencing was performed using a 316 chip on the Ion PGM machine as per the Users Guide (Life Technologies, Carlsbad, CA) at the Genomics Core Facility of the Huck Institutes of the Life Sciences at Penn State University. The resulting reads were assembled with Newbler assembler version 2.6 (Roche) resulting in 77 contiguous reads (contigs) of at least 500 bp with an average read depth of ~100x. Contigs were annotated using RAST (http://rast.nmpdr.org; Aziz et al., 2008) and manually curated. Genome completeness was evaluated using conserved housekeeping genes (Table S1) and phylogenetic marker genes identified with Phyla-AMPHORA (Wang & Wu, 2013). 179 5. A MODEL CYANOBACTERIUM FROM A PROTEROCOIC OCEAN ANALOG Phylogenetic analysis The 16S rRNA gene sequence from the LSS isolate was compared to sequences in public databases using BLASTN (Altschul et al., 1990). Sequences were added to an existing 16S rRNA alignment in ARB (Ludwig et al., 2004), and manually refined. Phylogenetic trees were computed using neighbor joining (general time-reversible model) with one thousand bootstrap replicates using PAUP (ver. 4.0) (Swofford, 2001). Neighbor-joining trees, also calculated using PAUP (ver. 4.0), were compared with maximum likelihood trees (general time-reversible model, site-specific rates and estimated base frequencies). For gene-specific phylogenetic analyses, full-length sequences were identified in the genome using functional annotation and BLASTX, translated, and verified by BLASTX/P. All BLAST analyses were performed using an e-value cut-off of 1e-5. Reference datasets were populated with sequences mined from the NCBI databases and JGI IMG-M using the same criteria. Protein sequences were aligned with CLUSTALX (version 2.1) with the Gonnet 250 protein substitution matrix and default gap extension and opening penalties (Larkin et al., 2007). The best evolutionary model was determined for each single-protein alignment using ProtTest (version 3, Darriba et al., 2012). The phylogeny was evaluated using PhyML (Guindon and Gascuel, 2003) or RAxML with one thousand bootstrap replicates, employing the best evolutionary model as identified by ProTest. Maximum parsimony and maximum likelihood trees yielded similar results in all analyses. Hopanoid analyses Samples of pure cultures were concentrated by centrifugation (12 000 g), freeze-dried, homogenized, weighed, and extracted using a microscale modification of the protocol prescribed by Talbot et al. (2003), substituting dichloromethane (DCM) for chloroform in the procedure. An aliquot of each total lipid extract (TLE) was acetylated using 1:1 pyridine: acetic anhydride at 60°C for 1 hr. Bacterophopanpolyols (BHPs) were characterized using an Agilent 6310 high-pressure liquid chromatography/mass spectrometry (HPLC/MSn) ion trap with an atmospheric pressure chemical ionization (APCI) chamber. Reverse-phase HPLC was performed using a Phenomenex Gemini 5 ȝm C18 column (150 mm x 3.0 mm i.d.) and a 5 ȝm pre-column with the same solid phase. Mobile phase flowed at 0.5 ml/min and was composed of (A) water, (B) methanol, and (C) isopropyl alcohol (0 min A: B: C 10: 90: 0, with a ramp to 25 min 1: 59: 40 and an isocratic flow to 45 min). LC/MS settings were the same as reported in Talbot et al., 180 II. Manuscripts 2003: positive APCI mode, mass scanning from 150-1300 m/z, and “Smart” fragmentation setting (i.e., 8000 nA corona voltage, 60 psi nebulizer pressure, dry gas of five l/min, dry temperature of 350 C, vaporizer temperature 490 C). The run was divided into three segments, with target masses of 285 (0-10 min), 1002 (10-17 min) and 655 (17- 50 min), all with normal optimization. Auto MSn settings for two precursor ions were an absolute threshold of 100,000 and a relative threshold of 5%. The most abundant ion is excluded after two measurements and released after 0.5 min. The acquisition parameter had a fragment amplitude 1.0V, with an average of 5.0 and an isolation width of 3.0 m/z. Auto MS (n > 2) settings with one precursor ion were as follows: absolute threshold of 1,000, relative threshold 5%, and fragment amplitude 1.0V. Standards consisted of individual BHP compounds from samples collected at Bear Meadows Bog, Rothrock State Forest, PA. Standards were fraction-collected following reverse phase separation. Each collected fraction was additionally purified via neutral phase separation on a silica column (150 mm x 3.0 mm, Waters) and an isocratic flow of 80:30 hexanes:isopropyl alcohol (IPA) at 1 ml/min. The fractions were then carefully transferred, dried and weighed using tin EA cups on a nanogram scale. Compounds collected for use as standards include: BHT (m/z 655), Aminotriol (m/z 714), and BHTriol cyclitol ether (m/z 1002). Microsensor measurements Rates of oxygenic and anoxygenic photosynthesis dependent on irradiance and H2S concentration were determined in biofilms formed by the cyanobacterium using microsensors. Mats were grown on pre-sterilized glass fiber filters to facilitate the measurements. Single filters were then transferred to a polyester fibrous web at the bottom of a 500 mL custom-made aquarium and held in place with pins. The aquarium was filled with B-HEPES media. When necessary, sulfide was added from a neutralized Na2S stock solution (pH 7 – 7.5). To limit diffusive loss of sulfide, a thin layer of paraffin oil (Merck, Kenilworth, NJ) was added to the surface of the medium. A circular flow of the water column above the biofilm was achieved by streaming N2 gas onto the paraffin- oil surface. Illumination in the visible range was provided by a halogen lamp (Schott KL- 2500, Mainz, Germany) mounted above the aquarium. The incident irradiance at the surface of the biofilm was determined with a submerged cosine-corrected quantum sensor connected to a LI-250A light meter (both LI-COR Biosciences GmbH, Germany). 181 5. A MODEL CYANOBACTERIUM FROM A PROTEROCOIC OCEAN ANALOG O2, pH and H2S microsensors with a tip diameter of 10–30 μm and response time of <1 s were built, calibrated and employed as previously described (Revsbech, 1989; Jeroschewski et al., 1996; de Beer et al., 1997; Klatt et al., 2015a). The microsensor tips were positioned at the biofilm surface and always separated by less than 50 μm during simultaneous O2, pH and H2S measurements. All measurements were performed at room temperature. Volumetric rates of gross oxygenic photosynthesis (GOP) were determined based on the dynamics of oxygen concentration after a light-dark shift as described previously (Revsbech and Jørgensen, 1986). Analogously, volumetric rates of gross anoxygenic photosynthesis (GAP) were calculated from the dynamics of H2S concentration and pH directly after a light-dark transition (i.e. light-driven Stot consumption rates) as previously described (Klatt et al., submitted). Prior to each measurement in the presence of sulfide, GOP as a function of irradiance (9 – 289 μmol photons m-2 s-1) was quantified (PI-curve). Irradiance was then adjusted to a specific value chosen from the PI-curve and GOP was determined again, prior to addition of sulfide. Following injection of sulfide into the water column, the local H2S concentration, GOP and GAP in the surface of the biofilm were monitored until complete depletion of sulfide and recovery of GOP. After recovery of GOP, the PSII inhibitor, 3-(3,4-dichlorphenyl)-1,1-dimethylurea (DCMU; Sigma-Aldrich, St. Louis, MO) was added to the water column (final concentration: 10 μM) during exposure to light and before the re-addition of sulfide. The local H2S concentration, GOP and GAP in the mat were again monitored until complete depletion of sulfide and recovery of GOP. This procedure was subsequently repeated at several light intensities on fresh mat samples. To compare GAP and GOP, all reactant consumption/production rates were converted to theoretical electron transport rates as previously described (Klatt et al., 2015a; Table S2). Briefly, assuming zero-valent sulfur is the primary product of anoxygenic photosynthesis, rates of total S consumption (Stot) were multiplied by a factor of 2 to account for two electrons per oxidized sulfide that are theoretically transported and + used for NADP reduction. Analogously, the rates of O2 production were multiplied by a factor of 4. To allow for the comparison of results obtained in different biofilm samples, we normalized GOP and GAP to GOPmax, the maximum rate of gross oxygenic photosynthesis in the respective mat sample which was determined using the light-dark method as described above. 182 II. Manuscripts CO2 photoassimilation The contribution of Photosystem II (PSII)-independent anoxygenic photoassimilation of CO2, with H2S as the electron donor, was determined in assays containing the PSII inhibitor DCMU. All experiments were carried out at 30 ºC in B-HEPES media at pH 7.2. Exponentially growing cells were harvested by centrifugation and resuspended in fresh O2-free media and introduced (10-mL suspensions) in 20-ml capped serum vials. Vials 13 were kept in the dark and purged with N2 to remove any remaining O2. NaH CO3 (Cambridge Isotope Laboratories, Inc., Andover MA) was added to each vial at a final concentration of 100 μM. To assess the PS-II independent anoxygenic CO2 assimilation, vials (n = 3 for each treatment) were amended with 100 μM neutralized Na2S and 10 μM DCMU (Sigma-Aldrich, St Louis MO). To analyze the affects of Na2S or DCMU alone on CO2 photoassimilation, vials (n = 3 for each treatment) were amended with 100 μM Na2S or 10 μM DCMU. A natural abundance control (n = 3 vials) received unlabeled -2 -1 NaHCO3 (100 μM). Vials were incubated at a light intensity of 100 ȝmol photons m s under cool white fluorescent lamps. An additional set of vials (n = 3 for each treatment) were prepared as described above and incubated under 700 nm wavelength light for 2 hours. To assess CO2 assimilation in the dark, vials (n = 3) were amended with 100 μM 13 NaH CO3 and placed in the dark for three hours. Following incubation, cells were harvested by centrifugation and freeze-dried. Dried samples were treated with concentrated HCl (1 M) to remove excess carbonate, washed with de-ionized H2O to remove excess acid and freeze-dried again. Samples were weighed and placed into tin dishes, sealed, and analyzed via an elemental analyzer with an isotope ratio mass spectrometer (EA-IRMS). All assays were performed in triplicate. GenBank accession numbers The draft genome of the LSS cyanobacterium is deposited at DDBJ/EMBL/GenBank under the accession numbers XXX; and the version described in this paper is XXX. Results Isolation of Leptolyngbya sp. LSS Red pinnacle mats collected from the sediment-water interface in Little Salt Spring, a karst cover-collapse sinkhole, were used for isolation. A serial dilution strategy coupled to multiple transfers of red filaments resulted in an axenic culture of a cyanobacterium 183 5. A MODEL CYANOBACTERIUM FROM A PROTEROCOIC OCEAN ANALOG which contains chlorophyll a, phycocyanin, phycoerythrin and allophycocyanin. The isolate is filamentous and non-mobile. The isolated cyanobacterium was named LSS. The phylogenetic position of Oscillatoriales cyanobacterium sp. LSS was evaluated by comparing a 1281 bp fragment of the 16S rRNA gene sequence with closely related cyanobacteria. Based on this analysis, the isolate is a member of the Oscillatoria. The 16S rRNA sequence of the isolate is most closely related to uncultured clones from biological soil crusts on copper mine tailings (KF287742) and clones from banana plantation soil (JX133481) and among cultured cyanobacteria, Geitlerinema sp. PCC 7407, and Oscillatoria acuminata PCC 6304 (~ 94% sequence identity)(Fig. 1). The 16S rRNA sequence of the isolate is identical to a sequence recovered from the red pinnacle mat in Little Salt Spring (Hamilton et al., submitted) (Fig. 1). A highly enriched culture of Oscillatoriales cyanobacterium UVFP2 was obtained from Fuente Podrida, a sulfide-rich spring (0.24-1.12 mM) close to the Cabriel River in Valencia, Spain (Camacho et al., 2005) while the isolation source of Geitlerinema sp. PCC 7407 and O. acuminata PCC 6304 are not known. Phylogenetic analysis of full-length 16S rRNA sequences from complete to nearly complete sequenced genomes (n=79) available from JGI indicates the isolate is most closely related to Geitlerinema sp. PCC 7407 and Oscillatoriales cyanobacterium JSC-1 (~ 94% sequence identity) (Fig. S1). Oscillatoriales cyanobacterium JSC-1 was isolated from a ferrous iron-rich circumneutral hot spring in Yellowstone National Park, WY, USA (Brown et al., 2010). Genome features A draft genome of Oscillatoriales cyanobacterium sp. LSS was assembled, containing 77 contigs and 5,940,030 bp with an average GC content of 52.3% (Table 1). The draft genome encodes 61 tRNAs and 5627 protein coding genes, including all of the conserved housekeeping genes (Table S1) and >96% of the phylum specific marker genes identified with PhylaAMPHORA, suggesting it is nearly complete. The genome encodes the enzymes necessary for aerobic photoautotrophic growth including Form I RuBisCO and a complete Calvin-Benson cycle; two high affinity terminal oxidases, a cytochrome c oxidase and a bd-type quinol oxidase; photosystems I and II and chlorophyll biosynthesis; and a cytochrome b6f complex. The bd-type quinol oxidase and cytochrome c oxidase differ in their affinity for oxygen (0.35 μM vs 1.0 μM) (Pils and Schmetter, 2001), and as a result, the former is postulated to operate under conditions of low oxygen tension (Hart et al., 2005). Genes for an F-type ATPase and a NAD(P)H:quinone oxidoreductase 184 II. Manuscripts (NDH) are present. Strain LSS also encodes the enzymatic machinery necessary for assimilatory nitrate and sulfate reduction as well as nitrogen fixation via a Mo-dependent nitrogenase. Figure 1: Maximum likelihood based phylogenetic 16S rRNA gene tree of closely related Cyanobacteria and strain LSS. Accession numbers are provided in parentheses. Bootstrap support values based on 1000 bootstrap samplings >90 are noted. Full tree is shown in Figure S1. Three enzymes integral to photosynthesis—coproporphyrinogen III oxidase, heme oxygenase, and Mg-protoporphyrin IX monomethylester cyclase—require oxygen for activity; however, oxygen levels at the depth of the red pinnacle mats in Little Salt Spring sampled are low (<30 μm) and the water column experiences weakly sulfidic conditions (<30 μm)(Hamilton et al., submitted). In the cyanobacterium Synechocystis sp. PCC 6803, alternative forms of these enzymes are present that operate under low oxygen conditions (Aoki et al., 2011). Homologs of both the aerobic and anaerobic forms of these enzymes are encoded in the LSS draft genome. A gene with sequence homology to sulfide:quinone oxidoreductase (SQR), which has been implicated in anoyxgenic photosynthetic activity in cyanobacteria (Shahak et al., 1998), was also encoded in the genome. SQR catalyzes the oxidation of sulfide to zero- valent sulfur and may play a physiological role in both energy transduction and sulfide 185 5. A MODEL CYANOBACTERIUM FROM A PROTEROCOIC OCEAN ANALOG detoxification. The translated SQR sequence of the isolate clusters with Type F SQRs (SQRF) (Gregersen et al., 2011) (Fig. 2) and is only distantly related (25% sequence identity) to the SQR sequences of Aphanothece halophytica and Geitlerinema sp. PCC 9228 (formerly Oscillatoria limnetica), the latter has been implicated in sulfide-dependent anoxygenic photosynthesis. Geitlerinema sp. PCC 7105 also encodes an F type SQR. The well-characterized SQR of Geitlerinema sp. PCC 9228 is an A type (SQRA), as were all other cyanobacterial SQR sequences identified here (Fig. S2). Type F SQR has been characterized in the GSB Chlorobaculum tepidium and is necessary for growth at sulfide concentrations 4 mM (Chan et al., 2009). Table 1: Statistics for the LSS cyanobacterium draft genome Genome features Total bp 5940030 Contigs 77 N50 (bp) 137782 %GC 52.32 Longest (bp) 544817 tRNA 61 Protein coding genes 5627 Red pinnacle mats collected from Little Salt Spring contain hopanoids, including those methylated at the C-2 position (Hamilton et al., submitted). The LSS genome encodes homologs of two enzymes that are presumably necessary for the biosynthesis of 2-methylhopanoids—a squalene-hopene cyclase and a radical SAM methylase (HpnP) (Welander et al., 2010). The translated HpnP homolog branches with other cyanobacterial HpnP sequences (Fig. 3). The isolate sequence shares 80% sequence identity with the HpnP sequence of Nostoc punctiforme which makes 2-methyl hopanoids and 59% sequence identity with the characterized HpnP from Rhodospeudomonas palustris TIE-1. 186 II. Manuscripts Figure 2: Maximum likelihood phylogeny of Sqr sequences. Classification of Sqr othologs is based on Gregersen et al., 2011. Accession numbers are given in parentheses. Black circles indicate bootstrap support values >90 based on 1000 bootstrap samplings. Full tree is shown in Figure S2. Figure 3: Maximum likelihood phylogeny of HpnP sequences. Black circles indicate bootstrap support values >90 based on 1000 bootstrap samplings. Accession numbers for all sequences are provided in Table S3. 187 5. A MODEL CYANOBACTERIUM FROM A PROTEROCOIC OCEAN ANALOG Hopanoids Analysis of the hopanoid content of the isolate indicates the cyanobacterium makes bacteriohopanetetral (BHT) and 2-MeAnhydro-BHT (Fig. S3). These observations are consistent with the recovery of both structures from the red pinnacles mats in Little Salt Spring (Hamilton et al., submitted). Oxygenic Photosynthesis Microsensors were used to determine the gross rate of oxygenic photosynthesis (GOP) as a function of irradiance in the uppermost layer of a biofilm formed by the LSS cyanobacterium (Fig. S4). The experimental data were fitted with the model of Eilers and Peeters for photosynthesis-over-irradiance-curves (PI-curves) by non-linear regression to determine the optimal light intensity (137 μmol photons m-2 s-1) and the biomass- dependent GOP at this optimal light intensity (GOPmax). The PI-curve measured for oxygenic photosynthesis exhibited the expected features (Fig. S5)—GOP increases linearly in an irradiance-dependent manner before saturation occurs. Normalization of all rates from each separate measurement to GOPmax revealed that the activity relative to GOPmax was highly reproducible and independent of the biomass in the surface layer. Photosynthesis in the presence of sulfide—anoxygenic photosynthesis Microsensors were used to determine the gross rate of anoxygenic photosynthesis (GAP) as a function of irradiance in the uppermost layer of a biofilm formed by the LSS cyanobacterium (Fig. S4). Anoxygenic photosynthesis was instantaneously detectable when sulfide was locally available within the mat (Fig. S5). This result was observed both with and without DCMU (Fig. S5). GOP vs. GAP Oxygenic and anoxygenic photosynthesis were never performed simultaneously during our measurements presumably due to inhibition of oxygenic photosynthesis caused by local H2S concentrations <1 μM. GOP only started to recover when the local H2S concentrations remained <1 μM for ~ 30 min (Fig. 4). Intriguingly, the rate of GOP recovery decreased with increasing light intensity (Fig. 4). Normalization of GAP to GOPmax revealed a consistent pattern of H2S- and light-dependent kinetics (Fig. 5). For instance, GAP increased with increasing H2S concentration until a light-dependent maximum was reached. The initial increase of GAP over low H2S concentrations and the 188 II. Manuscripts saturation effect at higher concentrations resembled Michaelis-Menten-like kinetics. The initial slope of the increase and the maximum GAP were however also light-dependent. During exposure to 137 μmol photons m-2 s-1, the light-dependent maximum was observed at ~42 μM H2S, followed by a pronounced decrease in GAP with increasing H2S concentration. This pronounced decrease was not observed during exposure to lower light intensities (Fig. 5). The maximum GAP (in electrons) at an H2S concentration of 42 μM was roughly two times higher than GOPmax during exposure to 136 and 180 μmol photons m-2 s-1 (Fig. 5; Fig. S5). In contrast, during exposure to 36 μmol photons m-2 s-1, GAP did not exceed the corresponding GOP. All patterns were strictly dependent on H2S concentration and were not affected by the temporal dynamics of exposure to sulfide (e.g. H2S dynamics in Fig. S5) or by the addition of DCMU (Fig. 5; Fig. S5). 100 10 I = 137 I = 23 8 75 (%) 6 max 50 M) 4 μ OP/OP S ( 2 25 2 H 0 0 024682224 time (h) Figure 4: Recovery of the volumetric gross rates of oxygenic photosynthesis after the depletion of H2S within the LSS isolate biofilm during exposure to 23 (open triangles) and 136 (closed triangles) ȝmol photons m-2 s-1. The lines represent the output of the simulation of the experimental data based on the model described by Fig. 7 and in Table S2. CO2 fixation rates during oxygenic and anoxygenic photosynthesis To confirm that the LSS isolate does not only deplete sulfide dependent on light, but is capable of primary productivity via anoxygenic photosynthesis, we performed 13C- bicarbonate studies in the absence and presence of DCMU (Fig. 6). In the presence of sulfide alone, the cells incorporated higher amounts of 13C-bicarbonate (12.9 (±1.61) μmol C assimilated mg dry weight-1) compared to cells which received no treatment (9.07 (±0.92) μmol C assimilated mg dry weight-1) and cells that received both sulfide and DCMU (5.7 (±0.71) μmol C assimilated mg dry weight-1). No 13C incorporation was observed in cells incubated in the dark. 189 5. A MODEL CYANOBACTERIUM FROM A PROTEROCOIC OCEAN ANALOG I = 137 200 I = 137, +DCMU 150 (% ) max 100 P/OP 50 I = 36 I = 36, +DCMU 0 0 255075100125 H2S ȝM) Two sulfide-oxidizing enzymes model Light-dependent SQR activity model Figure 5: Volumetric gross rates of anoxygenic photosynthesis normalized to the maximum electron transport rate of oxygenic photosynthesis at the optimal irradiance 137 ȝmol -2 -1 photons m s (GOPmax) dependent on H2S concentration measured at 36 and 137 ȝmol photons m-2 s-1 in the absence and presence of DCMU. The lines represent the output of the simulation of the experimental data based on the model described by Fig. 7 and in Table S2. Figure 6: Carbon assimilation by the LSS isolate. Error bars obtained from triplicate measurements. Discussion The Utility of Hopanoids as Biomarkers Hopanoids, which derive primarily from bacteria, have been found throughout the geologic record and with robust analyses of hopanoids in extant organisms and modern 190 II. Manuscripts environments, their presence may aid in interpreting geochemical conditions as well as provide some taxonomic resolution of microbial communities present at the time of deposition. The majority of sedimentary 2-methylhopanes are thought to derive from ancient shallow marine environments (Eigenbrode et al., 2008; Waldbauer et al., 2009), where oxygenic phototrophs would have been dominant. However, hopanoid production is not observed in pure cultures of the most abundant marine cyanobacteria (Talbot et al., 2008) and none of the sequenced cyanobacteria that contain the hpnP methylase gene were isolated from marine environments (Welander et al., 2010). C-2 methylated hopanoids are however common in environmental samples from modern marine environments (Talbot et al., 2008); evidence bolstered by recent analyses of Shark Bay, Australia stromatolites where the majority of partial hpnP sequences recovered were most closely related to cyanobacteria (Garby et al., 2012). In the moderately saline waters of the Little Salt Spring (~3 ppt), the seasonal red biofilm is a large source of BHPs and 2- Me-BHPs (anhydro-BHT and 2Me-anhydro-BHT) (data not shown). In the present study, BHP synthesis by the LSS isolate has been demonstrated, including those methylated at the C2 position. This is the first identified species of this clade to produce BHPs and another possible cyanobacterial source of C2 methyl hopanes in the rock record. Interestingly, the purple cyanobacterial mats in sunlit sinkholes of Lake Huron, closely related to P. autumnale, do not appear to synthesize 2Me-BHPs (Voorhies et al., 2012) despite colonizing a similar environment, i.e. low oxygen, sulfidic waters. Despite the observation that many bacteria that encode an hpnP homolog are plant-associated (Ricci et al., 2013), many modern environments where 2-MeBHPs have been recovered are micro-oxic to anoxic. These results, combined with the Little Salt Spring isolate presented here, further precipitate the need for model organisms more representative of early- evolving bacteria to understand the physiological role of 2Me-BHPs and the environmental parameters selecting for the synthesis of these lipids. Our data support recent conclusions that 2-MeBHPs cannot be utilized as taxonomic markers in modern or ancient environments. Cyanobacteria and Anoxygenic Photosynthesis Cyanobacteria from stably oxygenated environments are not typically exposed to sulfide and are extremely sensitive to sulfide exposure. In contrast, several adaptations are observed in species inhabiting sulfidic environments including resistance of PSII to sulfide and the ability to perform anoxygenic photosynthesis. Several species of 191 5. A MODEL CYANOBACTERIUM FROM A PROTEROCOIC OCEAN ANALOG cyanobacteria are capable of utilizing sulfide as an electron donor, performing anoxygenic photosynthesis using only PS I. This physiological strategy has been considered a relic of ancient cyanobacteria prior to the evolution of cyanobacteria (Oren at al., 1977). In addition to LSS, this activity has been observed in pure cultures of other members of Oscillatoriales, including Geitlerinema sp. PCC 9228 (formerly Oscillatoria limnetica 'Solar Lake'), isolated from sulfide-rich mats in Solar Lake, Israel (Cohen et al., 1975a; 1975b) and Phormidiaceae cyanobacterium SAG 31.92 (formerly Microcoleus chthonoplastes strain 11) (De Wit and van Gemerden, 1987) and has been implicated in in situ studies where members of the Oscillatoriales are abundant (i.e. Voorhies et al., 2012). And, while a number of Oscillatoriales spp. have been isolated from sulfidic environments, the ability to perform anoxygenic photosynthesis is not a conserved trait among these isolates and has been observed in phylogenetically diverse cyanobacteria. These observations are compounded by the observation that the Oscillatoriales, as well as other groups of cyanobacteria, are not polyphyletic (Ishida et al., 2006); multiple evolutionary lineages of cyanobacteria are capable of living in sulfidic environments (Miller and Bebout, 2004); and there are relatively few sequenced genomes of cyanobacteria available, especially of isolates from sulfidic, low oxygen habitats. As such, conserved phylogenetic markers which indicate the genetic capability to perform anoxygenic photosynthesis among cyanobacteria warrant further investigation in model organisms such as the LSS isolate. Genetic Requirements for Cyanobacterial Anoxygenic Photosynthesis SQR is the only enzyme that has a demonstrated role in anoxygenic photosynthesis in cyanoabacteria—in Geitlerinema sp. PCC 9228—where the enzyme catalyzes sulfide- dependent reduction of the plastoquinone (PQ) pool. SQR mediates electron transfer from sulfide to PQ without requiring any other proteins, suggesting it could be the only requirement for anoxygenic photosynthesis in cyanobacteria. For SQR sequences in general, there is only rough correlation between the topology of phylogenetic trees of 16S rRNA and SQR sequences from the same organism (Pham et al., 2008). SQR also serves other functions such as sulfide detoxification and may be involved in anaerobic respiration in cyanobacteria (Garlick et al., 1977; Oren and Shilo, 1979). Analysis of cyanobacterial sequences for which both 16S rRNA sequences and SQR are available—a relatively small data set (n=27)—did not reveal a correlation between topology of cyanoabcterial 16S rRNA sequences and SQR (Fig. S6). In addition, the SQR sequences 192 II. Manuscripts of the LSS isolate, Geitlerinema sp. PCC 9228—both capable of anoxygenic photosynthesis—are only distantly related (Fig. 1) and a number of marine spp., which presumably are not exposed to sulfidic conditions, lack SQR homologs (Miller and Bebout, 2004). Several lines of evidence suggest sulfide tolerance is a dynamic trait that can be gained or lost including the variable tolerance of PSII to sulfide (Miller and Bebout, 2004). The widespread taxonomic distribution of cyanobacteria capable of performing anoxygenic photosynthesis in the absence of detectable heritability of this trait also supports this hypothesis. Unfortunately, these observations do not discern if SQR, or more specifically, anoxgyenic photosynthesis, is an ancestral trait in cyanobacteria or acquired by horizontal gene transfer. It is still unclear if SQR is the only additional requirement for all cyanobacterial anoxygenic photosynthesis. One more drawback is the observation that many organisms encode more than one homolog of SQR. There are several examples of this among cyanobacteria (Fig. S2; FigS6) and could be true for both Geitlerinema sp. PCC 9228 whose genome has not been sequenced and the LSS isolate, for which we only present a draft genome. The thylakoid membrane-bound type A SQRs of Geitlerinema sp. PCC 9228 and A. halophytic exhibit high affinity for sulfide (Km = 8 μM and 20 μM, respectively) (Arieli et al., 1994; Bronstein et al., 2000). The LSS isolate genome encodes a type F SQR and while Km values for this type have not been reported, levels of SQRF mRNA in C. tepidum are regulated by sulfide (Chan et al., 2009) and this SQR is membrane-associated. The LSS draft genome does not encode other sulfur oxidizing enzymes such as flavotocytochrome c (FCC), dissimilatory sulfur reductase (Dsr) or sulfite:cytochrome c oxidoreductase. However, the taxonomic and physiological breadth of cyanobacteria has been poorly characterized, both genetically and biochemically, thus the presence of novel or unknown pathways of sulfide oxidation cannot be dismissed. Clearly, there is disparity between experimental observations of cyanobacterial anoxygenic photosynthesis and the characterization, availability and genomic sequencing of relevant cultured isolates. The LSS draft genome presented here supports the role of SQR in anoxygeinc photosynthesis among cyanobacteria but evolutionary and biochemical characterization are required to determine the genetic underpinnings of this physiology which could have played a key role in Earth’s redox evolution. The genomic data, in combination with the microsensor data presented here suggests a many factors including H2S concentration, irradiance, kinetics, and enzyme 193 5. A MODEL CYANOBACTERIUM FROM A PROTEROCOIC OCEAN ANALOG affinity play a role in anoxygenic photosynthetic activity in cyanobacteria. Based on our data, we present a model of this interplay (Fig. 7 and Table S2) and will discuss these aspects of the model in more detail below. f E II k k R E PSIId OEC:H2 S PSII PSII* kD k S1 kS2 S0 OEC ox SQR kAP kOP H2 O H2 S kO2 PQox PQred O2 OEC k PQ S0 “SO” kAP2 cyt cox H2 S cyt cred NADP+ PSI CO2 kCO2 f I E k CH O tot 2 NADPH Figure 7: Proposed model for the kinetic control of the redox reactions involved in oxygenic and anoxygenic photosynthesis by the LSS isolate. Dark gray represents components involved only in anoxygenic photosynthesis; light gray represents components involved in both oxygenic and anoxygenic photosynthesis; no color are those components that are involved only in oxygenic photosynthesis. E is the photon flux; PSII, PSII* and PSIId are photosystem II in the ground, excited and degraded/inactive state, respectively; OEC is the oxygen evolving complex. OECox is an intermediate formed during H2O oxidation that is inhibited by H2S. OEC:H2S is the inhibited form of this intermediate; SQR is the sulfide:quinone:oxidoreductase that couples the oxidation of sulfide to the reduction of the oxidized part of the plastoquinone pool (PQox), which yields zero- valent sulfur and reduced PQ (PQred). Cyt cox is representative of any oxidized intermediate electron transport chain component between PQ and photosystem I (PSI) that serves as the electron acceptor for an unidentified sulfide oxidase. The PSI reaction centre can receive electrons from the reduced intermediate (cyt cred). These electrons are used to reduce NADP+ to NADPH, which serves as the electron donor during CO2 fixation. Definitions of the process rates (ki) are given in Table S2. Inhibition of oxygenic photosynthesis Oxygenic photosynthesis by the LSS isolate was immediately inhibited by H2S concentrations >1μM (Fig. S5). Sulfide-dependent inhibition of photosynthesis has been observed in a number of cyanobacteria (Castenholz, 1977; Garlick et al., 1977; Oren et al., 1979; Cohen et al., 1986; Miller and Bebout, 2004). In typical cyanobacteria, the mechanism behind the inhibition is most likely a direct interaction of H2S with the oxygen evolving complex (OEC) in PSII (Cohen et al., 1986; Garcia-Pichel and Castenholz, 1990). In contrast, for metabolically versatile cyanobacteria capable of anoxygenic photosynthesis, it has been suggested that the inhibition of OP can also occur via changes in the photosystem stoichiometry (Garcia-Pichel and Castenholz, 1990). 194 II. Manuscripts Previous studies in cyanobacteria have indicated PS stoichiometry is linked to the redox poise of the plastoquinone pool (Murakami et al., 1997; El Bissati et al., 2000; Li and Sherman, 2000). According to this hypothesis, PSI is up-regulated during anoxygenic photosynthesis while PSII is degraded. This hypothesis is supported by fluorescence measurements indicating a decrease in the amount of PSII during anoxygenic photosynthetic growth and the observation of a delayed recovery of oxygenic photosynthesis after depletion of H2S (Cohen et al., 1986; Garcia-Pichel and Castenholz, 1990). A mechanism for PSII degradation is not known. In the LSS cyanobacterium isolate we observed a ~30 min lag between sulfide depletion and the onset of oxygenic photosynthesis (Fig. 4). The recovery rate after this onset increased with decreasing irradiance. The short timeline contradicts an H2S- dependent modulation of PSII levels wherein PSII is degraded in the presence of sulfide and induction of PSII transcription occurs as a result of H2S depletion. Instead, we hypothesize that the delay in onset of recovery and the light-dependent rate of recovery of OP are both caused by an interplay between the kinetics of OEC inhibition and photoinhibition reactions based on the model depicted in Fig. 7. Specifically, the light- independent ~30 min delay of OP recovery can be explained by assuming that the inhibition of the OEC in the LSS cyanobacterium only slowly relaxes after H2S depletion—the back reaction to an active OEC (kS2 in Fig. 7 and Table S2) is slow. The excitation energy harvested in PSII cannot be used for the photochemical reactions after inhibition of the OEC. Thus, even at very low light intensities, rates of photoinhibition and degradation of PSII components (d1 subunit) (kD in Fig. 7 and Table S2) that are sufficiently high enough to exceed the rate of PSII repair (kR in Fig. 7 and Table S2) are plausible. Notably, this scenario does not invoke H2S-driven degradation of PSII. Instead, the rate of degradation will depend on light intensity and the level of OEC inhibition. Therefore, as soon as non-inhibited OEC is available, oxygenic photosynthesis can continue. If the light intensity is high, the rate of photoinhibition will still substantially exceed the rate of PSII repair, causing slow recovery. In contrast, low light intensities will allow for a rapid decrease in photoinhibition rates and consequently a fast recovery. We have implemented this concept into a numerical model (using the deSolve package in R (Soetaert et al., 2010; cran.r-project.org) based on previous work on the kinetics of inhibition by H2S (Klatt et al., 2015b) (Fig. 7 and Table S2), the results of which are in remarkable agreement with the experimental data (see lines in Fig. 4). These results 195 5. A MODEL CYANOBACTERIUM FROM A PROTEROCOIC OCEAN ANALOG support the hypothesis that the kinetics of OEC inhibition and photoinhibition reactions affect the recovery of GOP, a model that warrants further testing. 250 CO fixation rate Photoinhibition Light limitation 2 limitation 200 150 (%) max 100 P/OP 50 OP AP 0 0 50 100 150 200 250 Irradiance (μmol photons m-2 s-1) Figure 8: Volumetric gross rates of oxygenic and anoxygenic photosynthesis described in Fig. S5 with lines representing the output of the simulation of the experimental data based on the model described by Fig. 7 and in Table S2. The resultant PI-curve is divided into operational irradiance ranges based on the plausible rate limiting steps of the photosynthetic electron transport during oxygenic photosynthesis. Anoxygenic Photosynthesis Anoxygenic photosynthesis attributed to cyanobacteria has been observed in several sulfidic environments including benthic mats in Solar Lake (Cohen et al., 1975a; 1975b) and sulfidic sinkholes in Lake Huron (Voorhies et al., 2012) yet the mechanisms and regulation of this activity is poorly understood in pure cultures. Geitlerinema sp. PCC 9228 oxidizes sulfide to elemental sulfur which is deposited extracellularly (Cohen et al., 1975a; Cohen et al., 1975b; Castenholz and Utkilen, 1984). In contrast, Phormidiaceae cyanobacterium SAG 31.92 produces thiosulfate stoichiometrically from sulfide during anoxygenic photosynthesis (De Wit and van Gemerden, 1987). In Geitlerinema sp. PCC 9228, the shift from oxygenic to anoxygenic photosynthesis occurs after a 2 h incubation period in the presence of light and sulfide (Cohen et al., 1975a; 1975b; Oren and Padan, 1978) whereas variable induction times were observed in thermophilic strains of Oscillatoria amphigranulata (Garcia-Pichel and Castenholz, 1990). Here, we demonstrated sulfide-dependent anoxygenic photosynthesis in the LSS isolate immediately upon addition of sulfide and confirmed these results in the presence of sulfide and DCMU (Fig. S5). The pre-incubation time in Geitlerinema sp. PCC 9228 is presumably necessary for protein synthesis where SQR is the inducible factor (Oren and Padan, 1978) while the switch back to oxygenic photosynthesis requires no time. The 196 II. Manuscripts rapid switch from oxygenic to anoxygenic in the LSS isolate measured in the microsensor-based approach is supported by observations of 13C incorporation in DMCU and sulfide treated cells during the 2 h incubation time (and no pre-incubation)(Fig. 6). We observed elevated levels of 13C incorporation compared to the controls (light) and the incubations that received both DCMU and sulfide. The lower level of C assimilated in the incubation experiments in the presence of DCMU and sulfide suggest this treatment becomes limited with respect to electron donor. We hypothesize this difference represents high rates of anoxygenic photosynthesis followed by oxygenic photosynthesis in the sulfide treatment whereas the DCMU and sulfide treatment was not able to perform oxygenic photosynthesis following depletion of sulfide by anoxygenic photosynthesis. The rates of anoxygenic photosynthesis in the LSS isolate seem to be controlled by a complex interplay between irradiance and H2S concentration because: (i) GAP increases dependent on H2S concentrations until a light-dependent maximum at ~42 μM is reached and the slope of this increase is dependent on light (Fig. 5); (ii) the anoxygenic photosynthetic electron transport rate at ~42 μM is two times that of the oxygenic electron transport rate at saturating light intensities (Fig. 5; Fig. S5); (iii) GAP decreases at H2S concentrations greater than ~42 μM when exposed to high light (Fig. 5). SQR donates electrons derived from H2S oxidation into the plastoquinone pool, the oxidation of which is governed by light harvested in PSI. This explains the H2S- dependent increase of GAP and the light-dependent maximum GAP in the LSS isolate (Fig. 5). Specifically, H2S oxidation is expected to proceed at a rate that depends on the affinity of SQR to H2S and the oxidized part of the PQ pool (kAP; Fig. 7 and Table S2). The light energy harvested in PSI dictates the maximum electron transport rate in the irradiance range below saturation (ktot; Fig. 7 and Table S2) which also represents the maximum GAP. Thus maximum GAP is independent of the maximum H2S oxidation rate by SQR. Still, this does not explain the light-dependent slope of the increase in GAP with increasing H2S concentration (i.e. Fig. 5). Instead, this effect can be best explained by three plausible hypotheses: (a) The LSS isolate is equipped with at least two types of SQR characterized by different affinities towards H2S and the activity and expression of these enzymes is dependent on irradiance. We identified a single gene encoding an SQR in the draft genome and attempts to amplify more that one SQR-encoding gene have been unsuccessful, therefore we consider this hypothesis unlikely. (b) The LSS isolate possesses a single SQR and the expression of this SQR is dependent on irradiance. For 197 5. A MODEL CYANOBACTERIUM FROM A PROTEROCOIC OCEAN ANALOG instance, if, at higher light intensities the transcription of SQR is up-regulated, the result would be more active SQRs and an increased maximum rate of H2S oxidation (Ȟmax in kAP_B; Fig. 7 and Table S2). This would result in a steeper initial slope in GAP like that observed in Fig. 5. (c) There are two types of sulfide oxidizing enzymes with different affinities for H2S that donate electrons into the electron transport chain at different levels. A SQR and a FCC would be most plausible because they would theoretically fulfill these requirements (Griesbeck et al., 2000). No FCC-like enzyme was encoded in the draft genome, but the second enzyme might be an unidentified sulfide oxidase (see “SO” and kAP2 in Fig. 7 and Table S2). Notably, for this scenario it is not necessary to assume a direct effect of irradiance on enzyme activity or regulation of transcription. The PI-curve measured for oxygenic photosynthesis exhibits the expected features (Fig. S5; Fig. 8)—GOP increases linearly in an irradiance-dependent manner before saturation occurs. The saturation effect at high light intensities is most likely the result of rate limitation during CO2 fixation (Sukenik et al., 1987; Cardol et al., 2011). At even higher light intensities GOP decreases again (Fig. 8) presumably due to photoinhibition reactions in PSII that lead to degradation of PSII components (Shipton and Barber, 1991; Vass et al., 1992) (see above). Intriguingly, the maximum GAP at 42 μM H2S concentration was nearly double the GOP in the range of light intensity where rate limitation during CO2 fixation would be expected (Fig. 8); and, in contrast, the maximum GAP at lower light intensities did not considerably exceed the rate of oxygenic photosynthetic electron transport. It has been shown previously that H2S enhances the activity of RuBisCO in obligate oxygenic phototrophs (Chen et al., 2011; Klatt et al, 2015a). We hypothesize that H2S has a similar regulatory function in the LSS isolate and that the increase in electron transport rate is further supported by excitation energy transfer from PSII to PSI, which is regulated by the redox state of the PQ pool (kCO2; Fig. 7 and Table S2). At H2S concentrations above ~42 μM GAP decreased and the decrease was most pronounced at higher light intensities. To our knowledge, inhibition of the rate of anoxygenic photosynthesis by H2S concentration has not been reported previously. We hypothesize that H2S negatively affects a reaction downstream of PSI, which only plays a role when the maximum GAP is not exclusively controlled by irradiance, i.e., at light intensities where the rate of CO2 fixation is the limits the overall electron transport rate (e.g., kCO2 in Fig. 7 and Table S2). 198 II. Manuscripts Based on the experimental and genomic data presented here, we implemented the conceptual framework derived above (Fig. 7) in our numerical model of the photosynthetic electron transport chain reactions. Also, we separately tested our 2 most plausible hypotheses—two sulfide oxidizing enzymes (SQR and “SO”) or light-dependent activity of SQR. Both models gave remarkably good agreement with the experimental data (Fig. 5), precluding our ability to discern between these hypotheses. Future experiments to address these questions, including PS stoichiometry and the light and H2S dependence of anoxygenic photosynthesis are critical to understanding underlying intricacies of this physiology. Implications for Earth’s Redox Evolution Anoxygenic photosynthesis in cyanobacteria has been suggested to represent an ancient relic carried over from the early evolution of oxygenic photosynthesis (Padan, 1989). A proposed role of ancient cyanobacteria capable of this shift in physiology is in the protracted delay in oxygenation of the atmosphere following the Great Oxidation Event (Johnston et al., 2009). If physiologically versatile cyanobacteria contributed to maintaining low levels of oxygen throughout Earth’s middle age by performing anoxygenic photosynthesis, the timescale of the switch between PSI- and PSII-dependent photosynthesis (and back) could have been a competitive advantage over other photolithotrophs. This advantage would be two-fold considering oxygen produced by PSII-dependent photosynthesis would oxidize sulfide present in the photic zone. Assuming some form of SQR was utilized by ancient cyanobacteria, affinity of the enzyme for sulfide would also play a key role in determining the ecological success of these versatile cyanobacteria. Alternatively, the presence of SQR and primary productivity via PSI-dependent anoyxgenic photosynthesis in select cyanobacteria could reflect niche-specific adaptations acquired by horizontal gene transfer. However, this study presents several lines of evidence suggesting metabolically versatile cyanobacteria could have persisted in euxinic oceans during Earth’s middle age. For instance, rates of anoxygenic photosynthesis observed in the LSS isolate depend on both H2S concentration and light and, under the correct conditions, rates of PSI-dependent photosynthesis are over twice that of oxygenic photosynthesis. Our data suggest that PSII-dependent photosynthesis in LSS can only be initiated upon depletion of H2S and at higher light intensity; however, this could be a reflection of the conditions tested here. Further experiments are required to determine if these observations are consistent with in situ 199 5. A MODEL CYANOBACTERIUM FROM A PROTEROCOIC OCEAN ANALOG activity, but these data support our current knowledge of cyanobacteria in sulfidic sunlit environments where they co-occur with anoxygenic phototrophs, but preferentially occupy positions that receive the most light (i.e. the top layer of microbial mats). This same niche in the Proterozoic oceans, well-lit and sulfidic, could have harbored metabolically versatile cyanobacteria as well as other photolithotrophs, similar to stratified systems observed on Earth today where elevated tolerance to both light and oxygen presumably result in cyanobacteria dominating the upper levels of these niches. 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Manuscripts ANOXYGENIC PHOTOSYNTHESIS AND 2-METHYL HOPANOID PRODUCTION: A MODEL CYANOBACTERIUM ISOLATED FROM A PROTEROZOIC OCEAN ANALOG Trinity L. Hamilton, Judith M. Klatt, Laurence M. Bird, Katherine H. Freeman, Dirk de Beer, and Jennifer L. Macalady Supplementary Information 207 5. A MODEL CYANOBACTERIUM FROM A PROTEROCOIC OCEAN ANALOG Figure S1.Maximum likelihood phylogeny of cyanobacteria inferred from full-length 16S rRNA sequences from from complete to nearly complete genomes. Bootstrap support values >90 based on 1000 bootstrap samplings are shown for each node. Accession numbers are given in parentheses. 208 II. Manuscripts Figure S2. Maximum likelihood phylogeny of Sqr sequences. Classification of Sqr othologs is based on Gregersen et al., 2011. Accession numbers are given in parentheses. Bootstrap support values >90 based on 1000 bootstrap samplings are shown for each node. 209 5. A MODEL CYANOBACTERIUM FROM A PROTEROCOIC OCEAN ANALOG Figure S3. Structures of hopanoids made by the LSS isolate. Figure S4. Volumetric gross rates of oxygenic and anoxygenic photosynthesis normalized to the maximum electron transport rate of oxygenic photosynthesis at the optimal irradiance of 137 ȝmol photons m-2 s-1 (GOPmax) dependent on incident irradiance. Rates were determined in the uppermost layer of a mat formed by the LSS isolate. 210 II. Manuscripts Figure S5. Volumetric gross rates of oxygenic and anoxygenic photosynthesis, and local H2S concentration over time at 137 ȝmol photons m-2 s-1 incident irradiance in the absence (A) and presence (B) of DCMU. The measurements were performed in the uppermost layer of two different LSS isolate mat samples. Arrows indicate addition of DCMU or H2S. All rates were converted into photosynthetic electron transport rates. Figure S6. Maximum likelihood phylogeny of all cyanobacteria for which both 16S rRNA sequences (A) and Sqr sequences (B) are publicly available. Accession numbers are given in parentheses. Bootstrap support values >90 based on 1000 bootstrap samplings are shown for each node. 211 5. A MODEL CYANOBACTERIUM FROM A PROTEROCOIC OCEAN ANALOG 212 II. Manuscripts Table S2: Definition of the rate laws governing the redox reactions shown in Figure 7. Expression Description PSII The rate of generation of an excited catalytic Chl a k E f II E dimer in photosystem II (PSII*). It depends on PSII K E irradiance (E), the availability of the ground state Chl a in PSII (PSII) and the absorbance cross-section factor fII that describes the efficiency of conversion of the externally available photon flux (E) into a volumetric rate of excitation. OEC PQ ox The rate of PQ reduction by PSII, i.e., the rate of kOP Q OP PSII * oxygenic photosynthetic electron transport. It OEC K PQ K OEC ox PSII depends on the availability of the excited catalytic Chl a dimer in PSII (PSII*), non-inhibited oxygen evolving complex (OEC) and oxidized plastoquinone (PQox). This process results in the formation of a highly reactive oxidized oxygen evolving complex (OECox), reduced plastoquinone (PQred) and regeneration of ground state Chl a in PSII (PSII). OEC The rate of O2 release from the OEC, which depends k Q ox O2 O2 on the availability of oxidized OEC (OECox). OECox K O2 E PSII * The rate of PSII degradation by photoinhibition. It k D Q D depends on the availability of the excited catalytic E K PSII * K D1 D2 Chl a dimer in PSII (PSII*) and the irradiance (E). To account for light-dependent efficiency of photoinhibition (REF) the rate saturates at high light intensities. PSIId The rate of repair of the partially degraded, non- k R Q R active PSII (PSIId). PSIId K R The rate of H S oxidation coupled to PQ reduction [H 2S] PQ ox 2 ox k AP = vmax by SQR, i.e., the rate of anoxygenic photosynthetic K [H S] K PQ M 2 SQR ox electron transport. This process results in the formation of zero-valent sulfur and PQred. Assuming that a hypothetical additional sulfide [H 2S] PQ ox k AP _ B = vmaxD oxidizing enzyme (see “SO” in k ) exists or that K [H S] K PQ AP2 M 2 SQR ox two SQRs exist that are expressed dependent on the E light intensity (not shown), this rate is kAP and D (1 D ) depends exclusively on the H S concentration ([H S]) 0 E K 2 2 D and the availability of PQox. Assuming that the activity of SQR is directly regulated by the light intensity the maximum rate of H2S oxidation this rate is kAP_B. In this rate Ȟmax, additionally depends on a factor Į that increases with irradiance (E). The rate of H S oxidation coupled to the reduction of [H 2S] cyt c ox 2 k AP2 = vmax 2 another electron transport chain component, such as K M 2 [H 2S] K"SO" cyt c ox oxidized cytochrome c (cyt cox), by a hypothetical sulfide oxidizing enzyme “SO”. This rate is only included in the model when assuming that the activity of SQR is not directly regulated by the light intensity (see description of kAP and kAP_B). 213 5. A MODEL CYANOBACTERIUM FROM A PROTEROCOIC OCEAN ANALOG Table S2 (continued) cyt cox PQ red The rate of PQred oxidation coupled to the reduction k PQ Q C of any electron transport chain component, such as cyt cox K C1 PQ red K PQ oxidized cytochrome c (cyt cox), that can serve as the electron acceptor for the hypothetical sulfide oxidizing enzyme “SO”. This process results in the reformation of PQox, which is available again for the reduction by SQR or PSII. The rate of NADP+ reduction coupled to the NADP cyt c red ktot f I E E oxidation of an electron transport component (cyt NADP K cyt c K N red C 2 cred). This rate depends on the availability of cyt cred, + -2 -1 NADP , irradiance (IPSI in μmol photons m s ), and PQ red E (1 E 0 f II ) the absorbance cross-section factor fI that describes PQ tot the efficiency of conversion of the externally available photon flux (E) into a volumetric rate of excitation in PSI. fI is increased by excitation energy transfer from PSII to PSI, which is described by the transfer factor ȕ. ȕ depends on the redox state of the plastoquinone pool where PQred refers to the reduced part of the total PQ pool (PQtot). NADPH The rate of CO2 fixation coupled to NADPH kCO2 Q CO2 J G oxidation, which depends on the maximum rate of NADPH K CO2 CO2 reduction (ȞCO2) and the availability of NADPH. Ȟ is enhanced when H S available, which is [H 2S] CO2 2 J 1 J 0 described by the factor Ȗ. At high H2S concentrations [H 2S] K E (KE< 214 II. Manuscripts 6. ANOXYGENIC PHOTOSYNTHESIS CONTROLS OXYGENIC PHOTOSYNTHESIS IN A CYANOBACTERIUM FROM A SULFIDIC SPRING Judith M. Klatt1*, Mohammad A. A. Al-Najjar1,2, Pelin Yilmaz1, Gaute Lavik1, Dirk de Beer1, Lubos Polerecky1,3 1Max Planck Institute for Marine Microbiology, Celsiusstr.1, 28359 Bremen, Germany 2 Red Sea Research Center, KAUST, Thuwal, Saudi Arabia 3Department of Earth Sciences – Geochemistry, Faculty of Geosciences, Utrecht University, Princetonplein 9, 3584 CC, Utrecht, The Netherlands Published in Applied and Environmental Microbiology *Address correspondence to Judith M. Klatt, [email protected] 215 6. COMPETITION BETWEEN PHOTOSYNTHETIC MODES Abstract Before the Earth’s complete oxygenation (0.58–0.55 Ga ago), the photic zone of the Proterozoic oceans was probably redox stratified and characterized by an upper slightly aerobic nutrient-limited layer above a light-limited layer that tended towards euxinia. In such oceans, cyanobacteria capable of both oxygenic and sulfide-driven anoxygenic photosynthesis played a fundamental role in the global carbon, oxygen and sulfur cycle. We isolated a cyanobacterium, Pseudanabaena FS39, in which this versatility is still conserved, and show that the transition between the two photosynthetic modes follows a surprisingly simple kinetic regulation controlled by its affinity to H2S. Specifically, oxygenic photosynthesis is only performed additionally to anoxygenic photosynthesis when H2S becomes limiting and decreases below a threshold, which increases predictably with the available ambient light. The carbon-based growth rates during oxygenic and anoxygenic photosynthesis were similar. However, Pseudanabaena FS39 additionally - assimilated NO3 during anoxygenic photosynthesis. Thus, the transition between anoxygenic and oxygenic photosynthesis was accompanied by a shift of the C:N ratio of the total bulk biomass. These mechanisms offer new insights into how versatile cyanobacteria might have promoted oxygenic photosynthesis and total primary productivity despite nutrient limitation in the oxic photic zone in the mid-Proterozoic oceans, a key step that enabled complete oxygenation of our planet and subsequent diversification of life. Introduction Oxygenic photosynthesis (P) couples the power of two photosystems (PSI and PSII) for the extraction of electrons from water (Eq. 1) to reduce CO2. - 2 H2O Æ O2 + 4 e . (1) Nowadays, this metabolism is driving the major part of Earth´s ecosystems by providing its products oxygen and organic carbon to other organisms. However, oxygenic P has not always been the dominant photosynthetic process. From its evolution, which occurred possibly more than 3 billion years (Ga) ago (1), until its final success in oxygenating the atmosphere and biosphere (0.58–0.55 Ga ago), oxygenic P always had to compete with anoxygenic P, which uses alternative reduced electron donors, such as hydrogen sulfide (Eq. 2) (2). 0 - H2S Æ S + 2 e . (2) 216 II. Manuscripts Intriguingly, cyanobacteria could majorly contribute to primary productivity by both oxygenic P and anoxygenic P for billions of years (2), and this photosynthetic versatility was probably the key to their success in a reduced world. The balance between oxygenic P and anoxygenic P was plausibly dependent on the environmental conditions, e.g., the light, nutrient and H2S gradients. An understanding of how and why the global balance between oxygenic and anoxygenic P has changed on a geological timescale is therefore tightly connected to understanding (i) the regulation of the switch between the two photosynthetic modes dependent on relevant environmental conditions and (ii) the C- and N-based growth rates accompanying the respective photosynthetic mode. This can only be achieved by studying contemporary model organisms. It has been known since the 1970s that few extant cyanobacteria are still able to switch between anoxygenic and oxygenic P (3). Such cyanobacteria are presently exclusively found in systems where they are regularly exposed to sulfide, such as phototrophic microbial mats (4). So far, several factors were identified that appear to play a regulatory role on the partitioning between oxygenic and anoxygenic P: (i) H2S concentration, which is connected to both the toxicity of H2S for photosystem II (5) and to the concentration-dependent activity of the sulfide:quinone:oxidoreductase (SQR) that drives H2S oxidation in anoxygenic P (6, 7), (ii) exposure time to sulfide and light, because the de-novo synthesis of SQR first has to be induced (8), and (iii) the spectral quality of the light, because anoxygenic P is exclusively driven by PSI, while oxygenic P depends on PSI and PSII, both of which have different absorption spectra (9). However, little progress has been made on the details of the regulation of the transition between anoxygenic and oxygenic P in versatile cyanobacteria. In this study we investigated how the transition between anoxygenic and oxygenic P is regulated in cyanobacteria that are capable of simultaneous anoxygenic and oxygenic photosynthesis. Specifically, we isolated a mono-cyanobacterial culture of Pseudanabaena sp. from thin microbial mats forming in a cold sulfidic spring in Frasassi, Italy (10), and quantified its growth rate and nutrient requirements during the two photosynthetic modes.. Our overall aim was to gain insights into possible selective advantages that versatile cyanobacteria might have had in a redox-stratified Proterozoic ocean and to identify external parameters that might have promoted the success of oxygenic P. 217 6. COMPETITION BETWEEN PHOTOSYNTHETIC MODES Materials and Methods Experimental setup The experimental batch bioreactor without headspace (Fig. S1) was constructed using a 250 mL round-bottom flask with a ring-joint flange (Duran, Germany). It was closed with a custom-made lid through which sensors for real-time monitoring of O2, pH and variable fluorescence were inserted. Tightness was achieved by using O-rings and polyurethane- based sealing material (2K-PUR, Kemper System, Germany). The bioreactor was additionally equipped with two ports for injection, subsampling and degassing. A cut off glass syringe (100 mL) was used as an extension of the bioreactor volume to compensate for the injected/subsampled volume. Subsampling therefore did not lead to dilution or other disturbance of the culture and its activity. Turbulent mixing was achieved by using two magnet-filled glass bars simultaneously. Temperature was kept constant (15°C) by submerging the bioreactor in a flow-through water bath connected to a large temperature- controlled water reservoir. Illumination was provided by red (Ȝmax = 690 nm) and orange (Ȝmax = 590 nm) light emitting diodes (H2A1 series, Roithner-Lasertechnik, Austria) arranged circularly around the transparent water bath and the glass syringe (Fig. S1). Photon flux in the centre of the medium-filled bioreactor was measured with a fiber-optic scalar irradiance microprobe (11) connected to a spectrometer (USB4000, Ocean Optics, USA) and calibrated against a scalar irradiance sensor connected to a LI-250A light meter (LI-COR Biosciences GmbH, Germany). The current through the diodes was adjusted such that the culture was always exposed to a mixture of half the intensity of red light and half the intensity of orange light. Vertical and horizontal movement of the light-sensors during the calibration confirmed that the light was distributed homogeneously in the bioreactor. Experimental protocol The studied filamentous cyanobacterium strain FS39 was isolated and cultivated as described in the Supplement. When the culture reached a mid-exponential growth phase - - (Fig. S3), it was transferred into the bioreactor and the NO3 and HCO3 concentrations - - were readjusted to their initial values (3mM NO3 and 8 mM HCO3 ). Subsequently the bioreactor was tightly closed, and the culture was incubated at specific light conditions. When anoxygenic P was measured, H2S was additionally injected to the bioreactor (final concentration 0.5–10 mM). For selected incubations, the PSII inhibitor DCMU (3-(3,4- Dichlorophenyl)-1,1-dimethylurea; dissolved in ethanol) was added to a final 218 II. Manuscripts concentration of 5 μM (12). During all incubations, O2 concentrations, pH and variable fluorescence were monitored every 5–60 s using sensors inserted inside the bioreactor, while total sulfide concentrations (H2S) were analyzed in sub-samples collected every 13 15 10-20 min. During selected incubations, C-labeled HCO3 (~10% label) and N-labeled - NO3 (~20% label) were additionally injected to the bioreactor to enable quantification of inorganic carbon and nitrate uptake rates into biomass. Prior to each culture incubation, culture-free medium was incubated for at least 48 h to confirm tightness of the bioreactor and to determine rates of oxygen and sulfide removal by abiotic sulfide oxidation. Analytical procedures Concentration of dissolved O2 was measured using an O2 microelectrode (tip diameter 100-250 μm). pH was measured with a LIX-microsensor (tip diameter >50 μm) with a double junction reference electrode (REF251, Radiometer Analytical, France) and additionally with a standard laboratory pH probe (InLab Routine-L, Mettler Toledo, Switzerland). Microsensors were built and calibrated as previously described (13, 14). Total sulfide concentration (H2S) was determined according to Cline (15). The corresponding concentration of H2S was then calculated from this H2S according to Millero (16) using the pK value of 7.14 and the measured pH. Ammonium concentration in filtered and frozen samples was determined using flow injection analysis (17). 13 15 - Assimilation of C-HCO3 and N-NO3 into biomass was determined as previously described (18). Sub-samples were collected from the reactor and filtered through a pre-combusted GF/F filter placed in a syringe filter unit. The filtrate was fixed 13 12 with HgCl2 and later used to determine the C/ C ratio on a gas chromatography-isotope ratio monitoring mass spectrometry (VG Optima, Micromass, Manchester, UK) (18). To 15 14 - determine the N/ N ratio in the filtrate, NO3 concentrations just before and directly 15 - after the addition of N-NO3 were quantified using an NOx analyzer equipped with a chemoluminescence detector (Model CLD 66, Eco Physics, Switzerland) (19). The 13C/12C and 15N/14N ratios in the biomass on the GF/F filters were determined using an automated elemental analyser (Thermo Flash EA, 1112 Series) coupled to a Delta Plus Advantage mass spectrometer (Finnigan Deltaplus XP, Thermo Scientific) after freeze- drying (18). 219 6. COMPETITION BETWEEN PHOTOSYNTHETIC MODES Quantification of rates Rates of oxygenic and anoxygenic P were calculated from the measured concentrations of O2 and H2S as OxyP = d[O2]/dt and AnoxyP = -d[H2S]/dt, respectively. The CO2 and - NO3 assimilation rates were calculated as the rate of increase in the isotopic labeling of the biomass corrected for the labeling of the inorganic substrate in the medium (18). All rates were corrected for growth of the culture by using a growth factor derived from the overall increase of particulate organic carbon, as determined from mass spectrometry measurements. Results and discussion Pseudanabaena FS39 Phylogenetic analysis revealed that the 16S rRNA sequence of the studied cyanobacterium clustered with other Pseudanabaena spp. (Fig. S4). Taking additionally into account its morphology (Fig. S2), we decided upon the name Pseudanabaena FS39. Pseudanabaena FS39 is capable of exclusive oxygenic P, simultaneous anoxygenic and oxygenic, and exclusive anoxygenic P (see below). It can perform anoxygenic P at total sulfide (H2S) concentrations of at least 10 mM. We amplified a sqr-like gene (data not shown), which indicated that the initial H2S oxidation step of anoxygenic P in this strain is catalyzed by SQR, similar to Geitlerinema sp. PCC 9228 (formerly Oscillatoria limentica), the cyanobacterium that has been most intensively studied concerning its ability to perform anoxygenic P (7). Environmental relevance of the induction time for anoxygenic P In microbial mats local light, [H2S] and nutrient availability generally fluctuate dramatically over a diurnal cycle and cyanobacteria are expected to perform anoxygenic P in the early mornings when sulfide is abundant in the photic zone (4). In this context it appears paradoxical that cyanobacteria require an induction time of 2–3 hours before anoxygenic P starts (8). Also in Pseudanabaena FS39 anoxygenic P first had to be induced after cultivation in the absence of sulfide, and only occurred after ~2 h of exposure to light and sulfide (Fig. 1). However, subsequent injections of sulfide within 36 h of non-sulfidic or dark conditions resulted in an instant start of anoxygenic P, that is, the induction lag-phase disappeared (Fig. 1). Thus, the induction time is not of importance in 220 II. Manuscripts the environment if the organism is exposed to light and sulfide at least once within a diurnal cycle dark light dark light 3500 3000 H S injected 2500 2 (μM) S 2000 2 H 2S injected no delay H 1500 Σ ~ 2 h delay 1000 500 0 0 2 14 16 18 20 22 24 26 28 40 42 44 46 48 50 time (h) Figure 1: Concentration of total sulfide (H2S) in the bioreactor with Pseudanabaena str. FS39 as a function of time. After the initial injection of sulfide, the culture had to be exposed for ~2 h to light for anoxygenic photosynthesis (i.e., H2S consumption) to start. When the cyanobacteria were left exposed to sulfide in the dark for more than 18 h, no induction time was required and anoxygenic photosynthesis started immediately after the light was switched on. Anoxygenic P in Pseudanabaena FS39 is controlled by H2S concentration and light intensity To be able to quantify the rates of exclusive anoxygenic P, we inhibited oxygenic P by using DCMU (Fig. S7). The measurements in this experiment were done at two light intensities, 200 and 50 μmol photons m-2 s-1 (Fig. 2). Unexpectedly, the rates of anoxygenic P (anoxyP) were either exclusively determined by light or exclusively by the H2S concentration ([H2S]) within the boundaries of two distinct [H2S] ranges operationally denoted as A and B (Fig. 2). In the concentration range A, i.e., at low [H2S], the rates of anoxygenic P followed Michaelis-Menten-like (MM) enzyme kinetics with the sulfide species H2S (not H2S) as the substrate and were therefore light-independent (Fig. 2). Specifically, the apparent affinity constant was always KM = 38 μM (+/– 1.2 μM; N=8) (e.g., Fig. 2 and 3). Upon transition to the [H2S] range B anoxyP abruptly stopped increasing and remained constant at a maximum rate AnoxyPmax (Fig. 2). The threshold [H2S], above which transition from range A to B occurred, as well as the value of AnoxyPmax increased with light intensity. We propose that in the operational [H2S] range A anoxyP is determined by the activity of sulfide:quinone:oxidoreductase (SQR) (kH2S in Fig. 4). This enzyme catalyzes the oxidation of H2S to zero-valent sulfur coupled to the reduction of the plastoquinone (PQ) pool. This reaction is light-independent (8). The re-oxidation of the PQ pool is 221 6. COMPETITION BETWEEN PHOTOSYNTHETIC MODES indirectly governed by light, because light energy harvested in PSI drives the overall electron transport. This makes PSI the “bottleneck” of electron transport before the onset of light saturation effects downstream of PSI. We therefore suggest that AnoxyPmax reflects the maximum rate of electron flow through PSI (kPSI), which is dictated by the light quanta harvested by this photosystem. Thus, light-independent SQR enzyme kinetics seem to control the rate of anoxygenic P at low [H2S] (range A, Fig. 2), while light intensity determines the upper limit (AnoxyPmax) of the process at higher [H2S] (range B). 500 A I = 200 400 I = 200 + DCMU ) -1 h S 2 300 [H S] range A [H S] range B 2 2 H Σ (μM 200 I = 5 0 oxyP I = 50 + DCMU 100 An [H S] 0 2 thr 0024006 80 100 H2S(μM) 500 B 400 ) I=200 -1 h S 300 2 K K PSII/ SQR: H Σ 1000 100 200 (μM 10 1 oxyP 100 I=50 An 0 0 20406080100 H S(μM) 2 Figure 2: The rates of anoxygenic photosynthesis as a function of H2S concentration. (A) Measured rates at illumination with two different light intensities (I=50 μmol photon m-2 s-1 or I=200 μmol photon m-2 s-1) in the absence and presence of DCMU. Rates below the light- dependent H2S concentration threshold (i.e., in [H2S] range A) were fitted with the Michaelis- 2 Menten model for enzyme kinetics by non-linear regression (thick line) (KM = 39.1 μM, R = 0.998). Horizontal dashed lines indicate AnoxyPmax at the respective light intensity. The corresponding temporal dynamics of H2S and O2 are shown in Supplemental Figure S7. (B) Simulation of anoxygenic photosynthesis in Pseudanabaena str. FS39 under the same light conditions, based on Fig. 4 and Table 1. The use of different ratios of KPSII/KSQR (Table 1) revealed that KPSII has to be at least two orders of magnitude greater than KSQR to achieve rates of anoxyP that are not significantly affected by the rates of oxyP. 222 II. Manuscripts 500 A1 A2 A3 B 400 ) 1 - h - 300 Me (μ 200 P OxyP 100 AnoxyP 0 --1 CO NO - 0.20 500 2 3 Σ(CO ,NO -) 400 2 3 0.19 300 N:C 200 0.18 100 0.17 0 0 1 2310 20 30 40 50 60 assimilation rate (μM e h ) H S(μM) 2 Figure 3: Rates of anoxygenic (anoxyP) and oxygenic (oxyP) photosynthesis (upper panel), and nitrate and carbon dioxide assimilation rates and the N:C ratio of the bulk biomass (lower panel) -2 -1 as a function of H2S concentration during exposure to 200 μmol photons m s . Modeled rates (based on Michaelis-Menten kinetics; KM = 37.1 μM) are represented by the thick lines. Rates - -1 -1 were converted to μM e h as follows: oxygenic photosynthesis in μM O2 h was multiplied by 4 -1 (Eq. 1), anoxygenic photosynthesis in μM H2S h was multiplied by 2 (Eq. 2), rates of CO2 and - NO3 assimilation were multiplied by 4 and 8, respectively. Annotations on top indicate the operational [H2S] ranges (A1, A2, A3, B) of the complex switch between the photosynthetic modes. Further examples are shown in the Supplemental Information 5. Oxygenic P in Pseudanabaena FS39 does not affect anoxygenic P In the absence of DCMU, we measured simultaneous oxygenic and anoxygenic P. As oxygenic and anoxygenic P driven electron transport intersect in the PQ pool (Fig. 4), we expected that the rates influence each other by competing for the PQ pool. However, our data showed that the kinetics of anoxygenic P were not significantly affected by oxygenic P, because the H2S removal rates in the absence and presence of DCMU were indistinguishable (Fig. 2). Therefore, anoxygenic P always occurred at a rate that was determined either by [H2S] or the light intensity. It was, however, not affected by oxygenic P. Effects of H2S on oxygenic P During exclusive oxygenic P, which occurred in the absence of sulfide, the rate of oxygen production at a given light intensity was immediately constant after switching on the light and remained at this constant level independent of oxygen concentration in the range of 0–900 μM (Fig. S5). As expected, the rate of oxygenic P (oxyP) increased with the light 223 6. COMPETITION BETWEEN PHOTOSYNTHETIC MODES intensity. Saturation was reached at about 550 μmol photons m-2 s-1. Above this intensity oxyP decreased due to light inhibition (Fig. S6). In the presence of H2S a complex relation between anoxyP and oxyP was observed. As described above, anoxyP increased with [H2S] following MM kinetics below the light-specific [H2S] threshold (range A) and then remained constant at a light- dependent value of AnoxyPmax (range B) (Fig. 2 and 3). In the [H2S] range B, Pseudanabaena FS39 exclusively performed anoxygenic P. To describe the complex relation between anoxyP and oxyP in [H2S] range A, we further sub-devided it into [H2S] ranges A1, A2 and A3 (Fig. 3). Also, for a more meaningful comparison between anoxyP and oxyP, we converted the rates of O2 production and H2S consumption into electron equivalents (e-) according to Eqs. 1 and 2 for oxyP and anoxyP, respectively. OxyP in range A1 (low [H2S]), was constant (OxyPmax1) (Fig. 3). The rates of oxyP abruptly increased going from range A1 to A2 and remained constant at a higher level (OxyPmax2) that was, however, still lower than AnoxyPmax. In the [H2S] range A3 oxyP started to decrease. However, the overall sum of photosynthetic electron transport rates and the corresponding CO2 fixation rates reached their maximum in the [H2S] range A3. In this range the sum of electron transport by oxyP and anoxyP remained constant at AnoxyPmax (413 μM e- h-1). OxyP could therefore be calculated by subtracting the momentary electron transport rate of anoxyP from the maximum rate observed (OxyP = AnoxyPmax – AnoxyP) (thick lines in Fig. 3). Overall, H2S has two effects on oxyP in Pseudanabaena FS39. First, H2S stimulates oxygenic P (OxyPmax2 in range A2; Fig. 3). Secondly, H2S determines the rate of anoxyP, which in turn defines and limits the rate of oxyP (ranges A3 and B; Fig. 3). Thus, anoxyP is exclusively regulated by external factors, namely the [H2S] and light intensity, while oxyP is additionally controlled by the rate of anoxyP. Oxygenic photosynthesis is regulated by anoxygenic photosynthesis via the redox state of the PQ pool. The most straight-forward explanation for the relation between oxyP and anoxyP is that the affinity of SQR to PQ is substantially higher than the affinity of PSII components to PQ. We therefore suggest that the mechanistical basis for the out-competition of oxyP by anoxyP in Pseudanabaena FS39 is simple kinetic regulation. The rate of H2S oxidation by SQR (kH2S in Fig. 4) is limiting anoxyP at low [H2S] (range A in Fig. 2). The reduction 224 II. Manuscripts of PQ by SQR (kanoxy in Fig. 4) is however extremely fast. Therefore, oxyP is kinetically out-competed by anoxyP. Only if the rate of H2S oxidation (kH2S) is lower than the light- specific maximum electron transport rate through PSI (kPSI in Fig. 4), there is sufficient oxidized PQ available to allow also oxygenic photosynthetic electron transport (koxy in Fig. 4) to occur. That is, oxyP fills up the “left-over” electron transport rate. At high [H2S] (range B in Fig. 2) the electron transport rate through PSI (kPSI) is limiting. Thus, a light-dependent AnoxyPmax is reached and there is no “left-over” electron transport rate left for oxyP. These kinetic mechanisms imply that the rates of oxygenic P and anoxygenic P are regulated via the redox state of the PQ pool. This is because the rates of reduction (kanoxy and koxy) and oxidation (kPSI) of the PQ pool depend on its redox state, which in turn depends on [H2S] and light. kH2S H2S SQR S0 kanoxy H O 2 k koxy PSI PSII PQ PSI O2 I PSII IPSI Chl a Figure 4: Simplified model of the intersecting oxygenic and anoxygenic photosynthetic electron transport pathways. Chlorophyll a (Chl a) is the main light harvesting pigment in photosystem I (PSI). The phycobilisomes (blue) are the main accessory light harvesting antennae of photosystem II (PSII) that transfer excitation energy into the Chl a (red) containing reaction centre. Excitation energy can be redistributed between the photosystems in state 2 as indicated by the orange and red arrows. koxy is the rate of plastoquinone (PQ) reduction by PSII, i.e., the rate of oxygenic P. kanoxy is the reduction rate by sulfide:quinone:oxidoreductase (SQR), i.e., the rate of anoxygenic P. kPSI is the PQ oxidation rate by PS I, which corresponds to the overall photosynthetic electron transport rate. We implemented this concept into a numerical model based on Fig. 4 and Table 1 (details in the Supplemental Information 6), which gave a very good agreement with our experimental data (compare Fig. 2A and Fig. 2B). Also, this model revealed that the affinity constant of PSII components to PQ (KPSII in the rate koxy in Table 1) has to be at least two orders of magnitude higher than the affinity constant of SQR (KSQR in the rate 225 6. COMPETITION BETWEEN PHOTOSYNTHETIC MODES kanoxy in Table 1) to fit our experimental data. Also, our model confirmed that the [H2S] threshold, at which oxygenic and anoxygenic P can occur simultaneously, is predictable for any given light intensity (Supplemental Information 6.3). Table 1: Definition of the rate laws governing the reactions shown in Figure 4. Expression Description [H S] The H S oxidation rate by SQR. k = v 2 2 H 2S max K [H S] M 2 KM (38 ±1.2 μM) and vmax (biomass-dependent) were derived by fitting the data below AnoxyPmax in Figs. 2, 3 and S8. PQ The rate of PQ reduction by SQR. k = k ox anoxy H 2S K PQ SQR ox PQox refers to the oxidized part of the total PQ pool. KSQR describes the apparent affinity of SQR towards oxidized PQ. PQ The reduction rate of PQ by PSII. k = I ox oxy PSII K PQ PSII ox KPSII describes the apparent affinity of PSII towards oxidized PQ. IPSII describes the rate of exciton generation by the light energy harvested in PSII, for which we assume a linear relationship to electron transport at a given redox state of the PQ pool for the non-saturating light intensities used in our experiments. IPSII also defines the maximum rate of oxygenic P, i.e., OxyPmax. PQ The total rate of photosynthetic electron transport. k = I red PSI PSI K PQ PSI red KPSI describes the apparent affinity of PSI towards the reduced PQ. IPSI describes the rate of exciton generation by the light energy harvested in PSI, and defines the maximum rate of kPSI, i.e., AnoxyPmax. The model assumes that in a steady state kPSI = koxy + kanoxy. Light intensity and H2S concentration determine the maximal rates of oxygenic photosynthesis In the absence of sulfide, oxyP (OxyPmax1) was substantially lower than AnoxyPmax (Fig. 3). While the maximal anoxygenic photosynthetic electron transport exclusively depends on the light energy harvested in PSI, oxygenic photosynthetic electron transport depends on light energy harvested in both PSI and PSII. As OxyPmax1 was lower than AnoxyPmax, we suggest that oxyP (koxy) was limited by light energy harvested in PSII. OxyPmax1 remained constant at increasing [H2S] because it was not sufficiently high to fill the “left- 226 II. Manuscripts over” electron transport rate and therefore there was no competition for the PQ pool. This also implies that PSI was substantially more efficient in light energy harvesting than PSII. Remarkably, OxyPmax1 abruptly increased to higher rate (OxyPmax2) when both H2S and anoxyP reached a threshold. A stimulation of oxygenic P by H2S has already been reported for plants (20). These studies, however, involved long-term exposure to H2S, which lead to a gradual but substantial adjustment of the photosynthetic machinery (such as activity of RuBisCO, (20)). Also for a cyanobacterium an enhancement of CO2 fixation rates during oxygenic P was reported (6). The transition between OxyPmax1 and OxyPmax2 in our study organism Pseudanabaena FS39 occurred within less than 5 min. We therefore suggest that a plausible mechanism behind the increase of oxyP is state transition. State transitions are commonly used by cyanobacteria to adapt to short term fluctuations (e.g. in light intensity) to adjust the light energy distribution and the resultant electron transport rate between PSI and PSII (21). In state 1 the processes of energy harvesting in PSI and PSII are uncoupled, while excitation energy is transferred from PSII to PSI in state 2 to enhance PSI activity (21). Specifically, in state 1 the phycobilisomes are associated to PSII, whereas in state 2 the excitation energy harvested by the phycobilisomes is also driving PSI activity (22). Additionally, the energy harvested by chlorophyll a (Chl a) can be redistributed from PSII to PSI (23). To explain all our data, we needed to assume that excitation transfer is also possible in the other direction, i.e., from the Chl a in PSI to PSII. Although this has never been shown directly, it does not mean that it is theoretically impossible, especially when considering how the absorption, excitation and emission spectra of the photosystems I and II can vary among cyanobacterial species and depend on growth conditions (24). Also, bi- directional excitation transfer might not have been previously shown because under “normal conditions”, in the absence of H2S, state transition is induced by a reduced PQ pool, which can only be caused by limited light energy availability in PSI compared to PSII. The resulting “electron traffic jam” is removed by transferring more light energy to PSI (21). In contrast, in our experiment, light energy is limiting in PSII. Still, due to the combined activity of SQR (kanoxy) and oxyP (koxy) the PQ pool can become reduced. When their sum reaches a certain threshold, leading to a correspondingly highly reduced PQ pool, a state transition might occur. To test this hypothesis, we performed additional incubations at different spectral qualities of light (either targeting PSI or PSII) (Supplemental Information 5) and 227 6. COMPETITION BETWEEN PHOTOSYNTHETIC MODES implemented bi-directional state transition into our numerical model (Supplemental Information 6). The simulated data agreed with the experimental data remarkably well (compare Fig. S8 and S10). Also, opposed to changes in the pigment equipment of the cells or even photosystem stoichiometry (PSII:PSI ratio in the membranes), state transition can occur very fast. Thus, we conclude that state transition is the most probable mechanism controlling the increase of oxygenic P (going from range A1 to A2 in Fig. 3) in the presence of H2S. - CO2 and NO3 assimilation The electrons derived from either H2S during anoxygenic P or H2O during oxygenic P end up reducing NADP+ to NADPH in the last step of the electron chain. To understand the fate of the photosynthetically produced NADPH in Pseudanabaena FS39, we - converted the rates of CO2 and NO3 assimilation into rates in electron equivalents. Figure 3 shows that the CO2 assimilation rate per electron extracted during photosynthesis was lower during anoxygenic P than during oxygenic P. Specifically, up to 20 % of the - electrons available from H2S oxidation were channelled into NO3 reduction at the expense of CO2 reduction. In contrast, during oxygenic P up to 90 % of the NADPH potentially reduced during exclusive oxygenic P was channelled into CO2 reduction. The - NO3 reduction rates decreased to below detection limit (Fig. 3), despite lack of an alternative N-source, such as ammonium (data not shown). Based on this change in the - CO2:NO3 assimilation ratio, we suggest that during the first step of anoxygenic P, i.e., the oxidation of H2S to zero-valent sulfur by SQR, protons are not translocated across the membrane (7) and thereby do not contribute to the proton motive force and ATP generation. In contrast to oxygenic P, this leads to a higher cellular NADPH:ATP ratio and favours nitrate reduction over CO2 reduction, because less ATP compared to NADPH is required (25). Even though oxygenic P results in higher CO2 fixation rates per electron transported, growth rates are not necessarily lower during anoxygenic P. This is because the final CO2 fixation rate is a result of both the electron transport rate (higher during anoxyP) and the efficiency of converting reducing power into biomass (higher during oxyP). Interestingly, these two factors can compensate each other, which leads to equivalently high CO2 fixation rates during exclusive anoxygenic and exclusive oxygenic P (Fig. 3). Interestingly, the balance between electron transport rate and energy conservation efficiency (given by ATP generation per electron transported) is only 228 II. Manuscripts optimal during simultaneous oxygenic and anoxygenic P (range A3 in Fig. 3). Also, during this simultaneous anoxygenic and oxygenic P the N:C uptake ratio is close to Redfield (Fig. 3). Implications We have shown that the regulation of the switch between anoxygenic and oxygenic P in Pseudanabaena FS39 is predominantly determined by the rate of anoxygenic P. This is because anoxygenic P kinetically outcompetes oxygenic P in the step that transfers electrons to the PQ pool. This outcompetition of oxygenic P in the electron transport chain, however, does not imply that this process is outcompeted in the cyanobacterium under environmental conditions. Instead, this simple regulatory mechanism allows a very rapid and efficient adjustment of the photosynthetic activity upon changing environmental conditions. In microbial mats versatile cyanobacteria switch between anoxygenic and oxygenic P over a diurnal cycle, wherein anoxygenic P is expected to occur in the early mornings and evenings when sulfide is abundant (4). Intriguingly, the local concentration of sulfide is lowered by the activity of the cyanobacteria themselves due to anoxygenic P. Efficient sulfide removal and the flexible transition from anoxygenic to oxygenic P, as shown here for Pseudanabaena FS39, are therefore not only crucial for rapid adaptation to fluctuating light and sulfide concentrations, but also help the cyanobacterium switch to simultaneous oxygenic and anoxygenic P, which results in highest carbon-based growth rates (Fig. 3). Nowadays, adaptations of cyanobacteria to sulfide only play a role in some specific sulfidic ecosystems, such as the microbial mats in the Frasassi sulfidic springs, the habitat of Pseudanabaena FS39. Additionally, they do not seem to be linked to the phylogeny of cyanobacteria (5). Thus, most extant cyanobacteria are not equipped with the metabolic potential to perform anoxygenic P. However, oxygenic P probably evolved from anoxygenic forms of P (26) in a reduced environment and had to thrive in partially sulfidic oceans for billions of years. For instance, photosynthetic microbes in the mid-end-Proterozoic oceans probably lived in a euphotic zone characterized by downward gradients of light and oxygen opposed by upward gradients of sulfide and nutrients, similar to what we find nowadays condensed in microbial mats and biofilms (27, 28). In this scenario a stratified two-layer system might have established across the euphotic zone, with the lower, light-limited layer dominated 229 6. COMPETITION BETWEEN PHOTOSYNTHETIC MODES by anoxygenic P and the upper, oxic and nutrient-limited layer dominated by oxygenic P (2). Planktonic counterparts of Pseudanabaena FS39 might have been very successful in such oceans. This is because Pseudanabaena FS39 is highly adapted to an environment where aerobic conditions coincide with N-limitation and sulfidic conditions coincide with N-availability (Fig. 3). The preferential N-assimilation during anoxygenic P has also been shown in Geitlerinema sp. PCC 9228 (formerly Oscillatoria limnetica). Namely, this cyanobacterium increases its N2 fixation rates in the presence of sulphide. Cyanobacteria similar to Pseudanabaena FS39 or Geitlerinema sp. PCC 9228 would have assimilated N - (either in the form of NO3 or N2) during anoxygenic P in the oxic-anoxic interface of the photic zone of the Proterozoic oceans. Over a diurnal cycle this would have caused a downwards shift of this interface, leading to sulfide- and N-limitation in the zone where there is still light. However, this would not have caused limitation by the electron donor (H2S) in versatile cyanobacteria similar to Pseudanabaena FS39, because of their ability to rapidly shift to oxygenic P and thus continue growing at the same rate. Additionally, - possible carry-over of nutrients (e.g. NO3 ) taken up preferentially in deeper layers during anoxygenic P could have provided sufficient support for growth in the upper oxic zone while conducting oxygenic P in the presence of more light. This means that the proliferation of oxygenic P would not have been majorly disadvantaged by nutrient limitation, which might have driven primary productivity in general to higher levels. Thus, the adaptation strategy of cyanobacteria such as Pseudanabaena FS39 might represent an important intermediate step in evolution. This is because such cyanobacteria might have not only opened up the stage for obligate oxygenic phototrophs by their sulfide removal activity, but their adaptation strategy might have also created sufficient pressure on the obligate oxygenic phototrophs to evolve new strategies to cope with N- limitation, such as N2 fixation in the presence of oxygen, which is thought to represent a key factor that lead to the proliferation of oxygenic P, the consequent depletion of sulfide and outcompetiton of anoxygenic phototrophs – including Pseudanabaena FS39-like organisms. In general, the capability to perform both oxygenic and anoxygenic P was probably an important trait of cyanobacteria for large intervals of Earth’s history and played a crucial role in shaping primary productivity on a global scale (2). This is echoed in the astonishingly smooth and simple kinetic regulation of the transition between anoxygenic and oxygenic P in Pseudanabaena FS39. The simplicity of the regulation is especially remarkable when considering how complex the adjustment of photosynthesis 230 II. Manuscripts to changing environmental parameters can be. For instance, Gan et al. (24) recently reported that a Leptolyngbya sp. changes the relative transcript levels for >40% of its genome during photoacclimation to far-red light. In contrast, the simplicity of the control of anoxygenic P in Pseudanabaena FS39 suggests that anoxygenic P represent an extremely basic part of the photosynthetic metabolism, so important that it even regulates oxygenic P. Acknowledgements We thank the technicians from mechanical and electronical workshops for invaluable help during the construction of the setup, the technicians of the microsensor group for microsensor construction, M. Kuypers and H. Schulz-Vogt for providing access to lab facilities, G. Klockgether for her support during mass spectrometry measurements, K. Bischof for providing the diving PAM instrument, M. Kuypers, F. Widdel, U. Fischer and A. Chennu for inspiring discussions, J. Macalady and D. S. Jones for their support during sampling, A. Montanari and P. Metallo for the enjoyable atmosphere at the Osservatorio Geologico di Coldigioco during the sampling campaign, and A. Oren and D. Ionescu for providing the Oscillaroria limnetica culture. This work was financially supported by the Max-Planck-Society. References 1. Buick R. 2008. When did oxygenic photosynthesis evolve? Philos. Trans. R. Soc. Lond. B. Biol. Sci. 363:2731–2743. 2. 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The co-evolution of the nitrogen, carbon and oxygen cycles in the Proterozoic ocean. Am. J. Sci. 305:526–545. 233 6. COMPETITION BETWEEN PHOTOSYNTHETIC MODES ANOXYGENIC PHOTOSYNTHESIS CONTROLS OXYGENIC PHOTOSYNTHESIS IN A CYANOBACTERIUM FROM A SULFIDIC SPRING Judith M. Klatt, Mohammad A. A. Al-Najjar, Pelin Yilmaz, Gaute Lavik, Dirk de Beer, Lubos Polerecky Supplementary Information Content 1. Bioreactor 2. Isolation, cultivation and phylogenetic analysis of Pseudanabaena FS39 3. Oxygenic photosynthesis in Pseudanabaena FS39 4. Inhibition of oxygenic photosynthesis in Pseudanabaena FS39 by DCMU. 5. Effects of the spectral quality of light on oxygenic and anoxygenic photosynthesis 6. Model for the kinetic regulation of anoxygenic and oxygenic photosynthesis in Pseudanabaena FS39 via the redox state of the plastoquinone pool. 6.1. Formulation of the rate laws 6.2. Excitation energy transfer between PSII and PSI 6.3. Conditions determining the switch from anoxygenic to oxygenic P 7. Supplemental references 234 II. Manuscripts 1. Bioreactor Figure S1: Image of the experimental bioreactor. 2. Isolation, cultivation and phylogeenetic analyl sis of Pseudanabaenna FS39 Isolation and cultivation Strain FS39 (Fig. S2 and S3) was isolated from a biofilm growing iin a sulfidic spring emerging from the Frasassi cave system, Italy (1), by plating diluted samples on BG-11 agar medium. When growth was observed, isolated filaments were transferred to anoxic liquid Pfennig’s medium I supplemented with 1 mM total sulfide (H2S). Upon growth in this medium, filaments were picked under an inverted microscope and dilution series in BG-11 medium were done. Cyanobacteria growing in the tube with tthe highest dilution were transferred into sulfide gradiient medium. Specifically, the cyanobacteria were transferred into BG-11 agar (0.2 %, 50 °C), stirred and poured onto a solid sulfidic (10 mM) agar plug. Individual colonies along the sulfide gradient were picked and transferred to liquid BG-11 again. The complete procedure was repeated until the cyanobacterial strain was the only phototrophic mmicroorganism in the culture and populations of contaminant heterotrophic bacteria in actively growing cultures were minimized according to microscopy (cyanobaccterial-cell:other-cell ratio >200). Unfortunately, we were not able to obtain an axenic culture, because in the complete absence of contaminant 235 6. COMPETITION BETWEEN PHOTOSYNTHETIC MODES heterotrophic bacteria proliferation of the cyanobacterial filaments was suppressed. During the whole isolation procedure, the cultures were incubated under fluorescent lamps mimicking solar radiation (Envirolight, USA) at 200 μmol photons m-2 s-1 and at a temperature of 15 °C. For maintenance, cultures were kept in the same conditions like for isolation and transferred every 6 weeks alternating to fresh BG-11 and sulfidic medium. Cultures for the bioreactor incubations were grown in standard B-HEPES medium except - - for deviating NO3 and HCO3 concentration (4 and 8 mM, respectively). Phylogenetic analysis Biomass was harvested from 20–30 mL of the cyanobacterial cultures by centrifugation (3000 g, 15 min). DNA was extracted as previously described (2). The partial 16S rRNA gene sequence was amplified using the cyanobacteria-specific primers CYA106F and CYA781R (3). The partial sequence of the functional gene sqr encoding sulfide:quinone:oxidoreductase was amplified using primer SQR-G1-475F and SQR-G1- 964R (4). DNA extracted from the original strain of Oscillatoria limnetica from Solar Lake (5) served as a control for sqr amplification. Final concentrations of reactants in the polymerase chain reaction (PCR) and duration and temperature of the amplification steps for 16S rRNA and SQR partial sequences were used as described in Nübel et al. (1997) and Pham et al. (2008), respectively. The PCR product of the partial 16SrRNA sequence was purified using the QIAquick PCR purification kit (Qiagen). For sequencing the same cyanobacteria-specific reverse and forward primers were used and analyzed on an Applied Biosystems 3130xl DNAsequencer. The consensus partial 16S rRNA gene sequence was created in BioEdit and is deposited in the European Nucleotide Archive under the accession number LM993811. The sequence was submitted to BLAST (http://www.ncbi.nlm.nih.gov/BLAST/), which revealed that the strain is closely related to several Pseudanabaena spp., including P. catenata UAM 412, P. mucicola PMC269.06, P. biceps PCC 7429 and Pseudanabaena sp. PCC 7408. Identities were above 95%, which placed our strain within the genus boundaries of Pseudanabaena. Furthermore, the partial sequence had more than 99% identity with P. biceps PCC 7429 and Pseudanabaena sp. PCC 7408. For phylogenetic analysis the 16S rRNA sequence was aligned with SINA (6), and was merged with the SILVA SSURef_NR99_115 dataset (7) for further processing using software package ARB (8). Along with the strain FS39 16S rRNA gene sequence, 35 full- 236 II. Manuscripts length 16S rRNA gene sequences from other cultivated strains belonging to the Pseudanabaenaceae family were seelected for phylogenetic tree reconstruction. Several treeing algorithms (neighbour joining, maximum parsimony, and maximum likelihood) with and without alignment position and base conservation filters were applied, and each attempt returned almost identical tree topologies. Phylogenetic ttree reconstruction confirmed that the sequence forms a distinct cluster with other PPsseudanabaena spp. (Figure S4). Figure S2: Pseudanabaena strain FS39, as seen in a light microscope. Scale bar is 5 μm. Figure S3: DAPI stained Pseudanabaena FS39 filaments. Red colour represents autofluorescence, blue colour DAPI fluorescence A: Culture grown without stirring formed biofilms. B and C: Subsamples of Pseudanabaena FS39 culture taken during a bioreactor run. When grown in stirred culture and in the exponential growth phase filaments are shorter and do not form aggregates. 237 6. COMPETITION BETWEEN PHOTOSYNTHETIC MODES Figure S4: A phylogenetic tree showing the position of the cyanobacterial strain FS39 (Frasassi FS39) sequence with respect to other members of Pseudanabaenaceae famillyy. The tree was calculated using RAxML version 7.0.4, with a GTRGAMMA model, a 10% cyanobacterial base conservation filter, and with 1000 bootstraps. The strain FS39 sequence was added later to this tree using the ARB parsimony insertion tool, again applying the 10% base conserrvvation filter. The two Gloeobacter violaceus sequences servee as the out-group. Bootstrap values are only shown when they were above 60%, the bar indicates 0.01 substitutions per nucleotide possition. 238 II. Manuscripts 3. Oxygenic photosynthesis in Pseudanabaena FS39 dark light 120 1000 ) 100 -1 800 h 2 80 600 M) P M O M P ( 60 2 400 O 40 oxyP ( oxyP 20 200 0 0 0 4 8 12 16 20 time (h) Figure S5: Oxygen concentrations (O2) and the corresponding rates of oxygenic photosynthesis (oxyP, calculated as oxyP = dO2/dt) measured in the bioreactor with Pseudanabaena FS39 in the absence of sulfide in the dark and during illumination by orange and red light (total irradiance 200 μmol photons m-2 s-1; split equally between the orange and red light). 160 140 ) -1 120 h 2 100 80 M O M P 60 40 OxyP ( OxyP 20 0 0 200 400 600 800 1000 1200 I (μmol photons m-2 s-1) Figure S6: Oxygenic photosynthesis (oxyP) by Pseudanabaena FS39 as a function of irradiance (I) in the bioreactor (total irradiance was split equally between the orange and red light). Fitting -2 -1 by the model of Eilers and Peeters (9) gave Im = 553 μmol photons m s , Ik = 198 μmol photons -2 -1 -1 m s and Pm = 140 μM O2 h (Pm is biomass-dependent). 239 6. COMPETITION BETWEEN PHOTOSYNTHETIC MODES 4. Inhibition of oxygenic photosynthesis in Pseudanabaena FS39 by DCMU 700 2000 dark 50 200 200 50 600 DCMU injected 1500 500 (μM) 400 S 2 1000 2 H 300 O (μM) Σ 200 500 100 0 0 0 10 20 40 80 90 100 110 time (h) Figure S7: Inhibition of oxygenic photosynthesis in Pseudanabaena FS39 by DCMU. Shown are dynamics of oxygen and total sulfide concentrations in the bioreactor before and after the addition of DCMU. Measurements were performed in the dark and at irradiances of 50 μmol photons m-2 s-1 and 200 μmol photons m-2 s-1. 5. Effects of the spectral quality of light on oxygenic and anoxygenic photosynthesis We measured rates of oxygenic P (oxyP) and anoxygenic P (anoxyP) dependent on [H2S] under the exposure to different spectral qualities of light. Specifically, we used red light (Ȝmax=690 nm) to target Chlorophyll a (Chl a), which is the main light-harvesting pigment in PSI (10). PSII excitation energy is mainly provided by accessory pigments, the phycobilins (Pb) in the phycobilisomes, and only peripherally by Chl a (10). To target Pb, we used orange light (Ȝmax = 590 nm). Independent of the spectrum used, the overall photon flux was kept the same (200 μmol photons m-2 s-1). During constant orange illumination, anoxyP increased with [H2S] following Michaelis-Menten (MM) kinetics below the light-specific [H2S] threshold (range A) and then remained constant at a light-limited value of AnoxyPmax (range B in Fig. S8A). Conversion of the rates of O2 production and H2S consumption into electron equivalents (e-) according to Eqs. 1 and 2 for oxyP and anoxyP, respectively, revealed that the sum of electrons transported remained constant at 132 μM e- h-1 throughout the experiment, regardless of [H2S]. Only the partitioning between anoxyP and oxyP changed. The sum of the rates of CO2 and nitrate assimilation/reduction was constant in this experiment irrespective of the source of electrons, i.e., of the type of photosynthesis (~120 μM e- h-1; Fig. S8A; Table S1). 240 II. Manuscripts In contrast to orange illumination, the sum of the electron transport rate during constant red illumination did not remain constant over the complete range of [H2S] (Fig. S8B). Specifically, at low [H2S] a complex relation between anoxyP and oxyP was observed. We divided this pattern into operational [H2S] ranges A1, A2, A3 and B. AnoxyP followed the MM-kinetics over the concentration ranges A1, A2 and A3 and remained constant at AnoxyPmax in the [H2S] range B. OxyP in range A1 was constant (OxyPmax1) and lower than anoxyPmax. Intriguingly, the rates of oxyP abruptly increased going from range A1 to A2 and remained constant at a higher level (OxyPmax2). However, the overall sum of photosynthetic electron transport rates and the corresponding CO2 fixation rates reached their maximum only in the [H2S] range A3. In this range the sum of - -1 electron transport by oxyP and anoxyP remained constant at AnoxyPmax (694 μM e h ). As observed also during orange illumination, the sum of the rates of CO2 and nitrate - -1 assimilation/reduction was constant in this [H2S] range (635 μM e h ; Fig. S8B, Table S1). During exposure to a mixture of half the intensity of red light and half the intensity of orange light described above (red+orange light), which more realistically resembles ambient light conditions, we observed a similar complex pattern as during red illumination (Fig. 3 and S7C). The exceptions were a shift in the maximal values of oxyP and anoxyP (Table S1) and in the boundaries of the operational ranges A1-B (Fig. 3 and S7C). In summary, the spectral quality of the light had no effect on the MM kinetics of H2S oxidation (Fig. S8). It however impacted the maxima of oxyP and anoxyP (Table S1). Additionally, we observed that the partitioning between oxygenic and anoxygenic P followed different patterns dependent on [H2S]. 241 6. COMPETITION BETWEEN PHOTOSYNTHETIC MODES 150 A B ) -1 h - 100 Me P 50 AnoxyP rate ( OxyP - A CO2 +NO3 as s imilation [H2S]thr 0 0 5 10 15 20 25 P H2S( M) 800 A1 A2 A3 B ) -1 600 h - Me 400 P rate ( 200 [H S] B 2 thr 0 0 50 100 150 200 H S(PM) 2 500 A1 A2 A3 B 400 ) -1 h - 300 Me P 200 rate ( 100 [H S] 2 thr C 0 0,15 ’ m F 800 0,10 Y 700 0,05 F’ 600 0,00 0 1 2 3 10203040506070 H S(PM) 2 Figure S8: Rates of anoxygenic (anoxyP) and oxygenic (oxyP) photosynthesis, and the sum of nitrate and carbon dioxide assimilation rates as a function of H2S concentration during exposure to (A) orange light, (B) red light and (C) orange and red (red+orange) light. In all cases the total photon flux was 200 μmol photons m-2 s-1. Modeled rates are represented by the thick lines. Annotations on top of (B) and (C) indicate the operational [H2S] ranges (A1, A2, A3, B) of the complex switch between the photosynthetic modes. Panel (C) additionally shows variable fluorescence yields (F’ and Fm’) and the corresponding effective quantum yield of PSII (Y), which was measured with a fibre-bundle connected to a pulse amplitude modulated fluorometer (Diving- PAM, Walz, Germany). Specifically, fluorescence yields during actinic illumination just before -2 - (F’) and at the end (Fm’) of a saturating pulse (duration 0.8 s, intensity ~2000 μmol photons m s 1 ) were used to calculate the effective quantum yield of PSII at each time point as Y = (Fm’ – F’)/Fm’ (11). 242 II. Manuscripts 6. Model for the kinetic regulation of anoxygenic and oxygenic photosynthesis in Pseudanabaena FS39 via the redox state of the plastoquinone pool We propose the following simplified model of kinetic regulation of anoxyP and oxyP in Pseudanabaena FS39 as a mechanistic basis for the outcompetition of oxyP by anoxyP and for the conspicuous enhancement of oxyP at low [H2S] in the operational range A2 (Fig. S8). Using the model we show that the complete regulation of the switch between oxyP and anoxyP can be explained based on very limited regulation mechanisms: (i) the affinity of SQR to H2S, (ii) the resulting redox state of the PQ pool and (iii) light energy harvested in PSI and PSII, wherein H2S indirectly induces state transition via the redox state of the PQ pool allowing redistribution of this energy, which can lead to the increase of oxyP. We also show that the condition at which the switch between oxyP and anoxyP occurs can be predicted from a very simple relationship between [H2S] and available light. 6.1 Formulation of the rate laws The model focuses on the central component of the anoxygenic and oxygenic photosynthetic electron transport chain, namely the PQ pool, which is where both pathways intersect (Fig. 4). Specifically, the ratio of the reduction rate of PQ by SQR (kanoxy in Fig. 4) and the reduction rate of PQ by PSII (koxy) directly translates into the partitioning between anoxygenic and oxygenic electron transport, respectively, whereas the rate of PQ oxidation (kPSI) corresponds to the sum of both processes, i.e., the rate of electron flow through PSI. The rate of PQ reduction by SQR (kanoxy) depends on (i) the H2S oxidation rate by SQR (kH2S in Fig. 4) and (ii) the affinity of SQR to oxidized PQ according to [H 2S] PQox k anoxy = vmax (S1) K M [H 2S] K SQR PQox where PQox refers to the oxidized part of the total PQ pool and KSQR describes the affinity of SQR towards this oxidized PQ pool. KM (38 (+/– 1.2) μM) and vmax can be derived by fitting the data below AnoxyPmax in Figs. 2, 3 and S7 with Eq. S1 assuming PQox>>KSQR. The reduction rate of PQ by PSII (koxy) is described similarly using an affinity (KPSII) towards PQox. Additionally, koxy depends on the rate of exciton generation by the light energy harvested in PSII (IPSII), for which we assume a linear relationship to electron transport at a given redox state of the PQ pool for the non-saturating light intensities used in our experiment. Taken together, we get 243 6. COMPETITION BETWEEN PHOTOSYNTHETIC MODES PQox koxy = I PSII (S2) PQox + K PSII for the reduction kinetics of the PQ pool from the PSII side. In this equation the term IPSII describes the maximum of koxy, i.e., OxyPmax. The sum of both electron transport pathways, i.e., the total rate of photosynthetic electron transport (kPSI), is formulated as PQred k PSI = I PSI , (S3) PQred + K PSI where IPSI is the rate of exciton generation in PSI and PQred is the pool of reduced PQ. The term IPSI corresponds to the maximum kPSI, which is AnoxyPmax. The model is solved by finding a steady state solution to the differential equation dPQ red = k +k k , (S4) dt anoxy oxy PSI which describes the rate of change in the reduced fraction of the plastoquinone PQ pool (PQred) as a result of its reduction by PSII and SQR and oxidation by PSI. When finding a solution the sum of the reduced and oxidized fractions of the PQ pool are constant, i.e., PQred + PQox = PQtot. The kinetic laws that control the rates of reduction (kanoxy and koxy) and oxidation (kPSI) of the PQ pool depend on the H2S concentration ([H2S]), the rate at which excitation energy is harvested and converted to the flow of electrons through the electron transport chain by PSI (IPSI) and PSII (IPSII), and the redox state of the PQ pool, as given by Eqs. S1–S3. -1 -1 Since PQred is expressed in mol electrons L s , the rate of H2S consumption and O2 production corresponding to the rate of anoxygenic (kanoxy) and oxygenic (koxy) P is given as d[H S] k 2 = ȕ anoxy (S5) dt 2 d[O ] k 2 = oxy (S6) dt 4 Here, the factors 2 and 4 correspond to the amount of electrons transferred per molecule of H2S consumed and molecule of O2 evolved, respectively, and the factor ȕ relates the depletion of H2S from the total sulfide pool to the change in H2S concentration and thus depends on the pH (e.g., ȕ = 1/1.7 at pH=7, ȕ = 1/8.2 at pH=8). For a given value of [H2S], IPSI and IPSII, the steady-state redox state of the PQ pool is calculated by setting dPQred/dt = 0 in Eq. S4, from which the corresponding rates of oxygenic and anoxygenic P are calculated using Eqs. S1–S3. 244 II. Manuscripts The numerical implementation of this model revealed a remarkably good agreement with the experimental data when setting the affinity constant KPSII to an at least two orders of magnitude higher value than the affinity constant of SQR, KSQR. (compare Fig. 2A and B). Based on this we conclude that anoxygenic P in Pseudanabaena FS39 has a kinetic advantage over oxygenic P. 6.2 Excitation energy transfer between PSII and PSI Our data shows that OxyPmax abruptly increases in the [H2S] range A2 during red and red+orange illumination (Fig. 3, S7B and S7C). According to our model, an increase of OxyPmax (Eq. S2) can only be caused by an increase in the harvested photon flux. We propose that this increase in light energy is caused by state transition from state 1 to state 2, whereby in state 2 the two photosystems PSI and PSII are able to redistribute excitation energy (12). Due to the combined activity of SQR (kanoxy) and oxyP (koxy) the PQ pool can become reduced. When their sum reaches a certain threshold, leading to a correspondingly highly reduced PQ pool, a state transition might occur. This threshold indirectly depends on H2S, because [H2S] governs kanoxy (Eq. 3). We hypothesize that energy transfer from PSI to PSII leads to the increase of koxy in state 2 (going from range A1 to A2). This is consistent with the measurements of variable fluorescence, where we found that the H2S-induced increase in oxygenic P was accompanied by a decrease in the effective quantum yield of PSII, Y (Fig. S8C). The decrease in Y is indicative of a more reduced PQ pool, which induces state transition allowing the energy redistribution needed to enhance oxygenic P. For the implementation of state transition into our model, we postulate that for the given external photon fluxes of the red (Ir) and orange (Io) light, the rates of exciton generation at the reaction centre of the PSI and PSII are given, respectively, as I PSI = tĮo Io + 1 f 1 r Įr I r , (S7) I PSII = 1 t Įo Io + rĮr Ir + f 1 r Įr Ir . (S8) The parameters Įo and Įr describe how the externally available photons of the respective colour are harvested and converted into quanta of excitation energy (excitons) that eventually result in the supply of electrons from the reaction centre to the electron transport chain. Since the unit of the photon flux is mol photons m-2 s-1 and the unit of the -3 -1 electron generation rate is mol e m s , the unit of Įo and Įr is mol electrons (mol photon)-1 m-1, or shortly m-1. The factors t, r and f parameterize how the excitation photon energy is harvested and distributed between the two photosystems depending on the redox 245 6. COMPETITION BETWEEN PHOTOSYNTHETIC MODES state of the PQ pool, i.e., for the two possible states 1 and 2 (Fig. S9). Specifically, the factor t describes the fraction of the excitons generated by the orange light that drive PSI, while the complementary fraction (1–t) drives PSII. This factor is 0 in state 1 and 0 As follows from Eqs. S1–S3, IPSI and IPSII generally describe the maximal rates of anoxygenic and oxygenic P, reached when the PQ pool is fully reduced (PQred § PQtot) and oxidized (PQox § PQtot). The measured values of these maximal rates are given in Table S1 for the different illumination conditions. Because Eqs. S1–S3 and S7–S8 make it possible to find expressions for these maximal rates (see Table S1, last column), we combined these expressions with the measured maximal rates of anoxygenic and oxygenic P to estimate the values of the model parameters r, t, f, Įr and Įo. Specifically, -1 for the cyanobacterial biomass used in the bioreactor experiments we obtained Įr = 4 m -1 and Įo = 2 m . Furthermore, to simulate the experimentally observed abrupt decrease in oxygenic P at low H2S concentrations (compare Fig. S10 with Fig. S8B-C; transition from A2 to A1), we additionally had to assume that the parameters t and f depend on the redox state of the PQ pool as PQred t = tmax (S9) PQref + Kt PQred f = f max (S10) PQref + K f where tmax = 0.334, fmax = 0.13, Kt § 0.1PQtot and Kf § 0.01PQtot, while the parameter r = 0.14 was independent of the PQ pool redox state. This gave an excellent agreement with the experimental data (compare Figs 2A and 3 with Figs. 2B and S9, respectively). Taken together, our model predicts that for the studied Pseudanabena strain FS36 (i) about 14% of the utilized red light drives PSII while the remaining 86% drives PSI, (ii) under fully reduced PQ pool (state 2) about 33% of the excitation energy harvested by phycobilins associated with PSII (orange light) is transferred to PSI while about 13% of the excitation energy harvested by Chl a-containing antennae associated with PSI (red light) is transferred to PSII. 246 II. Manuscripts Figure S9: Schematic diagram of the photosynthetic electron transport pathways in Pseudanabaena FS39. The pathways driiven by oxygenic and anoxygenic phhootosynthesis intersect in the plastoquinone (PQ) pool. The redox state of the PQ pool depends onn the balance between the rates of electron transport from PSIII (oxygenic photosynthesis, koxy) and SQR (anoxygenic photosynthesis, kanoxy) to PQ and from PQ to PSI (kPSI). The rates koxy and kPPSI are determined by the light energy availability and the corresponding exciton generation in PSII (given by IPSII) and PSI (given by IPSI), respectively. Our data can be explained if, upon a transition to state 2 triggered by a sufficiently reduced PQ pool, a certain fraction of the excitation enerrgy generated by the accessory pigments associated with each photosystem is transferred to the other photosystem, as indicated by the dashed arrows: orange ffor energy transfer from PSII-associated phycobilins (Pb) to PSI, red for energy transfeer from PSI--associated Chl a to PSII. The magnittude of this transfer is defined by the factors t and f. In state 1, i.e., when the PQ pool is sufficiennttly oxidized, no such transfer occurs and both t and f are zero. 1000 A1 A2 A3 B ) 800 -1 h anoxyP - 600 oxyP Me P 400 rate ( ( 200 [H S] 2 thr 0 0 50 100 150 200 250 H S (PM) 2 Figure S10: Simulation of oxygenic and anoxygenic photosynthesis in Pseudanabaena FS39 under red illumination. Rates are pllotted as a function of H2S concentrations for external irradiance of 200 μmol photons m-2 s-1. Simulated data reproduce well the main features of the experimental data (compare with Fig. S8B). 247 6. COMPETITION BETWEEN PHOTOSYNTHETIC MODES r I r Į r r o ) I r I I r o Į –r r r Į Į ) I I r ) (1 ‡ r –r Į Į r –r = t ) (1 I r (1 + f + r PSI –r Į + r + k + I o r r o (1 + f I Į r = I = o o I Į = Į r oxy oxy max Į + r k PSI = o k = = t I oxy o = r k = Į PSI oxy k max1 = k max = = = oxy = AnoxyP = max1 k max max2 max = applicable. max2 = 132 132 = oxyP equation Corresponding max ) -1 h = 694 = 413 = anoxyP anoxyP = 115 oxyP - = 205 = 255 = oxyP 300 = oxyP oxyP max max max1 max2 max1 max2 (μM e † = AnoxyP = P OxyP FS39 under different combination and orange light. illumination by of red OxyP OxyP OxyP max AnoxyP AnoxyP OxyP § Pseudanabaena II and PSI (see schematic diagram in Figure S9). n.a. = not = n.a. S9). in Figure diagram schematic (see PSI II and r f § t as derived from the model (Eq. S1-S3 and S7-S10). S7-S10). and S1-S3 (Eq. model the from derived as S]) 2 n.a. 0.14 0.13 0.13 0.14 2 S]) n.a. 2 S]) 0.13 0.14 0 2 2 (r=red light, o=orange light). light). o=orange (r=red light, S]) 1 n.a. 0.14 0 0.14 n.a. 1 S]) 0 0.14 0 S]) 1 -1 2 2 S] 2 0.33 n.a. n.a. n.a. n.a. 0.33 2 S] s S]) 2 n.a. 0.14 0 0.14 n.a. 2 S]) 0 0.14 0.33 2 S]) 2 2 2 -2 on energy redistribution between PS redistribution on energy S] range S] range State 2 [H mol photons m mol μ Rates of oxygenic and anoxygenic photosynthesis in Rates of oxygenic and anoxygenic photosynthesis * r I * o 0 200 [H – medium A2 (low 0 200 A1 (0– low [H 0 200 B (high [H I 100 100 A1 (0– low [H 100 100 B (high [H 100 100 [H – medium (low A2 200 200 0 [H 0 – high Photon flux in flux in Photon describing the excitati Factors rates. determined Experimentally of photosynthesis, rates for the maximal Expressions Table S1: * § † ‡ 248 II. Manuscripts 6.3 Conditions determining the switch from anoxygenic to oxygenic P Since the switch between oxygenic and anoxygenic P in Pseudanabaena FS39 appears to be exclusively kinetically controlled, the remaining important question is: can it be predicted from the environmental conditions to which a single cell of Pseudanabaena FS39 is exposed? Our data show that the critical parameter defining whether or not oxygenic P occurs is the threshold sulfide concentration, [H2S]thr (Fig. 2 and 3). This threshold can be well approximated as the point where the curve characterizing the dependence of SQR activity on H2S intersects the line characterizing the maximal anoxygenic P at the given light. Using Eqs. S1–S3, this intersecting point is calculated as O [H S] = K (S11) 2 thr M 1 O where Ȝ = IPSI /vmax. It is important to note that both IPSI and vmax depend on the cyanobacterial biomass in the experimental reactor. However, as will be shown in the following, their ratio Ȝ does not. Specifically, let us assume that the concentration of the cyanobacterial biomass in the reactor, B, is given as the chlorophyll a concentration (e.g., mol Chl m-3), which is common in photosynthetic research. Under this assumption, the parameters Įo and Įr, which determine the conversion of the externally available flux of orange and red photons, respectively, into electrons that pass through the photosystems I and II (see Eqs. S7-S8) can be written as Di =KiV i B (S12) where Și is the quantum efficiency of photon-to-exciton conversion (expressed in units of -1 mol electrons (mol photons) ), ıi is the effective cross-section of the pigment-containing antennae that captures the photons (expressed in m2 (mol chl)-1), and i=o and i=r for orange and red light, respectively. Thus, Į in Eqs. S4-S5, is expressed as a product of fundamental properties of the photon-capture apparatus of the cell (Ș and ı) and the cellular biomass in the bioreactor (B). A quick check of units gives that Į is in mol e- (mol photon)-1 m-1, as required. In a similar way, the parameter vmax can be written as UmaxG SQR vmax = B (S13) Fchl where ȡmax is the maximal rate of H2S oxidation by a single molecule of the SQR enzyme under substrate non-limiting conditions (expressed in units of mol e- s-1 (mol SQR)-1), 249 6. COMPETITION BETWEEN PHOTOSYNTHETIC MODES įSQR is the density of the SQR enzyme in a cyanobacterial cell (expressed in mol SQR -1 -1 cell ) and Ȥchl is the cellular chlorophyll content (expressed in mol Chl cell ). Thus, vmax in Eq. S1 is expressed as a product of fundamental properties of the SQR enzyme (ȡmax), of the cell (įSQR and Ȥchl) and of the cellular cellular biomass in the bioreactor (B). A - -3 -1 quick check of units gives that vmax is in mol e m s , as required. Combining Eqs. S7, S12 and S13, we obtain that the parameter Ȝ is independent of the cyanobacterial biomass, B, and given by O tN o I o (1 f )(1 r)N r I r , (S14) where KiV i Fchl Ni = . (S15) UmaxG SQR The parameters ți (i=o and i=r for orange and red light, respectively, as above) can be calculated if each of the individual parameters involved is quantified separately. Although this is, at least in principle, experimentally possible, it would go far beyond the scope of this study. Nevertheless, by combining the complete set of H2S threshold values determined experimentally at the different light conditions (Table S2), we were able to 2 -1 determine these parameters empirically as țo = 2200 m s (mol photons) and țr = 5050 m2 s (mol photons)-1. Taken together, equations S11 and S14-S15 show that there is a direct relationship between the externally available light (Ir and Io) and the threshold H2S concentration below which the studied Pseudanabaena FS39 starts performing oxygenic P. This simple relationship makes it possible to link environmental parameters (light, O2 and H2S) with the fundamental processes that shape them (oxygenic and anoxygenic photosynthesis). 250 II. Manuscripts Table S2: Comparison of predicted and measured threshold H2S concentrations below which Pseudanabaena FS39 starts performing oxygenic photosynthesis. Predicted * * † Io Ir Ȝ Measured [H2S]thr † [H2S]thr 100 100 0.450 32 40 25 25 0.113 5 4.5 200 0 0.147 6.7 6.7 0 200 0.754 119.5 120 100 100 0.450 32 32 * Photon flux in μmol photons m-2 s-1 (r=red light, o=orange light). † Calculated from Eq. S8 and S11 using the following parameters values: t =0.334, r=0.142, f=0.13, 2 -1 2 -1 țo=2200 m s (mol photons) , țr = 5050 m s (mol photons) . 251 6. COMPETITION BETWEEN PHOTOSYNTHETIC MODES Supplemental references 1. Galdenzi S, Cocchioni M, Morichetti L, Amici V, Scuri S. 2008. Sulfidic ground-water chemistry in the Frasassi caves, Italy. J. Cave Karst Stud. 70:94–107. 2. Ionescu D, Voss B, Oren A, Hess WR, Muro-Pastor AM. 2010. Heterocyst- specific transcription of NsiR1, a non-coding RNA encoded in a tandem array of direct repeats in cyanobacteria. J. Mol. Biol. 398:177–88. 3. Nübel U, Garcia-Pichel F, Muyzer G. 1997. PCR primers to amplify 16S rRNA genes from cyanobacteria. Appl. Environ. Microbiol. 63:3327–32. 4. Pham VH, Yong J-J, Park S-J, Yoon D-N, Chung W-H, Rhee S-K. 2008. Molecular analysis of the diversity of the sulfideௗ: quinone reductase (sqr) gene in sediment environments. Microbiology 154:3112–21. 5. Cohen Y, Jørgensen BB, Padan E, Shilo M. 1975. Sulphide-dependent anoxygenic photosynthesis in the cyanobacterium Oscillatoria limnetica. Nature 257:489–492. 6. Pruesse E, Peplies J, Glöckner FO. 2012. SINA: accurate high-throughput multiple sequence alignment of ribosomal RNA genes. Bioinformatics 28:1823–9. 7. Quast C, Pruesse E, Yilmaz P, Gerken J, Schweer T, Yarza P, Peplies J, Glöckner FO. 2013. The SILVA ribosomal RNA gene database project: improved data processing and web-based tools. Nucleic Acids Res. 41:D590–6. 8. Ludwig W, Strunk O, Westram R, Richter L, Meier H, Yadhukumar, Buchner A, Lai T, Steppi S, Jobb G, Förster W, Brettske I, Gerber S, Ginhart AW, Gross O, Grumann S, Hermann S, Jost R, König A, Liss T, Lüssmann R, May M, Nonhoff B, Reichel B, Strehlow R, Stamatakis A, Stuckmann N, Vilbig A, Lenke M, Ludwig T, Bode A, Schleifer K-H. 2004. ARB: a software environment for sequence data. Nucleic Acids Res. 32:1363–71. 9. Eilers PHC, Peeters JCH. 1988. A model for the relationship between light intensity and the rate of photosynthesis in phytoplankton. Ecol. Modell. 42:199– 215. 10. Allen JF, Mullineaux CW, Sanders CE, Melis A. 1989. State transitions, photosystem stoichiometry adjustment and non-photochemical quenching in cyanobacterial cells acclimated to light absorbed by photosystem I or photosystem II. Photosynth. Res. 22:157–66. 11. Baker NR. 2008. Chlorophyll fluorescence: a probe of photosynthesis in vivo. Annu. Rev. Plant Biol. 59:89–113. 12. McConnell MD, Koop R, Vasil’ev S, Bruce D. 2002. Regulation of the distribution of chlorophyll and phycobilin-absorbed excitation energy in cyanobacteria. A structure-based model for the light state transition. Plant Physiol. 130:1201–12. 252 III. Manuscripts 7. REGULATION OF THE PARTITIONING BETWEEN OXYGENIC AND ANOXYGENIC PHOTOSYNTHESIS IN A CYANOBACTERIUM DOMINATING SULFIDIC SPRING MICROBIAL MATS Klatt, Judith M.1*; Häusler, Stefan1; de Beer, Dirk1; Polerecky, Lubos1,2 1Max Planck Institute for Marine Microbiology, Celsiusstr.1, 28359 Bremen, Germany 2Department of Earth Sciences – Geochemistry, Faculty of Geosciences, Utrecht University, Princetonplein 9, 3584 CC, Utrecht, The Netherlands In preparation *Corresponding author 253 7. H2S AND LIGHT CONTROL CYANOBACTERIAL PHOTOSYNTHESIS Abstract We used microsensors to study the regulation of oxygenic and anoxygenic photosynthesis by light and sulfide in a cyanobacterium dominating microbial mats from cold sulfidic springs. Both photosynthetic modes were performed simultaneously in a wide range of -2 -1 H2S concentrations (0–2200 μM) and irradiances (3–38 μmol photons m s ). At a given irradiance H2S controlled the partitioning between oxygenic and anoxygenic photosynthetic rates but not the overall photosynthetic electron transport rate. As this partitioning depended also on irradiance, we propose that it is kinetically controlled by the affinity of the electron transport components involved in anoxygenic (sulfide:quinone:reductase, SQR) and oxygenic (photosystem II, PSII) photosynthesis towards plastoquinone, which is where both electron transport pathways intersect. Fitting our data with a simple model of the electron transport chain revealed that the SQR enzyme in this cyanobacterium has a very low apparent affinity towards H2S whereas its apparent affinity towards oxidized plastoquinone is similar to that of PSII. We discuss how these specific regulation mechanisms make the studied cyanobacterium successful in its environment. Introduction Oxygenic photosynthesis (OP) is a process where light energy is used to extract electrons from water to reduce CO2. The evolution of this type of photosynthesis was predated by anoxygenic photosynthesis (AP), a process that uses another compound (e.g., H2S) as electron donor for the reduction of CO2 . While the more ancient AP requires only one photosystem (PSI) to drive the electron flow, OP requires the combined power of two photosystems (PSI+PSII), primarily because of the extraordinarily high energy demand of the water splitting reaction. OP probably evolved in cyanobacteria (2) inhabiting microbial mat-like structures. In these systems alternative electron donors for photosynthesis such as H2S were abundant (3, 4) and remained available until the end of the Proterozoic (5). Therefore, it has been hypothesized that AP performed by obligate anoxygenic phototrophs and cyanobacteria capable of both AP and OP (referred to as versatile cyanobacteria) was the dominant photosynthetic mode before the complete oxygenation of the Earth’s atmosphere and oceans (6). However, this oxygenation would not have been possible if cyanobacteria had not continuously produced O2 over billions of years. It is therefore intriguing why oxygenic phototrophs were so successful despite the widespread 254 III. Manuscripts availability of electron donors for anoxygenic phototrophs and the toxic effects of H2S on the components of OP, especially considering that AP is a biochemically less complicated and energetically less demanding process than OP (7, 8). New insights into possible mechanisms that allowed outcompetition of anoxygenic phototrophs by oxygenic phototrophs in the presence of H2S can be gained by studying adaptations of extant cyanobacteria living in sulfidic environments. The cold, light-exposed sulfidic springs at Frasassi, Italy (9) (J. M. Klatt, S. Meyer, S. Häusler, J. L. Macalady, D. de Beer, L. Polerecky; submitted for publication), which harbour thin microbial mats inhabited by diverse anoxygenic, oxygenic and versatile phototrophs, are one example of such an environment. Our recent studies of cyanobacteria isolated from this system revealed two novel mechanisms of cyanobacterial adaptation to H2S. The first mechanism, observed for a versatile cyanobacterium Pseudoanabaena str. FS39, included partitioning between AP and OP that is regulated kinetically by the apparent affinity of the electron transport components involved in AP (sulfide:quinone:reductase, SQR) and OP (photosystem II, PSII) towards plastoquinone, which is where both electron transport pathways intersect, and H2S (8). Specifically, OP in this cyanobacterial strain is active only when the electron transport chain is not fully used by AP, which occurs when H2S is limiting. The second mechanism, observed for the obligatory oxygenic cyanobacterium Planktothrix str. FS34, included acceleration of the recovery of OP by H2S after prolonged exposure to darkness and anoxia combined with an enhancement of OP rates by H2S at low irradiance and a delayed inhibition of OP leading to a temporary resistance to H2S (10). Although both Pseudanabaena str. FS39 and Planktothrix str. FS34 have adaptations that could make them successful in the microbial mats from the Frasassi sulfidic springs, which are characterized by a microenvironment with rapidly fluctuating availability of light and H2S (J. M. Klatt, S. Meyer, S. Häusler, J. L. Macalady, D. de Beer, L. Polerecky; submitted for publication), microscopic observations revealed that neither of them appeared to be the key player. Instead, the mats where dominated by yet another cyanobacterial morphotype. The aim of this study was to understand the specific adaptations responsible for the success of this dominant cyanobacterium in the system. We hypothesized that the dominant cyanobacterium possesses a mechanism that allows it to rapidly switch between AP and OP based on the instantaneous availability of light and H2S and thus be adapted to the rapidly fluctuating microenvironmental conditions in the mats. However, we expected that the mechanism differs from that found in 255 7. H2S AND LIGHT CONTROL CYANOBACTERIAL PHOTOSYNTHESIS Pseudanabaena str. FS39, given that the abundance of this strain in the mats is very low. To test this hypothesis, we quantified the combinatory effects of light and H2S on the partitioning between AP and OP performed by the dominant cyanobacterium. Because our attempts to isolate it were not successful, we performed our measurements in a natural mat sample dominated by the cyanobacterium using microsensors. Methods Microsensors O2, H2S and pH microsensors with a tip diameter of 10–50 μm and response time of <2 s were built, calibrated and used for profiling and monitoring of local concentration dynamics as previously described (11–13) (J. M. Klatt, S. Meyer, S. Häusler, J. L. Macalady, D. de Beer, L. Polerecky; submitted for publication) A fiber-optic microprobe for scalar irradiance (14) was used to measure the locally available light in the mat. The microprobe was connected to a spectrometer (USB4000, Ocean Optics, USA) and calibrated against a PAR scalar irradiance sensor connected to a light meter (LI-250, both from LI-COR Biosciences, Lincoln, NE, USA) as previously described (15). Mat sampling and measurement protocol Thin cyanobacterial mats together with the underlying sediment were sampled from the Frasassi sulfidic springs (J. M. Klatt, S. Meyer, S. Häusler, J. L. Macalady, D. de Beer, L. Polerecky; submitted for publication) in September 2009. Immediately after sampling they were placed in a flow-through chamber and submersed in the spring water. Volumetric rates of gross OP in the mats were measured using the previously described O2- based light-dark shift method (16). Gross AP rates were determined as - 2- -dStot/dt, where the total sulfide concentration (Stot=H2S+HS +S ) was calculated from the measured H2S concentration and pH. As described previously (J. M. Klatt, S. Meyer, S. Häusler, J. L. Macalady, D. de Beer, L. Polerecky; submitted for publication), the variation in pH with time during a light-dark shift measurement was ignored as it was too slow compared with the variation in H2S. During the measurements the tips of the microsensors were separated by <50 μm. First, the light-dark shift measurements were done at a constant irradiance at various depths of the mat to identify the depth with the highest photosynthetic activity. At this depth (0.83 mm) gross OP and AP were subsequently determined at incident irradiance levels between 26 and 340 μmol photons -2 -1 m s . During the measurements H2S was gradually depleted in the mat (Supplementary 256 III. Manuscripts Fig. S1). Therefore, the dark incubation of the mat was occasionally longer to allow for the re-establishment of sulfidic conditions at the depth of measurements. After the light- dark shift measurements, a depth profile of scalar irradiance was measured in the same spot to determine the locally available light. Finally, a sub-sample of the measured mat was examined by microscopy. Calculations Total potential photosynthetic electron transport rates were calculated as a sum of the potential electron transport rates due to OP and AP. The former were calculated by multiplying the gross rates of O2 production measured by the light-dark shift method with + - a factor of 4, as implied by the reaction 2H2O Æ O2 + 4H + 4e . Analogously, the latter were calculated by multiplying the gross rates of Stot depletion measured by the light-dark 0 + - shift method with a factor of 2, as implied by the reaction H2S Æ S + 2H + 2e . Results Microscopy The measured sample of the cyanobacterial mat was dominated by a single filamentous cyanobacterium with a diameter of ~4 μm diameter (Fig. 1). Besides this cyanobacterial morphotype we only observed unicellular non-pigmented prokaryotes. Figure 1: Microscopic images of the cyanobacteria dominating the cyanobacterial mats in the Frasassi sulfidic springs. (A): Light microscope image. (B): Auto-flourescence image (Excitation maximum: 547 nm, Emission >590 nm). The scale bars are 10 μm. 257 7. H2S AND LIGHT CONTROL CYANOBACTERIAL PHOTOSYNTHESIS 8.1 μmol photons m -2 s-1 A 8 ) -1 s -1 6 L - 4 P (μmol e P 2 0 -2 -1 12 12.0 μmol photons m s B 10 ) -1 s 8 -1 L - 6 4 P (μmol e P 2 0 25 26.9 μmol photons m-2 s-1 C 20 ) -1 s -1 L - 15 10 P (μmol e P 5 0 35 37.9 μmol photons m-2 s-1 D 30 ) -1 25 s -1 L - 20 15 P (μmol e P 10 5 AP AP modelled OP OP modelled AP+OP AP+OP modelled 0 0 500 1000 1500 2000 2500 H2S (μM) Figure 2: Volumetric rates of gross photosynthesis in the studied cyanobacterium as a function of the H2S concentration and the locally available scalar irradiance. Shown are rates of anoxygenic (AP) and oxygenic (OP) photosynthesis and their sum (AP+OP), all expressed in μM electrons s-1. Results of the numerical model that fits the experimental data-points are shown by lines. 258 III. Manuscripts Microsensor measurements Scalar irradiance decreased steeply with depth in the cyanobacterial mat, reaching about 10% of the surface value at depth of 0.83 mm (Supplementary Fig. S2). The parallel light- dark shift measurements at this depth revealed that the studied cyanobacteria performed both types of photosynthesis (anoxygenic and oxygenic) simultaneously (Supplementary Fig. S3). While the partitioning between the gross rates of OP and AP depended strongly on the H2S concentration, the total photosynthetic electron transport rate determined at a given local irradiance did not (Fig. 2). Specifically, OP dominated at lower H2S concentrations while AP dominated at higher H2S concentrations, with the threshold H2S concentration above which AP dominated increasing with light (Fig. 2). Importantly, even -2 -1 at very low local light intensities (~2.9 μmol photons m s ) and high H2S concentrations (> 1.7 mM) we always observed simultaneous OP and AP and never only AP (e.g., Fig. 2A). The total potential electron transport rate, i.e., the sum of electron transport rates due to OP and AP, increased linearly in the studied range of the locally available light (Fig. 3). 30 ) -1 s 25 -1 L - 20 15 10 5 AP+OP (μmol e AP+OP 0 010203040 Figure 3: Total photosynthetic electron transport rates in the studied cyanobacterium as a function of the locally available scalar irradiance. The fit of the data-points with a linear model (R=0.9992, P<0.0001) is also shown. Discussion Regulation of OP and AP by H2S and light The studied cyanobacterium performed OP and AP simultaneously over a wide range of H2S concentrations and light intensities. Although the regulation of each photosynthetic mode by H2S and light appeared to be complex, the regulation of the total photosynthetic electron transport given by the sum of electron transports driven by AP and OP was 259 7. H2S AND LIGHT CONTROL CYANOBACTERIAL PHOTOSYNTHESIS astonishingly simple. Specifically, the total electron transport rate increased linearly with irradiance and did not depend on H2S at all. When explaining this pattern, we exclude the effects of H2S on the light- harvesting components in the cell (pigments) and on the photosystem stoichiometry (i.e., the ratio of PSI:PSII in the thylacoid membrane) because all our measurements were performed within a short time (~6 hours) and because the observed pattern emerged even though we did not follow a strict order (e.g., either gradually increasing or gradually decreasing) of light intensities and H2S concentrations. Thus, the rapid adaptation of the studied cyanobacterium to the instantaneous light and H2S conditions in their microenvironment strongly suggests that its photosynthetic electron transport, and specifically the partitioning between oxygenic and anoxygenic photosynthetic modes, is kinetically controlled. We have observed such regulation previously in Pseudanabaena str. FS39, a versatile cyanobacterium enriched from the same environment (8). Specifically, we identified that the partitioning between OP and AP in this cyanobacterium can be explained if the light-driven electron transport from PSII and the H2S-driven electron transport from sulfide:quinone:reductase (SQR) intersect in the plastoquinone (PQ) pool and are controlled by its redox state determined by the relationship between the rates of PQ reduction by PSII (koxy) and SQR (kanoxy) and the rate of PQ oxidation by PSI (kPSI) (Fig. 4A). Based on this model we determined that in Pseudanabaena str. FS39 the apparent affinity of SQR to H2S is very high (KM § 39 μM; see notation in Fig. 4A) and, more importantly, the apparent affinity of SQR to oxidized PQ is at least two orders of magnitude higher than that of PSII (KPSII / KSQR 100). As a consequence, OP in Pseudanabaena str. FS39 is kinetically outcompeted by AP, which is manifested by the fact that OP is performed only in addition to AP at high light intensities or when H2S is limiting (Fig. 4B). When applying the same model for the cyanobacterium in this study, we found an excellent agreement between the measured and predicted rates of OP and AP and their light- and H2S-dependent partitioning (compare symbols with lines in Fig. 2). However, in contrast to Pseudanabaena str. FS39, we found that to fit the experimental data for the cyanobacterium studied here the apparent affinity of SQR to H2S must be much lower (KM § 6.5 mM) while its apparent affinity to oxidized PQ must be very similar to that of the PSII (KPSII / KSQR § 1). As a consequence, OP is kinetically competitive against AP and thus performed simultaneously with AP irrespective of the light intensity and H2S 260 III. Manuscripts concentration (Fig. 4C), even at H2S concentrations that are higher than ever observed in- situ in the naturally occurring mats (J. M. Klatt, S. Meyer, S. Häusler, J. L. Macalady, D. de Beer, L. Polerecky; submitted for publication). A IPSII IPSI PQox PQ red H O koxy = IPSII k = I 2 + PSI PSI + K PSII PQox K PSI PQ red PSI I PQ PSI O2 PQ k = k ox anoxy H 2S + [H S] K SQR PQ ox k = v 2 H 2S max + K M [H 2S] H2S SQR S0 120 K = 39 μM K / K = 200 B Pseudanabaena FS39 M PSII SQR 100 80 60 P (%) P 40 20 0 120 C Spring cyanobacterium KM = 6500 μM KPSII / KSQR = 1 100 80 2 μM H S OP 2 2 μM H S AP 60 2 P (%) P 40 1000 μM H S OP 2 1000 μM H S AP 20 2 0 0 100 200 300 400 500 600 700 -2 -1 Scalar irradiance (μmol photons m s ) Figure 4: (A) Simplified diagram of the model describing partitioning between AP and OP in versatile cyanobacteria, as proposed by Klatt and co-workers (8). The AP and OP pathways intersect at plastoquinone (PQ) and their partitioning is regulated by the PQ redox state. In the AP pathway, PQ is reduced in a light-independent reaction by the enzyme sulfide:quinone:reductase (SQR), which obtains the electrons from the oxidation of H2S to zero-valent sulfur. In contrast, PQ reduction in the OP pathway is driven by the light-dependent activity of photosystem II (PSII), with electrons originating from the oxidation of H2O to O2. In both pathways PQ oxidation is driven by the light-dependent activity of photosystem I (PSI). IPSI and IPSII denote light energy harvested in PSI and PSII, respectively. The rate-laws describing the kinetic regulation of the rates of OP (koxy), AP (kanoxy), H2S oxidation by SQR (kH2S) and of the total photosynthetic electron transport (kPSI) are shown by the formulae. Partitioning between AP and OP predicted by the model for Pseudanabaena str. FS39 and for the cyanobacterium studied here at low and high H2S concentrations is shown in panels B and C, respectively. The reason for the striking differences between Pseudanabaena str. FS39 and the dominant spring cyanobacterium might be that their SQR enzymes belong to different enzyme structure classes and therefore have different affinities to PQ and H2S (17). As the diffusion time of PQ between PSII and the cytochrome b6f complex in the membrane 261 7. H2S AND LIGHT CONTROL CYANOBACTERIAL PHOTOSYNTHESIS is the rate limiting step of the oxygenic photosynthetic electron transport, the apparent affinity of SQR and its competitiveness with PSII might additionally be affected by differences in diffusion time of SQR, which is also membrane associated (18). For example, clustering of SQR, PQ and the cytochrome b6f complex would make it possible for the reactions between these three components to occur very fast. The relative position of PSII to the other electron transport components would in turn then also affect the apparent affinity of PSII. Further research is required to explore these hypotheses. Implications for the competition among phototrophs in sulfidic environments When studying selective advantages of an organism performing a certain process, thermodynamic considerations often provide the first level of understanding. While AP is driven by only one photosystem (PSI), OP is driven by two photosystems (PSI+PSII). Thus, theoretically, the photon flux required to drive a certain electron transport and growth rate is twice as large for OP than for AP. Due to this lower energy demand of AP, one could expect that AP is favourable for cyanobacteria when H2S is not limiting. However, this expected advantage of AP seems not to necessarily hold true in versatile cyanobacteria. For the studied cyanobacterium and also for Pseudanabaena str. FS39 we have shown that the total electron transport rate driven by a specific photon flux is independent of H2S concentrations and the photosynthetic mode. A constant electron transport implies that also the quantum efficiency is constant when calculated in electrons per photons. Thus, the light energy potentially harvested in PSII is basically wasted due to the partitioning between AP and OP being limited by the kinetics of the electron transport by PSI. AP would only be advantageous if the excitation energy were transferred from PSII to PSI or if the photosystem stoichiometry changed (8). Since this does not seem to occur in the cyanobacterium dominating the Frasassi sulfidic springs, AP does not provide a thermodynamic advantage to this phototroph. Considering that quantum efficiency in cyanobacteria is constant irrespective of the photosynthetic mode, obligate oxygenic phototrophic cyanobacteria can, from a thermodynamic perspective, theoretically be as successful as versatile cyanobacteria. Although obligate oxygenic phototrophs are present in the Frasassi sulfidic springs (e.g., Planktothrix str. FS34,(10)), they do not dominate the photosynthetic community. A possible reason could be toxicity of H2S for PSII. Although we have found that Planktothrix str. FS34 is temporarily resistant against H2S, this resistance is apparently not sufficient for success in this environment. This is possibly because the niche based on 262 III. Manuscripts temporary H2S-resistance is very narrow (10), making this cyanobacterium vulnerable to as minor environmental fluctuations as a change between sunny and cloudy days. Photosynthetic versatility and the associated active depletion of sulfide and production of oxygen appear to play a much more important role in the sulfidic springs setting. First, versatile cyanobacteria are generally never limited by electron donor availability. Second, by performing OP they additionally produce O2, which is toxic to most obligate anoxygenic phototrophs. Together, this provides them with a clear competitive advantage over obligate anoxygenic phototrophs. Still, this does not explain the dominance of the studied cyanobacterium in the Frasassi sulfidic spring mats over Pseudanabaena str. FS39, which is also a photosynthetically versatile cyanobacterium. We suggest that here the different apparent affinities of SQR and PSII towards the oxidized PQ play the critical role. Specifically, due to the much higher affinity of SQR towards oxidized PQ in Pseudanabaena str. FS39, this cyanobacterium performs primarily AP and switches to OP only when H2S becomes limiting, e.g., at high light (8) (Fig. 4B). This decreases their competitiveness against anoxygenic phototrophs, because their production of O2 is limited to a short interval during the day or might not occur at all. In contrast, the affinities of SQR and PSII towards the oxidized PQ in the cyanobacterium studied here are very similar, which makes it possible for them to perform simultaneous AP and OP at all times during the diurnal cycle. Additionally, their AP actively removes H2S, which leads to a higher O2 production because the rate of OP increases relative to the rate of AP with decreasing H2S concentrations (Fig. 2). Together, this increases their competitiveness against anoxygenic phototrophs relative to Pseudanabaena str. FS39 and thus leads to their dominance in the system. Evolutionary implications The capability to perform OP at very low light and high H2S concentrations might have provided the ancestors of contemporary cyanobacteria with a crucial advantage in the microbial mats, where OP evolved. Also in the mid-end Proterozoic oceans, which were characterized by abundant H2S in the photic zone of the water column, versatile cyanobacteria that produce toxic O2 while removing H2S might have played an essential role in the evolution of the Earth. Specifically, by depleting H2S, the indispensable substrate for obligate anoxygenic phototrophs, they might have sustained O2 production and eventually shifted the balance from AP to OP, thus contributing to the oxygenation of the atmosphere and oceans. 263 7. H2S AND LIGHT CONTROL CYANOBACTERIAL PHOTOSYNTHESIS Acknowledgements We thank the technicians of the microsensor group for the construction of microsensors, J. Macalady and D. S. Jones for their support during field campaigns and A. Montanari and P. Metallo for their hospitality during the sampling campaign at the Osservatorio Geologico di Coldigioco. This work was financially supported by the Max-Planck- Society. References 1. Blankenship RE. 2010. Early evolution of photosynthesis. Plant Physiol. 154:434–8. 2. Mulkidjanian AY, Koonin E V, Makarova KS, Mekhedov SL, Sorokin A, Wolf YI, Dufresne A, Partensky F, Burd H, Kaznadzey D, Haselkorn R, Galperin MY. 2006. The cyanobacterial genome core and the origin of photosynthesis. Proc. Natl. Acad. Sci. U. S. A. 103:13126–31. 3. Buick R. 2008. When did oxygenic photosynthesis evolve? Philos. Trans. R. Soc. Lond. B. Biol. Sci. 363:2731–2743. 4. Nisbet EG, Fowler CMR. 1999. Archaean metabolic evolution of microbial mats. Proc. R. Soc. B Biol. Sci. 266:2375–2382. 5. Canfield DE. 1998. A new model for Proterozoic ocean chemistry. Nature 396:450–453. 6. Johnston DT, Wolfe-Simon F, Pearson A, Knoll AH. 2009. Anoxygenic photosynthesis modulated Proterozoic oxygen and sustained Earth’s middle age. Proc. Natl. Acad. Sci. U. S. A. 106:16925–16929. 7. Miller SR, Bebout BM. 2004. Variation in sulfide tolerance of photosystem II in phylogenetically diverse cyanobacteria from sulfidic habitats. Appl. Environ. Microbiol. 70:736–44. 8. Klatt JM, Al-Najjar MAA, Yilmaz P, Lavik G, de Beer D, Polerecky L. 2015. Anoxygenic photosynthesis controls oxygenic photosynthesis in a cyanobacterium from a sulfidic spring. Appl. Environ. Microbiol. AEM.03579–14. 9. Galdenzi S, Cocchioni M, Morichetti L, Amici V, Scuri S. 2008. Sulfidic ground-water chemistry in the Frasassi caves, Italy. J. Cave Karst Stud. 70:94–107. 10. Klatt JM, Haas S, Yilmaz P, de Beer D, Polerecky L. H2S can inhibit and enhance oxygenic photosynthesis in a cyanobacterium from sulphidic springs. Environ. Microbiol. , in press. 11. Revsbech NP. 1989. An oxygen microsensor with a guard cathode. Limnol. Oceanogr. 34:474–478. 264 III. Manuscripts 12. Jeroschewski P, Steuckart C, Kühl M. 1996. An Amperometric Microsensor for the Determination of H2S in Aquatic Environments. Anal. Chem. 68:4351–4357. 13. De Beer D, Schramm A, Santegoeds CM, Kühl M. 1997. A nitrite microsensor for profiling environmental biofilms. Appl. Environ. Microbiol. 63:973–977. 14. Lassen C, Ploug H, Jørgensen BB. 1992. A fibre-optic scalar irradiance microsensor: application for spectral light measurements in sediments. FEMS Microbiol. Lett. 86:247–254. 15. Al-Najjar MAA, de Beer D, Jørgensen BB, Kühl M, Polerecky L. 2010. Conversion and conservation of light energy in a photosynthetic microbial mat ecosystem. ISME J. 4:440–449. 16. Revsbech NP, Jørgensen BB. 1983. Photosynthesis of benthic microflora measured with high spatial resolution by the oxygen microprofile method: capabilities and limitations of the method. Limnol. Oceanogr. 28:749–759. 17. Gregersen LH, Bryant DA, Frigaard N-U. 2011. Mechanisms and evolution of oxidative sulfur metabolism in green sulfur bacteria. Front. Microbiol. 2:116. 18. Shahak Y, Hauska G. 2008. Sulfide oxidation from cyanobacteria to humans: sulfide–quinone oxidoreductase (SQR), p. 319–335. In Hell, R, Dahl, C, Knaff, D, Leustek, T (eds.), Sulfur Metabolism in Phototrophic Organisms. Springer Netherlands, Dordrecht. 265 7. H2S AND LIGHT CONTROL CYANOBACTERIAL PHOTOSYNTHESIS REGULATION OF THE PARTITIONING BETWEEN OXYGENIC AND ANOXYGENIC PHOTOSYNTHESIS IN A CYANOBACTERIUM DOMINATING SULFIDIC SPRING MICROBIAL MATS Klatt, Judith M.; Häusler, Stefan; de Beer, Dirk; Polerecky, Lubos Supplementary Information H S (mM) 2 02468 -1.0 -0.5 0.0 0.5 1.0 Depth (mm) Depth 1.5 2.0 0 100 200 300 400 O (μM) 2 Figure S1: Examples of transient depth profiles of O2 and H2S concentration measured in the cyanobacterial mat during illumination at incident irradiance of 169 μmol photons m-2 s-1. The profiles shown with open and filled symbols were separated by about 2 hours. 266 III. Manuscripts scalar irradiance (%) 110100 0.0 0.2 0.4 0.6 depth (mm)depth 0.8 1.0 0 20406080100120 scalar irradiance (%) Figure S2: A depth profile of local scalar irradiance in the studied cyanobacterial mat. The scalar irradiance values were integrated over the wavelengths of photosynthetically active radiation (400 – 700 nm) and normalized to the value at the mat surface. Error bars represent the standard deviation of measurements in 3 replicate spots of the mat. Note that the filled and open symbols show the same profile in linear and logarithmic scale, respectively. 100 1250 75 1000 (μM) S (μM) 2 50 2 O H 750 25 0 500 0 60 120 180 240 time (s ) Figure S3: O2 and H2S concentrations recorded in the studied cyanobacterial mat during light- dark shift measurements. Measurements were conducted 0.83 mm below the mat surface with microsensor tips separated by <50 μm. During the light intervals (white areas) the local scalar irradiance was 18.8 μmol photons m-2 s-1. Shaded areas indicate dark intervals. 267 268 III. Perspective 269 270 III. Perspective 8. DISCUSSION The persistence of partially sulfidic oceans during the Proterozoic inspired Johnston et al (2009) to set up a model for biogeochemical cycling, which identified anoxygenic photosynthesis (AnoxyP) as the driver sustaining euxinia and tempering the net release of O2 release by oxygenic photosynthesis (OxyP) (section 1.1). To date, there is no alternative comprehensive model available that would explain the sustainment of oxygen deficiency for a time period as long as 1 – 2 billion years after the great oxidation event (GOE). Johnston et al. speculated that photosynthetically versatile cyanobacteria may have played an important role, however their activity is not explicitly accounted for in the model. Similarly, by considering extant microbial mats, Oren et al. (1977) and Cohen et al. (1986) among others have speculated that the evolution of AnoxyP might have predated the onset of OxyP in the cyanobacterial ancestor, and regard versatility as an important adaptation strategy to cope with the possibly reducing conditions in ancient microbial mat like structures. As OxyP likely evolved in such structures, these represent the strategic centers for the global oxygenation of the biosphere by cyanobacterial activity. Thus, if the Cohen et al.,Oren et al. and Johnston et al. hypotheses are right, cyanobacterial AnoxyP might have been a process of global importance; substantially contributing to shaping the history of Earth. Currently we cannot directly infer the history of cyanobacterial AnoxyP based on molecular tools or signatures left in the geological records (section 1.2). It is therefore crucial to further explore the selective advantage of versatile cyanobacteria in contemporary ecosystems to be able to evaluate if they might have played an important role in ancient systems. One possible approach is to study their activity in ecosystems that are analogous to the ancient systems that we wish to explore. The phototrophic microbial mats distributed along the flow path of the Frasassi sulfidic spring water and the Little Salt spring mats represent such ecosystems and can be considered as ancient Earth analogues (Chapter 2 and 5). Intriguingly, irrespective of the low O2 concentrations and partially high sulfide concentrations, both the Little Salt Spring mats and Frasassi sulfidic spring mats are dominated by photosynthetically versatile cyanobacteria while obligate anoxygenic and obligate oxygenic phototrophs are almost completely absent. Thus, the capability to perform both OxyP and AnoxyP indeed seems to be a key to success. However, it remains unclear what exactly the selective advantage of versatility over more specialized phototrophy is. 271 8. DISCUSSION In the following, first the general selective advantages of OxyP, AnoxyP and photosynthetic versatility are discussed and the consequences of cyanobacterial activity for other microbially mediated processes are further explored using the microbial mats forming at the bottom of the Frasassi sulfidic springs as an example. Secondly, specific adaptation strategies among extant cyanobacteria are analyzed and compared. Finally, as a synthesis of this study, possible implications in the context of current models of ancient ocean chemistry and early stromatolites are discussed. 8.1. The advantage of photosynthetic versatility Photosynthetically versatile cyanobacteria are capable of both OxyP and AnoxyP. During AnoxyP they oxidize sulfide to an incompletely reduced sulfur compound, mainly S0. In the environment they have to compete with other phototrophs, such as obligate oxygenic phototrophs and obligate anoxygenic phototrophs. The latter functional group of phototrophs can oxidize sulfide completely to sulfate or to intermediately reduced sulfur compounds that other phototrophs can further oxidize (section 1.3). We have shown in Chapter 2 that other autotrophic organisms, namely aerobic sulfide oxidizers, can also compete with versatile cyanobacteria. This type of competition and the role of versatility in determining its outcome, is however not discussed in the following. So far, there are three main hypothesizes concerning the advantage of photosynthetic versatility: (A) The rapid removal of H2S mainly serves a detoxification purpose (Jørgensen et al., 1986; de Wit and van Gemerden, 1987; Stal, 2002). (B) AnoxyP is energetically more favourable than OxyP. (C) Obligate anoxygenic phototrophs are directly out-competed by rapid sulfide removal (Stal, 2002). In the following these hypotheses are explored and subsequently a novel hypothesis built upon (C) is offered that tries to explain the success of versatile cyanobacteria by their modulation of the sulfur cycle. 8.1.1. “Detoxification” hypothesis It has been repeatedly suggested that, at least in some cyanobacterial species, AnoxyP is mainly used to remove the potentially toxic H2S. For cyanobacteria that cannot grow by AnoxyP (Padan and Cohen, 1982) this is inarguable. For other species, AnoxyP was suggested to be a detoxification mechanism because the organisms exhibited a very low 272 III. Perspective apparent affinity to H2S, which is interpreted such that they are not competitive with obligate anoxygenic phototrophs for sulfide (Wit and Gemerden, 1987). To consider AnoxyP as a simple detoxification mechanism, however, also comes with the implication that the use of H2S as an electron donor is not advantageous in itself. If this were the case, the main selective advantage gained by versatile cyanobacteria would be if AnoxyP was only used to allow OxyP to be performed at its maximal rate and duration. However, as shown in Chapter 4, there are also obligate oxygenic phototrophs that can perform OxyP in the presence of sulfide and even increase their photosynthetic rates upon exposure to H2S. Thus, they do not have to temporarily lower their O2 production rate whilst performing AnoxyP. Also, as H2S appears to be the toxic sulfide species, an active removal of sulfide is not crucial for detoxification. Instead the H2S concentration can also be modulated indirectly by performing OxyP, which results in pH shifts (Chapter 4). Therefore, the use of AnoxyP, which even involves de novo synthesis of proteins, as a mere detoxification mechanism would likely not lead to an increase in fitness in the environment unless other advantages of active photosynthetic sulfide removal are being considered. 8.1.2. “Energetics” hypothesis The energetics hypothesis is based on calculations of the Gibbs free energy of the reactions performed during OxyP and AnoxyP. Using the redox midpoint potentials of the involved redox carrier couples and electron donors listed in Table 1 (section 1.2.) (e.g. Alberty, 2003), such calculation yields ~15 kJ (mol NADP+)-1 and ~219 kJ (mol NADP+)-1 for the cyanobacterial AnoxyP and OxyP, respectively. First of all, it is worthwhile mentioning that these thermodynamic calculations are performed at standard physiological conditions. However, the activity (often approximated by the concentration) of the reactants would actually have to be considered. The effect of an adjustment of the calculation would be most pronounced for AnoxyP, as it depends on the concentration of H2S. The calculated energy requirement therefore increases, and consequently also the apparent thermodynamic advantage vanishes, with decreasing H2S availability. The question is, however, if the cyanobacteria exploit the apparent thermodynamic advantage of AnoxyP when H2S concentration is not limiting. To answer this question, the photosynthetic light harvesting and electron transport processes during AnoxyP and OxyP have to be understood. Oxygenic phototrophs employ two photosystems that operate in series. Thus, two photons (one in PSII and one 273 8. DISCUSSION in PSI) are used to drive the transport of one electron. In contrast, for AnoxyP the absorption of only one photon in one photosystem is required to transport one electron derived from the oxidation of H2S. This can indeed be considered an energetic advantage. However, as was shown in Chapter 6 and 7, not all versatile cyanobacteria seem to exploit this advantage because the electron transport rate per absorbed photon is not necessarily higher after a switch from OxyP to AnoxyP. This is because photons are still absorbed in PSII. Their energy is, however, not used for NADPH reduction and is “lost”. In the linearly organized photosynthetic electron transport chain the photosystems are provided with light energy from almost separate light harvesting accessory pigments or pigment complexes. Such design only allows for limited transfer of excitation energy between the photosystems during state transition (Chapter 6). A full exploitation of the advantage of the theoretical photon:electron ratio of 1 would therefore only be achieved by a complete replacement of PSIIs by PSIs upon switching from OxyP to AnoxyP, which has never been observed. Photosynthetic electron transport involves redox carrier (NADP+) reduction that is tightly coupled to proton translocation, thus generating a proton motive force (pmf) that can then be used for ATP synthesis. An absorbed photon therefore provides both the energy for redox carrier reduction and ATP generation. In other words, a fraction of the energy that is not used for NADP+ reduction is utilized to generate a proton motive force. During oxygenic photosynthetic electron transport, about 3 H+ per electron are translocated across the membrane, which results in the production of 0.75 ATPs assuming that ATP synthase generates 0.25 ATP per H+ (Steigmiller et al., 2008). When normalized to an absorbed photon instead of an electron however, only 0.375 ATP per photon are produced. It is not yet resolved how many protons are translocated during cyanobacterial AnoxyP (Griesbeck et al., 2000). Even if during the H2S oxidation reaction protons are not translocated across the membrane, AnoxyP results in 0.5 ATP per photon absorbed in PSI (Chapter 6). This means that also 0.5 ATP per electron are generated. Thus, the energy conserved as ATP per photon absorbed in PSI is higher during AnoxyP. Interestingly, this high ATP output per photon also implies that AnoxyP in versatile Cyanobacteria is actually as efficient as in obligate anoxygenic phototrophs – if the “loss” of photon energy harvested in PSII is not considered. However, in comparison to OxyP, the ATP yield per electron transported might be lower compared to OxyP unless the H2S oxidation step contributes to pmf generation. 274 III. Perspective Overall, when calculating the energy requirements of AnoxyP at physiological standard conditions, it appears to be the thermodynamically more favourable process. Indeed, during AnoxyP more ATP and NADPH can be generated per photon absorbed in PSI. However, excitation energy harvested in PSII is “lost” and therefore rates of NADP+ reduction cannot be increased unless excitation transfer occurs or the photosystem stoichiometry is changed. Also, during AnoxyP the ATP yield per electron transported might be lower compared to OxyP. Still, it should be noted that the mere energy gain during photosynthetic electron transport is not necessarily linked to success in the environment (Chapter 3). Both quantum efficiency and the actual rate of photosynthesis shape the fitness of the phototrophs under a certain set of conditions. Both these factors are generally rather the result of cooperative tuning of the kinetics of all different redox reaction steps than a matter of the efficiency of ATP generation during the individual reactions. 8.1.3. “Direct competition” hypothesis Stal (2002) suggested that there might be a direct competition between obligate anoxygenic phototrophs and versatile cyanobacteria for sulfide. This is a competition that might not necessarily be won by cyanobacteria as their apparent affinity for H2S covers a wide range. However, it is also suggested that due to extensive production of S0 by the versatile cyanobacteria in the morning, this product would not be available for further phototrophic oxidation in the aerobic layers of a mat. By pointing towards the importance of the product of cyanobacterial AnoxyP, Stal (2002) raises an interesting thought that needs further exploration, which will be done in the following. 8.1.4. “Modulation” hypothesis The governing hypothesis of this section is that cyanobacterial AnoxyP indirectly outcompetes obligate anoxygenic phototrophs by modulating the S-cycle. Namely, the 0 simultaneous production of S and O2 is expected to have a negative feedback effect on sulfide production rates, which tempers AnoxyP by obligate anoxygenic phototrophs. To arrive at this hypothesis several feedback mechanisms, such as the dependence of the sulfide production rate on the availability of organic material required for sulfate reduction, need to be considered. To be able to account for these factors that impact the competition among phototrophs in the following a simplified model of the sulfur, carbon and oxygen cycle in microbial mats is introduced. To arrive at an understanding of the 275 8. DISCUSSION feedback effects of cyanobacterial activity on possible competitors, this model is first used to identify additional factors that, in addition to external electron donor availability, likely shape the maximal rate of AnoxyP and thus its competitiveness. Then, it is derived how obligate oxygenic phototrophs likely impact the sulfur, carbon and oxygen cycle. This is finally contrasted to the impact of photosynthetically versatile cyanobacteria. 8.1.4.1. An efficiency-based model of the C-, S- and O-cycle in microbial mats This section briefly introduces a model of the C-, S- and O-cycle in microbial mats and outlines the major assumptions that were made. Afterwards the model is used to identify possible factors that might affect the outcome of the competition among obligate anoxygenic phototrophs, obligate oxygenic phototrophs and versatile cyanobacteria in a given environment. The model as depicted in Figure 8.1 is based on the substrate utilization efficiencies of the major processes involved in C-, S- and O-cycle in microbial mats. For instance, OxyP generates a given flux of O2, a fraction of which is escaping into the water column. Here, this fraction is defined by İO2 (Fig. 8.1, Table 1). The remainder of the O2 produced by OxyP is consumed by aerobic respiration at a rate that is characterized by a utilization efficiency İAR. The still remaining fraction ((1 – İAR) İO2 JO2,OP; Table 1) and any O2 available from an external source (JO2,ext; Table 1), is utilized by chemolithotrophic reduced sulfur oxidation (SOX). The model differentiates between three possible SOX pathways: aerobic S0 oxidation, aerobic sulfide oxidation to zero-valent sulfur and aerobic sulfide oxidation to sulfide. The utilization of O2 by these three processes is again dependent on substrate utilization efficiencies. Overall, the model is designed such that each process rate can be calculated from given values for the substrate utilization efficiencies and given external fluxes of O2 and sulfide, and the rate of OxyP. It should be noted that the substrate utilization efficiency represents the descriptor of the competitiveness. For instance, the partitioning of the sulfide flux between cyanobacterial AnoxyP and AnoxyP by obligate anoxygenic phototrophs is exclusively dependent on İCYA. One of the main assumptions in the model is that organisms performing AnoxyP are not competitive against sulfide oxidizing bacteria (SOB) (van Gemerden (1993); see section 1.3). Sulfide is consumed by SOX processes at a rate that is exclusively dependent on the O2 flux and the energy conservation efficiency of the SOB (Chapter 3, represented by the factors ySO4 and yS0 in the notions for JS,SOX,H2S->SO4 and JS,SOX,H2S->S0, respectively; 276 III. Perspective Table 8.1). The utilization efficiency İSOX determines how the O2 and sulfide fluxes are distributed among the sulfide-utilizing SOX processes that compete for it. The remaining fraction of the sulfide flux, which is available for other organisms such as obligate anoxygenic phototrophs, is therefore not calculated based on sulfide utilization efficiency. Thus, the model does not account for competition between SOX and AnoxyP and assumes that SOX processes always “outcompete” AnoxyP for sulfide if O2 is abundant. OP AP H2S SO4 HO2 H H AP H AR H2S APH2S S0 AR HCya CYAH2S S0 SOX H2S S0 HSOX SOX H2S SO4 HS0 HSOX,S0 HAP,S0 APS0 SO4 SOXS0 SO4 SRS0 H2S HSR HCorg SRSO4 H2S Figure 8.1: Proposed model for the S-, C- and O-cycle in microbial mats. The arrows represent the fluxes of compounds utilized or produced by microbially mediated processes indicated in the boxes. APH2S->SO4 and APH2S->S0 refer to AnoxyP by obligate anoxygenic 2- 0 phototrophs that oxidize sulfide to SO4 and S , respectively. CYA refers to cyanobacterial 0 0 AnoxyP yielding S from the oxidation of sulfide. APS0->SO4 is the S -driven AnoxyP performed 0 2- by obligate anoxygenic phototrophs. SRS0->H2S and SRSO4->H2S refers to S and SO4 reduction, respectively, coupled to organic carbon (Corg) oxidation. OP is OxyP. AR is aerobic respiration using O2 and Corg provided by OxyP. SOXH2S->S0 and SOXH2S->SO4 refer to incomplete and 0 complete aerobic sulfide oxidation, respectively. SOXS0->SO4 is aerobic S oxidation. If the SOX processes are performed by autotrophic SOB, the reduced sulfur oxidation is coupled to both the reduction of O2 and CO2, which yields Corg (indicated by the dashed arrows. The efficiencies of substrate utilization (İi) of the different processes are also indicated. The corresponding definitions of the fluxes are shown in Table 8.1. 277 8. DISCUSSION Furthermore, the model implicitly assumes that that in an ecosystem driven by the external supply of reductant, OxyP is not needed, unless electron donor supply becomes the limiting factor for anoxygenic primary productivity. The reasoning behind this assumption is the apparent energetic advantage of obligate anoxygenic phototrophs over oxygenic phototrophs. In the model OxyP is therefore a superimposed process, i.e., it is a given input parameter, and can be chosen to increase until total productivity is limited. In the current form of the model the maximal OxyP can be defined. The model does not consider the possibility of an inhibitory effect of O2 and H2S on the rates of AnoxyP and OxyP, respectively. Also, a possible decrease in light availability for photosynthetic processes due to increased activity and biomass production by other forms of photosynthesis or SOX processes (as observed in B/C mats in the Frasassi sulfidic springs; Chapter 2) is not incorporated. Table 8.1: Definition of the fluxes shown in Figure 8.1. Expression Description JO2,OP Rate of O2 production by oxyP JO2,AR = İAR İO2 JO2,OP Rate of O2 consumption by aerobic respiration (AR), which depends on the rate of OxyP, a fraction of which defined by İO2 escapes the mat system and the O2 utilization efficiency of AR İAR 0 JO2,SOX,S0->SO4 = 1.5y İS0 İSOX,S0 (JS0,AP, H2S->S0 Rate of O2 consumption by aerobic S oxidation. A 0 + J + J ) part, defined by (1 – İS0), of the S produced by S0,SOX,H2S->S0 S0,CYA obligate anoxygenic phototrophs, aerobic sulfide oxidizers and cyanobacteria at rates JS,AP, H2S->S0, JS,SOX,H2S->S0 and JS,CYA, respectively, escapes further oxidation or re-reduction. The remainder of this S0 flux is used by aerobic S0 oxidation at a utilization efficiency İSOX,S0. Dependent on the fluxes 2 energy conservation efficiency of the aerobic S0 O oxidizers, a fraction 1.5y of the utilized S0 is used for O2 reduction (Chapter XXX). JO2,SOX,H2S->SO4 = İSOX ((1 – İAR) İO2 JO2,OP Rate of O2 consumption by aerobic sulfide 2- oxidation to SO4 , which is fuelled by the external + JO2,ext – JO2,SOX,S0->SO4) O2 flux (JO2,ext) and by the remainder of the photosynthetically produced O2 (JO2,OP), which was not consumed by AR and aerobic S0 oxidizers. The partitioning of this total available O2 flux between 2- 0 SOX yielding SO4 and S is determined by İSO. JO2,SOX,H2S->S0 = (1 – İSOX) ((1 – İAR) İO2 JO2,OP Rate of O2 consumption by aerobic sulfide oxidation to S0. + JO2,ext – JO2,SOX,S0->SO4) 278 III. Perspective Table 8.1 (continued) Expression Description JC,OP = JO2,OP Rate of organic carbon (Corg) production by oxyP 0 1 y Rate of Corg production by aerobic S J C,SOX,S0-!SO4 J O2,SOX,S0-!SO4 oxidation, which can be calculated y from the rate of O2 consumption by aerobic S0 oxidation using the parameter y, which described the partitioning of the S0 pool between O2 and CO2 reduction. For non- autotrophic SOB, for instance, y would be equal to 1 (Chapter XXX) 1 y Rate of Corg production by aerobic J SO4 J 2- C,SOX,H2S-!SO4 O2,SOX,H2S-!SO4 sulfide oxidation to SO4 , which is ySO4 calculated from the rate of O2 consumption JO2,SOX,H2S->SO4 using the parameter ySO4. 1 y Rate of Corg production by aerobic S0 2- J C,SOX,H2S-!S0 J O2,SOX,H2S-!S0 sulfide oxidation to SO4 , which is yS0 calculated from the rate of O2 consumption JO2,SOX,H2S->S0 using the parameter yS0. JC,CYA = 0.5 İCYA (JS,ext + JS,SR,SO4->H2S + JS,SR,S0->H2S Rate of Corg production during fluxes – JS,SOX,H2S->S0 – JS,SOX,H2S->S0) cyanobacterial AnoxyP. Sulfide is org produced by SR processes (JS,SR,SO4- C >H2S and JS,SR,S0->H2S) and derived from an external source (JS,ext). A part of this available sulfide flux is consumed by SOX (JS,SOX,H2S->S0 and JS,SOX,H2S->S0). The remainder is exploited by cyanobacterial AnoxyP at a utilization efficiency İCYA and oxidized to S0. The oxidation is coupled to Corg production (H2S:Corg= 0.5). JC,AP,H2S->SO4 = 2 (1 – İCYA) İH2S İAP (JS,ext + JS,SR,SO4->H2S Rate of Corg production by AnoxyP 2- + J – J – J ) oxidizing sulfide to SO4 . The S,SR,S0->H2S S,SOX,H2S->S0 S,SOX,H2S->S0 fraction of the available sulfide flux not consumed by SOX and versatile cyanobacteria is used by obligate anoxygenic phototrophs at a utilization efficiency İH2S. The distribution of sulfide utilization among anomxygenic phototrophs 0 2- producing S or SO4 is dependent on İAP. JC,AP,H2S->S0 = 0.5 (1 – İCYA) İH2S İAP (JS,ext + JS,SR,SO4->H2S Rate of Corg production by AnoxyP oxidizing sulfide to S0. + JS,SR,S0->H2S – JS,SOX,H2S->S0 – JS,SOX,H2S->S0) 279 8. DISCUSSION Table 8.1 (continued) Expression Description JC,AP,S0->SO4 = 1.5 İS0 (1 – İSOX,S0) İAP,S0 (JS0,AP,H2S->S0 Rate of Corg production by AnoxyP 0 2- oxidizing S to SO4 . A part, + JS0,SOX,H2S->S0 + JS0,CYA) 0 defined by (1 – İS0), of the S produced by obligate anoxygenic phototrophs, aerobic sulfide oxidizers and cyanobacteria at rates JS,AP, H2S->S0, JS,SOX,H2S->S0 and JS,CYA, respectively, escapes further oxidation or re-reduction. The remainder of this S0 flux is used by aerobic S0 oxidation at a utilization efficiency İSOX,S0. The distribution of the still remaining fraction among anoxygenic phototrophs or 0 S reducers is defined by İAP,S0. fluxes JC,tot= JC,OP + JC,SOX,H2S->SO4 + JC,SOX,H2S->S0 + JC,SOX,S0->SO4 Rate of total CO2 fixation. org C + JC,AP,H2S->SO4 + JC,AP,H2S->S0 + JC,AP,S0->SO4 0 JC,SR,S0->H2S = 1.5 İS0 (1 – İSOX,S0) (1 – İAP,S0) Rate of Corg consumption by S reducers. (JS0,AP,H2S->S0 + JS0,SOX,H2S->S0 + JS0,CYA) 2- JC,SR,SO4->H2S = İCorg (JC,tot – İAR JC,OP – JS,SR,S0->H2S) Rate of Corg consumption by SO4 reducers. Any Corg produced by the processes listed above that is not consumed by aerobic respiration or S0 reduction is used at an efficiency İCorg. The remainder, defined by (1 – İCorg), escapes remineralisation. JC,AR = JO2,AR Rate of Corg consumption by aerobic respiration (AR) 1 Rate of sulfide consumption by J J 2- S,SOX,H2S-!SO4 O2,SOX,H2S-!SO4 aerobic sulfide oxidation to SO4 . 2ySO4 1 Rate of sulfide consumption by J J 2- S,SOX,H2S-!S0 O2,SOX,H2S-!S0 aerobic sulfide oxidation to SO4 . 0.5yS0 1 fluxes J J Rate of sulfide consumption during tot S,CYA C,CYA S 0.5 cyanobacterial AnoxyP. 1 JS,AP,H2S-!SO4 J C,AP,H2S-!SO4 Rate of sulfide consumption by 2 2- AnoxyP oxidizing sulfide to SO4 . 1 JS,AP,H2S-!S0 J C,AP,H2S-!S0 Rate of sulfide consumption by 0.5 AnoxyP oxidizing sulfide to S0. 280 III. Perspective Table 8.1 (continued) Expression Description 1 J J 0 S0,SOX,S0-!SO4 1.5y O2,SOX,S0-!SO4 Rate of S consumption by aerobic S0 oxidation Rate of S0 production by aerobic JS0,SOX,H2S-!S0 JS,SOX,H2S-!S0 sulfide oxidation to S0. JS0,CYA JS,CYA Rate of S0 production during cyanobacterial AnoxyP. fluxes 0 0 JS0,AP,H2S->S0 = JS,AP,H2S->S0 Rate of S production by AnoxyP S oxidizing sulfide to S0. 1 0 J J Rate of S consumption by S0,AP,H2S-!SO4 C,AP,S0-!SO4 0 2- 1.5 AnoxyP oxidizing S to SO4 . 1 Rate of S0 consumption by S0 JS0,SR,S0-!H2S J C,SR,S0-!H2S 1.5 reduction. 8.1.4.2. Microbial mats driven by anoxygenic photosynthesis As opposed to OxyP, AnoxyP relies on the supply of a suitable electron donor, such as sulfide, which is supplied from an external source, e.g. by the influx from the water column, and by remineralisation of the organic matter provided by AnoxyP. Thus, the rates of AnoxyP do not only depend on light and external supply of electron donor. This is because there is a feedback effect of AnoxyP on its own rate: if there is less AnoxyP, less biomass is generated, less SR can produce sulfide by oxidizing this biomass and therefore less sulfide is available for AnoxyP. The following paragraphs will show how this feedback can be described. The factors that determine the gross rate of AnoxyP are derived using the model of the S-, C- and O-cycle. Assuming that the only microbially mediated processes in a hypothetical mat are SR and sulfide-driven AnoxyP producing sulfate and using the equations for JC,SR,SO4->H2S and JC,AP,H2S->SO4 in Table 8.1 gives H H2S J C,AP,H2S-!SO4 2 J S,ext . (1 H CorgH H2S ) This shows that sulfide-removal by AnoxyP could theoretically increase infinitely if only the electron donor availability is considered and not light limitation. In reality, the substrate utilization efficiencies will, however, never approach a value of 1. In case of AnoxyP, İH2S is to a large extent influenced by diurnal dynamics and strongly depends on the temporal coupling between SR and AnoxyP. As AnoxyP cannot be performed during 281 8. DISCUSSION night, sulfide produced by SR in the dark is therefore lost to the water column. If SR would therefore exclusively depend on degrading cell material, İH2S would immediately be <0.5 (assuming 12 h light and 12 h darkness) because at least half of the sulfide produced by SR would escape AnoxyP. Such inefficient coupling was indeed observed in a meromictic salt lake where SR is instead driven by OxyP (Overmann et al., 1996). In pink mats dominated by purple sulfur bacteria (as inferred from microscopy and hyperspectral imaging) forming in the Frasassi sulfidic springs, however, we observed a very strong temporal coupling between SR and AnoxyP (Fig. 8.2). This coupling seems to allow for the high rates of sulfide consumption by AnoxyP in the pink mats that, when converted into potential gross CO2 fixation rates, were comparable to the rates observed in OxyP driven cyanobacterial mats, despite the moderate external availability of sulfide. Overall, the competitiveness of processes (here most prominent in İH2S) is not only dependent on obvious characteristics such as the substrate affinity of the microorganisms performing this process but also on the spatial and temporal coupling to other processes. -0.50 dark 67 μmol photons m-2 s-1 -0.25 239 μmol photons m-2 s-1 0.00 0.25 0.50 Depth (mm)Depth 0.75 1.00 1.25 0 200 400 600 800 1000 S (μM) tot Figure 8.2: Photograph of cyanobacterial, SOB-dominated and pink mats forming along the Frasassi sulfidic spring water flow and depth profiles of Stot concentration in a pink mat. Stot concentration was calculated from in-situ depth profiles of pH and H2S concentration measured in-situ during night, in the early morning (irradiance: 67 μmol photons m-2 s-1) and in the afternoon (irradiance: 239 μmol photons m-2 s-1). 8.1.4.3. Colonization of previously anaerobic microbial mats by cyanobacteria As discussed above, the coupling efficiency between AnoxyP and remineralisation processes is a crucial factor determining both photosynthetic and remineralisation rates. This concept is expected to also be important in the competition among phototrophs. 282 III. Perspective Namely, a positive feedback effect on AnoxyP is expected when additional Corg is provided, e.g. by OxyP. If, however, photosynthetically produced O2 and Corg are immediately consumed by aerobic respiration (AR), i.e., if OxyP and AR are well coupled (İAR=1), there is no additional export production and the rates of SR and AnoxyP are not enhanced. A negative feedback is expected when the coupling between AnoxyP and SR is disturbed. In the following, invasions of AnoxyP-driven microbial mats are simulated to show that the key to success of versatile cyanobacteria in the competition with obligate anoxygenic phototrophs might indeed be such disturbance of the coupling between AnoxyP and remineralisation. In Figure 8.3 simulations of invasion scenarios based on the model of the S-, C- and O-cycle described in 8.1.4.1 are shown. It was assumed that an AnoxyP-driven mat is first invaded by obligate oxygenic phototrophs (phase 1) and subsequently by versatile cyanobacteria (phase 2) to understand the conditions under which versatility is an advantage. An advantage is here defined by the decrease of CO2 fixation rates by other phototrophs because such decrease creates “space” for an increase in cyanobacterial CO2 fixation. As expected, simulations of the invasion by obligate oxygenic phototrophs (phase 1) show that SR generally increases. This increased availability of electron donor also results in increased rates of AnoxyP by obligate anoxygenic phototrophs. Also, the O2 production stimulates SOX processes that can provide S0, which in turn can stimulate S0- driven AnoxyP. A more detailed description of some examples of such simulations is provided in Figure 8.3. For the simulations of an invasion of mats inhabited by obligate oxygenic and anoxygenic phototrophs by versatile cyanobacteria (phase 2) it was assumed that total Corg production by cyanobacterial photosynthesis is limited upon invasion. Thus, Corg production by OxyP was displaced by Corg production by cyanobacterial AnoxyP. In contrast the total CO2 assimilation, i.e., the sum of Corg production of all autotrophic processes, was not restricted. An increase in the rates of AnoxyP by obligate anoxygenic phototrophs in phase 2 can be interpreted as an increase in competitiveness. In contrast, a decrease in CO2 fixation rate by obligate anoxygenic phototrophs would allow for an increase in cyanobacterial activity and therefore represents an advantage, i.e., an increase in fitness, of the cyanobacteria. 283 8. DISCUSSION phase 1 phase 2 16 -2 -1 ) A J (mmol CO m d ) C,tot 2 -1 14 d 2 12 -2 -1 - J (mmol O m d ) O2,OP 2 10 8 Sulfide fluxes ( mmol S m-2 d-1) 6 tot J +J 4 S,SOX.H2S SO4 S,SOX.H2S S0 JS,SR.SO4 H2S ux (mmol m fl 2 JS,AP.H2S SO4 0 J S,CYA 16 ) H =0 0 0 -2 -1 B AP,S0 S fluxes ( mmol S m d ) -1 140 H =1 O2 d JS0,SOX.S0 SO4 2 - 12 J 10 S0,SR.S0 H2S J 8 S0,AP.S0 SO4 6 4 ux ux (mmol m fl 2 16 C J = 1mmol m-2 d-1 14 O2,ext H =0 AP,S0 -2 -1 12 y=y =y =0.7 10 SO4 S0 8 6 4 ux ux (mmol m d ) fl 2 0 16 H =0.65 ) D S0 -1 14 d 12 -2 10 8 6 4 ux ux (mmol m fl 2 0 16 H =0.8 E S0 14 H =0.5 SOX,S0 12 -2 -1 10 8 6 4 ux ux (mmol m d ) fl 2 0 0 0.2 0.4 0.6 0.8 1.0 time 284 III. Perspective Figure 8.3 (previous page): Simulations of the invasion of AnoxyP-driven microbial mats by obligate oxygenic phototrophs (phase 1) and versatile cyanobacteria (phase 2) using the model of the C-, S- and O-cycle described in Figure 8.1 and Table 8.1. Unless stated differently in the respective simulations A – F, the input parameters used were as follows: JO2,ext=0 -2 -1 -2 -1 mmol m d , JS,ext=1 mmol m d , İAR=0, İO2=0.5, İS0=1, İSOX,S0=0, İSOX=0.5, İH2S=0.7, İAP=1, İAP,S0=0.8, İCorg=0.8 and y=ySO4=yS0=1. In phase 1, JO2,OP was increased until total Corg production reached 10 mmol m-2 d-1, which was assumed to be the maximal rate supported by a limiting factor. In phase 2, this limitation was not considered and JO2,OP was decreased until it reached zero or until it could not be replaced anymore by cyanobacterial AnoxyP. İcya was set to 0 in phase 1 and was then increased in phase 2 until OxyP was completely replaced by cyanobacterial AnoxyP or until the sulfide flux became limiting. The rates at time point 0 represent the rates in a AnoxyP- driven mat in steady state before the invasion calculated based on the initial values for the input parameters. A: In phase 1, the increasing rate of O2 production by OxyP stimulates SOX processes including incomplete sulfide oxidation to S0, which serves as an electron donor for AnoxyP (JS,AP,S0->SO4) and as an electron acceptor for anaerobic respiration (JS,SR,S0->H2S). Also, due to increasing OxyP additional Corg becomes available for the anaerobic respiratory processes leading to an increase in JS,SR,S0->H2S and JS,SR,SO4->H2S. The increased availability of sulfide stimulates sulfide-driven AnoxyP (JS,AP,H2S->SO4). In phase 2, rates of SR continue to increase because more of the photosythetically produced Corg is directly available for anaerobic respiration. Furthermore, there is a steep increase in S0-driven AnoxyP by obligate anoxygenic phototrophs, which is the result of increased sulfide production rates and decreased activity of the O2- dependent SOX processes. Sulfide-driven AnoxyP by obligate anoxygenic phototrophs decreases, however, because sulfide is consumed by cyanobacterial AnoxyP. B: In this simulation İO2 was chosen to be equal to 1, which means that there is no net O2 production. Thus, all photosynthetically produced O2 is available for mat-intrinsic processes, which leads to steeply increasing rates of aerobic sulfide oxidation (JS,SOX,H2S->SO4+ JS,SOX,H2S->S0). As İAP,S0 is equal to 0, 0 0 0 all S produced by aerobic sulfide oxidation to S is available for S reduction. The Corg required for this process is directly available from OxyP, as İAR is equal to 0. Rates of sulfide-driven AnoxyP decrease with increasing OxyP because there is sufficient O2 available for SOX to consume the majority of the available sulfide. In phase 2, S0 reduction continues to increase because of the additional S0 supply from cyanobacterial AnoxyP. The rate of AnoxyP by obligate anoxygenic phototrophs increases in phase 2 because the O2 supply for SOX decreases with decreasing OxyP. In a system where aerobic sulfide oxidation plays a major role, photosynthetic versatility is therefore clearly not of advantage. C: The trends observed in phase 1 of this simulation are very similar to the general trends in (B). The rates of AnoxyP, however, increase with increasing OxyP because there is less O2 available for SOX to outcompete the photosynthetic consumption of sulfide. In contrast to the simulation A–B, the overall rates of CO2 fixation decrease in phase 2. In such system, photosynthetic versatility would therefore likely be of advantage, as there is potential for an increase in cyanobacterial activity created. The decrease in CO2 fixation is the result of the decreased rates of sulfate reduction. These rates decrease because 0 most of the Corg is utilized by S reduction and because SOX processes (that do not use the sulfide exclusively for Corg production) still play a role. Most importantly, the decrease in overall CO2 fixation is, however, only possible in this scenario because S0-driven AnoxyP is excluded and cannot contribute to Corg generation. It is, however, very unlikely that the vast majority of the 0 available S is used for respiratory processes. D: For this simulation İS0 was chosen to be 0.65, which means that 45% of the produced S0 escape further reduction or oxidation by the processes considered in the model. This does not have substantial consequences in phase 1 when compared to (A). In phase 2, however, sulfide production rates, which is caused by an imbalance between anoxygenic photosynthetic Corg production and Corg consumption during anaerobic respiration (see text in section 8.1.4.3.). Consequently, also sulfide-driven AnoxyP by obligate anoxygenic phototrophs decreases rapidly, wherein it should to be noted that cyanobacterial AnoxyP is considered more competitive for sulfide in all simulations. E: As opposed to the other simulations, here also aerobic S0 oxidation is considered. Due to a similar nexus of reasons as in (E), this leads to a decrease in sulfate reduction rates and sulfide-driven AnoxyP by obligate anoxygenic phototrophs in phase 2 (see text in section 8.1.4.3.). 285 8. DISCUSSION The simulations showed that a shift from OxyP to AnoxyP in phase 2 is of advantage for the cyanobacteria if a considerable fraction of the S0 produced by versatile cyanobacteria escapes further photosynthetic oxidation, i.e., when İS0 or İSOX,S0 are high and/or if İAP,S0 is low. The simulations under these assumptions lead to a decrease in sulfide production, which has a negative feedback effect on sulfide-driven AnoxyP performed by obligate anoxygenic phototrophs (Fig. 8.3.C–E). As a result, total CO2 fixation decreases, which implies that total productivity is not limited anymore. Therefore, the cyanobacteria could increase their oxygenic photosynthetic activity and thus increase their overall CO2 fixation rates. This means that the photosynthetic versatility of the cyanobacteria is of advantage as it helps the cyanobacteria outcompete obligate anoxygenic phototrophs by capping their electron donor supply. The decrease in sulfide production is a result of the production of S0 in the 0 presence of O2. The production of S according to 0 H2S + 0.5 CO2 Æ S + 0.5 CH2O + 0.5 H2O (1) would not have a negative effect on sulfide production rates if it was further oxidized by phototrophs because this would overall yield the same Corg that was needed to produce sulfide during SR. The cyanobacterial activity can, however, stimulate aerobic S0 0 oxidation. Even if these S oxidizers were autotrophs, they do not produce Corg 0 0 stoichiometrically from S oxidation, as they oxidize a major fraction of S with O2 for energy generation (Chapter 3). As the S0 is not available for S0 reducers, SR remineralizes the photosynthetically produced Corg (see Eq.1) according to 2- + SO4 + 2 CH2O + 2 H Æ H2S + 2 CO2 + 2 H2O (2) Therefore, there is a stoichiometrical imbalance between sulfide and Corg production and consumption, and a negative feedback arises. H2S is not stoichiometrically 2- oxidized with CO2 to SO4 . Instead, a fraction of the sulfur pool is lost/buried or oxidized with O2 instead of CO2. Thus, there is substantially less Corg produced by AnoxyP than would be needed to sustain the sulfide production rates by SR that were present before the invasion by versatile cyanobacteria. The cyanobacteria thus take advantage of modulating the S-cycle and decreasing the coupling efficiency between AnoxyP and anaerobic remineralization. It should be noted that a shortcoming of using the model for such simulations is that the model does not account for likely changes in the substrate utilization efficiencies dependent on the magnitude of the available substrate flux. Namely, it would be expected that rates of substrates consumption do not linearly depend on substrate flux but are rather 286 III. Perspective defined by the affinities towards the substrate. Therefore, also the relative contribution of the functional groups to substrate consumption, as defined by the substrate utilization efficiencies, will change with substrate flux. Corresponding adjustments of the model will, however, not invalidate the general conclusions derived here. The hypothesis that S0 (or any incompletely reduced sulfur) production is the key to the success of photosynthetically versatile cyanobacteria, is supported by the fact that we indeed observed that sulfide production and AnoxyP are not well coupled in the cyanobacterial mats in the Frasassi sulfidic springs. Instead, sulfide production was almost entirely dependent on OxyP. After inhibition of OxyP with DCMU, sulfide production immediately started to decrease, even though AnoxyP continued at rates comparable to the rates observed in the pink mats (Fig.8.4). In this specific experiment the effect was likely amplified because the produced S0 was probably not used at all in the absence of O2 after abruptly inhibiting OxyP. Also, the experiment highlights another possible advantage of versatile cyanobacteria. During OxyP cyanobacteria excreted freshly assimilated DOC (data not shown). This was not observed during cyanobacterial AnoxyP. This indicates that the cyanobacteria disrupt the temporal coupling between AnoxyP and anaerobic remineralisation (decrease İH2S), which likely exerts a negative feedback effect on the activity of both functional groups (section 8.1.4.2). Overall, the results obtained from the model of the S-, C- and O-cycle suggest that environmental parameters, such as the water column sulfide concentration, do not allow prediction of whether photosynthetic versatility is of advantage in a microbial mat. Instead, the advantage of versatility is linked to the processes in the mat that compete for the incompletely oxidized sulfur product of the cyanobacteria. Namely, versatility seems to be of advantage when photosynthetically produced S0 is not further oxidized by photosynthetic processes. The environmentally most relevant alternative S0 sink is likely 0 aerobic S oxidation by SOB co-inhabiting the microbial mat. As these SOB require O2, this advantage of photosynthetic versatility can therefore only be exploited when AnoxyP and OxyP are performed simultaneously. 287 8. DISCUSSION + DCMU + DCMU + 6 h S (μM) S (μM) Stot (μM) tot tot 0 100 200 300 400 500 0 100 200 300 400 500 0100200300400500 -1 -1 -1 0 0 0 1 1 1 2 2 2 Depth (mm) Depth (mm) 3 3 Depth (mm) 3 4 4 4 0100200300400 0 100 200 300 400 0 100 200 300 400 O (μM) O (μM) 2 2 O2 (μM) AnoxyP (μM s-1) AnoxyP (μM s-1) AnoxyP (μM s-1) -2 0 2 4 -2024 -2024 -1 -1 -1 0 0 0 1 1 1 2 2 2 Depth (mm) Depth (mm) Depth (mm) 3 3 3 4 4 4 -2 0 2 4 OxyP (μM s-1) Figure 8.4: Depth profiles of Stot and O2 concentration (upper panel) and volumetric gross rates of O2 production and Stot consumption by OxyP and AnoxyP, respectively, (lower panel) in the absence and presence of DCMU, an inhibitor of OxyP. Depth profiles and rates were determined in a freshly collected cyanobacterial mat (C/B-type; see Chapter 2) placed in a -2 -1 flow through chamber. Incident irradiance was 315 μmol photons m s . Stot concentration and photosynthetic consumption was calculated from depth profiles and the light-induced dynamics of the pH and the H2S concentration, respectively (Chapter 2, 5 and 7). After the addition of DCMU, OxyP was inhibited and only AnoxyP was performed in the cyanobacterial layer. The right panel shows examples for depth profiles and rates determined after 6 h of incubation in the presence of DCMU. Rates of AnoxyP and sulfide production (as seen in the shape of the depth profile at depth below photosynthetic activity), had decreased substantially. After 1 day the mat had started to disintegrate (data not shown). 288 III. Perspective 8.2. Specific adaptations strategies among cyanobacteria to sulfidic conditions The aim of the previous section was the identification of general advantages of photosynthetic versatility. The spectrum of specific adaptations to sulfidic conditions found among cyanobacterial species is, however, multifaceted. This section attempts to enlighten why this diversity has emerged. Cohen et al. (1986) already pointed out that there is probably continuity between the four categories of adaptations of cyanobacteria to sulfide (section 1.3). Besides (i) sulfide-intolerance, these types of adaptations are (ii) sulfide-resistance of OxyP, (iii) the capability to perform simultaneous AnoxyP and OxyP without inhibition of OxyP at high H2S, and (iv) the capability for AnoxyP and OxyP; however, not necessarily simultaneous operation of both modes due to inhibition of OxyP at high H2S concentrations. The results of this thesis and other studies on sulfide-adapted cyanobacteria (Cohen et al., 1986; Garcia-Pichel and Castenholz, 1990) indeed revealed the complexity of the repertoire of responses to sulfide, which blurs the boundaries between the adaptation types and shows that a four-type categorization does not have sufficient capacity for the complete spectrum of adaptations. Most prominently, the traditional classification scheme does not consider the complex interplay between the effects of light and H2S concentration on photosynthetic activity, and ignores the time scale at which the different physiological responses take effect. These factors and the reasons why it is crucial to consider them when aiming to understand adaptation strategies in the environmental context will be discussed in the following. 8.2.1. Effects of H2S on cyanobacterial photosynthesis – an overview The effects of H2S on photosynthesis are highly diverse and complex, both in cyanobacteria as well as plants (Chapter 4–7; Wang (2012)), and vary greatly among species and even among strains (Garcia-Pichel and Castenholz, 1990). The various responses can be a result of direct interaction of H2S with enzymes or due to regulatory functions of H2S on the expression level. As oxygenic and H2S-driven anoxygenic photosynthetic electron transport intersect in the PQ pool (Fig. 8.5; Chapter 5–7), H2S concentration can also have a series of indirect effects on OxyP. The most important sites for direct interactions of H2S with the photosynthetic light and dark reactions are summarized in Figure 8.5. 289 8. DISCUSSION Ͳ1.5 P700* A0 Ͳ1.0 A1 FeͲSx FeͲSA P680* FeͲSB Fd Ͳ0.5 Pheo FNR NADP H2S QA [V] SQR 0.0 QB CO2 m E PQ hQ cytb6f PC 0.5 P700 NADPH hQ H2OOEC 1.0 P680 G3P 1.5 Figure 8.5: Sites of interaction of H2S with the photosynthetic electron transport chain and CO2 fixation pathway, the Calvin cycle, in cyanobacteria. Interaction with sites marked in green and red lead to an enhancement or decrease of photosynthetic rates, respectively. The dashed green line indicates excitation energy transfer between the two photosystems. Direct inhibition of OxyP The most striking direct negative effect of H2S on oxygenic photosynthesis is inhibition of the initial water-oxidation reaction in the oxygen evolving complex (OEC) of PSII (Chapter 4; Fig. 8.5). However, cyanobacteria inhabiting sulfidic environments show an increased resistance against inhibition and are thus able to perpetuate OxyP in the presence of sulfide, wherein the H2S thresholds causing inhibition are highly variable among species (Miller and Bebout, 2004). Additionally, the reversibility of the inhibition is variable. Namely, in some species the inhibition is irreversible (Cohen et al., 1986), while OxyP in other cyanobacteria instantaneously recovers upon removal of H2S (Chapter 4). Also, there can be a delay between depletion of H2S and recovery of OxyP (Chapter 5). Interestingly, some cyanobacteria seem to synthesize a more resistant form of PSII when they are exposed for several hours to sulfide and light (Garcia-Pichel and Castenholz, 1990). Indirect inhibition of OxyP Besides the direct inhibition of PSII, H2S can also indirectly suppress rates of OxyP because it is used for anoxygenic photosynthetic electron transport (Chaper 6–7). Oxygenic and anoxygenic photosynthetic electron transport intersect in the plastoquinone (PQ) pool and therefore electrons derived from H2O and H2S “compete” for this pool. The apparent affinities of the immediate electron donors for oxidized PQ, QB in OxyP and 290 III. Perspective SQR in AnoxyP (Figure 8.5), determine the relative contribution of both photosynthetic processes to the overall electron transport rate, which in turn is determined by the light energy harvested in PSI. The irradiance will thus also determine the threshold H2S concentration, below which simultaneous OxyP and AnoxyP is possible (Chapter 6). Overall, the rates of OxyP can be very low or even completely absent in the presence of sulfide, even though PSII is not inhibited. This is, however, only the case when SQR is active. Some cyanobacteria can immediately perform AnoxyP upon exposure to sulfide, even if they were grown under non-sulfidic conditions (Garcia-Pichel and Castenholz, 1990) (Chapter 5). Other species require an induction time of up to 4 h upon exposure to light and H2S for the de novo synthesis of SQR (Garcia-Pichel and Castenholz, 1990). Also, the time frame, over which SQR is kept in the cells seems to be variable among species (Chapter 6). Enhanced light energy conversion efficiency In some cyanobacteria, H2S can enhance the rates of OxyP when the concentrations are below a species-specific threshold (Cohen et al., 1986). In Chapter 4 and 6 we hypothesize that this enhancement might be caused by two main factors: (a) excitation energy transfer from PSI to PSII, which is indirectly triggered by the reduction of the plastoquinone pool due to anoxygenic photosynthetic electron transport (Chapter 6) and (b) a change in the absorbance cross sections of PSII and PSI, which directly depends on the H2S concentration (Chapter 4). The impact of both of these factors is most pronounced at low light intensities before light saturation effects occur. Also, these responses take effect immediately dependent on the redox state of the PQ pool and the momentary H2S concentration. A similar optimization of light energy utilization has been observed during AnoxyP (Chapter 5 and 6). Namely, due to state transition, i.e., excitation energy transfer from PSII to PSI, rates of anoxygenic electron transport exceeded the rates of oxygenic photosynthetic electron transport. This implies that light energy harvested by the phycobilisomes, which is normally transferred to PSII, is not entirely “wasted” when AnoxyP (and thus only PSI) is active. In addition to these immediate responses to H2S, it has been suggested that long term exposure to H2S alters the photosystem stoichiometry, i.e., the ratio between PSIIs and PSIs in the thylakoid membrane (Garcia-Pichel and Castenholz, 1990). An increase of the number of PSIs during AnoxyP leads to a more efficient utilization of light energy 291 8. DISCUSSION per photosystem, and thus per cell when assuming that the sum of photosystems remains constant. Enhanced maximal CO2 fixation rates In plants H2S was shown to increase the maximal rates of OxyP at saturating irradiances. This effect can mainly be assigned to an increased expression level of RuBisCo und thus to enhanced rates of CO2 fixation, which otherwise become rate limiting at high light intensities (Chen et al., 2011). Such an effect on the maximal rates of oxygenic photosynthetic electron transport in cyanobacteria has not yet been shown clearly. Still, in Chapter 5, we found indications of this effect of H2S when the cyanobacterium performed AnoxyP. Enhanced assimilation of inorganic nitrogen - In Chapter 6 we show that some cyanobacteria preferentially couple AnoxyP to NO3 , instead of CO2, assimilation. The regulation of this transition in electron acceptor - utilization remains to be enlightened. In Chapter 6, we hypothesized that NO3 assimilation is stimulated by a possible shift in the cellular NADPH:ATP ratio during AnoxyP. In some other cyanobacteria N2 fixation rates are enhanced during AnoxyP (Padan and Cohen (1982); Villbrandt and Stal (1996)) 8.2.2. Specific adaptation strategies and their advantages in the environment Studies on microbial mat ecosystems have shown that, aside from temperature, the primary variables influencing the distribution of obligate anoxygenic phototrophs vs oxygenic phototrophs are light availability and sulfide concentration (Boyd et al., 2012). This is not surprising when considering the multiplicity of effects of sulfide on photosynthesis and the interrelation to light. Unfortunately, the data on the distribution of the different responses to sulfide (8.2.1.) among cyanobacteria is very limited, which hampers our ability to make sophisticated correlations between certain traits or combinations of traits to environmental parameters. Also, the assessment of these potential controlling environmental factors (such as light and H2S) is experimentally difficult because their local availability inside microbial mats is highly dynamic over a diurnal cycle (Chapter 2). Moreover, these fluctuations would also have to be related to the temporal scale at which the cyanobacterial responses become effective. In the 292 III. Perspective following it is attempted to formulate hypotheses on the advantages of the different responses using specific examples from this study. 8.2.2.1. Adaptations of obligate oxygenic photosynthetic cyanobacteria to H2S In section 8.1 it is shown that versatility is only advantageous if phototrophic S0 oxidation is not a dominant sink of the S0 produced by the versatile cyanobacteria. Therefore, obligate oxygenic phototrophs might dominate over versatile cyanobacteria in some sulfidic environments. Fluctuating sulfide concentrations are characteristic of microbial mats in general. Despite lacking the ability to use H2S as an electron donor for photosynthesis, obligate oxygenic photosynthetic cyanobacteria are exposed to these fluctuations and have evolved various responses. Most importantly, all of them likely possess a sulfide-resistant PSII that is continuously present and that does not have to be exchanged for a more resistant form upon sunrise. Here it is also important to consider that OxyP of some cyanobacteria, such as Planktothrix str. FS34, is only temporarily resistant to H2S and that the duration of resistance is determined by the interplay between the irradiance and H2S concentration (Chapter 4). In a given environment, it is likely that only cyanobacteria that can continuously maintain their activity over the range of local light and H2S fluctuations that they are regularly exposed to will be most successful. H2S can also enhance the rates of OxyP (8.2.1), wherein it has to be differentiated between two distinct responses: (i) an increase of OxyP, which is most pronounced when the organism is exposed to non-saturating irradiances and therefore changes the initial slope of the photosynthesis-over-irradiance (PI) curve and (ii) an increase of the maximum rate of OxyP (Pmax on a PI curve) when the organism is exposed to irradiances, at which saturation or even light inhibition would occur in the absence of H2S. While (i) is an immediate and flexible adjustment, (ii) represents a long-lasting alteration. The remaining question is what the advantages of these specific sulfide-dependent regulations of the photosynthetic rate are. In cyanobacteria that regulate their activity dependent on H2S concentration the shape of the PI curve can differ considerably from the traditional shape (blue line in Fig. 8.6). This substantially more flexible response to irradiance should lead to increase in fitness of the cyanobacteria, because the available energy can be exploited more efficiently. To illustrate this Figure 8.6 shows examples for PI curves of oxygenic phototrophs that hypothetically compete for a habitat that is exposed to an irradiance of up to 170 μmol photons m-2 s-1. The area under the PI curves from zero to 170 μmol 293 8. DISCUSSION photons m-2 s-1 is representative of total photosynthesis that can be performed between sunrise and zenith assuming that irradiance increases linearly over time. In the absence of H2S the first effects of photoinhibition would reduce photosynthesis during the zenith (solid black line). In the presence of H2S, a sulfide-adapted cyanobacterium, such as Planktothrix str. FS34 (Chapter 4), yields a high integral. Similar total photosynthetic rates could alternatively only be reached by an adjustment of the optimal irradiance (Imax) towards lower irradiances (dotted black line in Fig. 8.6), which would, however, possibly cause severe photoinhibition during zenith. Also, the maximal photosynthetic rate (Pmax) could be adjusted (dashed black line in Fig. 8.6), e.g., by increasing the cellular photosystem and/or pigment content, which would, however, not be an economic solution. 140 200 120 100 150 80 100 S (μM) 2 60 H OxyP (%) OxyP 40 50 20 0 0 0 20 40 60 80 100 120 140 160 Irradiance (μmol photons m-2 s-1) Figure 8.6: Examples for PI-curves of differently adapted oxygenic phototrophs. Black lines correspond to the PI-curves of cyanobacteria, the activity of which is not affected by H2S. A PI curve similar to the one represented by the solid line would be expected in the environment. Namely, Pmax is just reached upon zenith (here: 170 μmol photons m-2 s-1). Total photosynthesis (integral of the PI curve) can, for instance, be increased by a steeper slope and lowered Imax (dotted line) or increased Imax and Pmax (dashed line). The blue line represents the PI curve of a Planktothrix-like cyanobacterium that modulates its rate of OxyP dependent on H2S. The solid line represents the PI curve OxyP is expressed in percent of Pmax of the Planktothrix-like organism. When the H2S concentrations in the photic zone of a microbial mat are low and rapidly decrease in the morning, an adjustment of the PI curve described by the blue line in Figure 8.6 would likely not be of advantage, as an alteration of the PI curve over a 294 III. Perspective small light interval would not lead to a significant increase of the total photosynthetic rate. In such mats another type of H2S-dependent modulation of OxyP might be advantageous. Namely, in microbial mats that are not exposed to sulfide, but have an internal sulfide source, the H2S-dependent adjustment of Pmax might be of selective advantage, especially when it is exposed to infrequently changing average irradiances. In a microbial mat that is exposed to relatively low irradiances, e.g., because days are normally cloudy, phototrophs will exhibit a PI curve that is adjusted according to the available light range. On rare sunny days, the cyanobacteria will migrate downwards when a light level is reached that exceeds the maximum irradiance of a cloudy day. A cyanobacterium that is not adapted will remain in the deeper layers to prevent photoinhibition. During migration into these deeper layers it might transiently encounter H2S. Some specialized cyanobacteria will respond to this H2S with an increase in the maximal CO2 fixation rate and thereby increase Pmax. The phototactic or photophobic cyanobacteria are thereby able to migrate upwards again (Tamulonis et al., 2011). Such adjustments normally only start playing a role when shifts in the irradiance are long- lasting. Thus, a spontaneous increase Pmax dependent on H2S concentration likely increases the fitness of sulfide-adapted cyanobacteria in environments exposed to fluctuating average irradiances. 8.2.2.2. Adaptations of versatile photosynthetic cyanobacteria to H2S The most unintuitive type of adaptation strategy listed by Cohen et al is type 4 (section 1.3). Type 4 cyanobacteria are able to perform both OxyP and AnoxyP. PSII is, however, partially inhibited by H2S, which leads to a “dip” in the overall photosynthetic CO2 fixation rates at intermediate H2S concentrations. This type classification was based on measurements at one light intensity where only the H2S concentration was varied. In the environment, however, both light and H2S concomitantly fluctuate. It is therefore very likely that in the natural environment of these cyanobacteria, such a “dip” would not manifest. This is because the transition between photosynthetic modes is influenced by various direct and indirect mechanisms (section 8.2.1). First, as pointed out above, the inhibitory effects might not always become manifest due to the temporal dynamics of the inhibition (Chapter 4). Second, some cyanobacteria synthesize a resistant PSII upon exposure to sulfide and light (Garcia-Pichel and Castenholz, 1990). Given that the timing of synthesis and exposure to sulfide and light in the environment are well tuned, the sulfide-sensitive PSII might never be inhibited. Namely, sulfide concentrations in mats in 295 8. DISCUSSION the morning are generally relatively high. A well-adapted cyanobacterium should have synthesized a sulfide-resistant PSII before the sulfide levels decrease below the thresholds that allow for OxyP. Third, the “dip” might still not occur in the environment, even if the inhibition of PSII is not delayed after exposure to sulfide, if the cyanobacteria do not synthesize a sulfide-resistant PSII and if the inhibition is instead strictly dependent on H2S concentration. This is because the transition between AnoxyP and OxyP is regulated based on the complex interplay between H2S and light availability. If, for instance, the light intensity was still low enough when the H2S concentrations cause inhibition, rates of OxyP might not necessarily be lower than rates in the absence of H2S. An exception to this concept is, however, the cyanobacterium isolated from Little Salt Spring (LSS) (Chapter 5). In this cyanobacterium, AnoxyP comes with a set of disadvantages. Namely, AnoxyP and OxyP were never performed simultaneously. Thus, there was always a phase of photosynthetic inactivity after depletion of H2S. Intriguingly, the LSS isolate is still highly adapted to sulfidic conditions as it instantaneously starts to perform AnoxyP, i.e., AnoxyP does not require an induction time. Possibly, this cyanobacterium would be successful in environments where it is exposed to infrequent pulses of sulfide. Clearly, it is not successful in the LSS mats that are dominated by a different species (Chapter 5). Also, it remains to be understood what kind of metabolism the LSS isolate performs in the phase of photosynthetic inactivity. In section 8.1 it is discussed that S0 production during AnoxyP acting jointly with aerobic S0 oxidation is possibly the key to success of versatile cyanobacteria. Namely, when these processes cooperate, a net decrease in sulfide production can be triggered, which has a negative feedback effect on the activity of obligate anoxygenic phototrophs. 0 To achieve this feedback effect, S has to be produced in the presence of O2. Therefore, this advantage of versatility can only be exploited by simultaneous AnoxyP and OxyP. Consequently, the transition between OxyP and AnoxyP of the most successful versatile cyanobacterium in a given environment is expected to be regulated so that both photosynthetic modes operate simultaneously for most of the light phase. The main regulatory parameter of the transition is likely the affinity of SQR to H2S and the plastoquinone (PQ) pool in relation to the affinity of PSII components to this PQ pool (Chapter 6). The most successful cyanobacterium will therefore be equipped with an SQR that is highly tuned to the diurnal dynamics of local H2S concentration and irradiance, which is in agreement with our observations in the Frasassi sulfidic springs (Chapter 2 and 7). 296 III. Perspective 8.3 Implications 8.3.1. Stromatolites and oxygenic photosynthesis The value of lithified stromatolite and MISS (microbially induced sedimentary structures) in the geological records as a proof for the onset of OxyP is heavily debated (e.g., Grotzinger & Knoll, 1999). There is doubt that the lithified structures are of biological origin, while the question of whether they provide evidence of OxyP is only rarely raised. Also, extensive deposits of organic-rich shales are considered an indicator of early OxyP (Lyons et al., 2014). An intriguing question is if ancient microbial mats might have been driven by forms of AnoxyP and if the organic-rich deposits might have arisen from active AnoxyP and not OxyP. Our data obtained in AnoxyP-driven mats in the Frasassi sulfidic springs and the results of the model introduced in section 8.1 suggest that AnoxyP can indeed be maintained despite high rates of organic matter burial, as long an external source of electron donor is available to initiate an efficient coupling of organic matter generation and degradation. As external electron donors for photosynthesis were likely abundant in the Archean (section 1.1.), lithified stromatolites and shales could have been formed by AnoxyP-driven systems. Our studies of the physiology of cyanobacterial isolates revealed a wide spectrum of adaptations to sulfide (Chapter 4– 7), which leads to their dominance in contemporary microbial mats that represent analogues to their ancient counterparts (Chapter 2 and 5). This is because photosynthetic versatility represents a selective advantage for the cyanobacteria over other types of photosynthesizers that have a limited repertoire of photosynthetic capabilities. Exploiting this advantage has, however, consequences for the net O2 budget of microbial mats. Mats dominated by either obligate anoxygenic phototrophs, obligate oxygenic phototrophs or versatile cyanobacteria will have different net O2 budgets. Transitions between the relative coverage of the shallow coastal regions by these different mat times over a geological timescale might represent an explanation for possibly biologically driven fluctuations in atmospheric O2 concentration, such as the “whiffs” of O2 in the Archean, suggested by the rock records (Anbar et al., 2007). In this hypothesis the evolutionary timing of the onset of the different photosynthetic modes is the crucial factor determining global changes in O2 concentration. Thus, such hypotheses cannot further be fostered without the identification of suitable biosignatures of cyanobacterial activity or further insights into the evolution of photosynthesis based on biochemical and molecular data. 297 8. DISCUSSION 8.3.2. The importance of sulfur oxidation intermediates in the Proterozoic C-, S- and O-cycle Proterozoic oceans were likely redox stratified, with a low-oxygen water layer above anoxic deep waters. Open ocean deep waters were likely characterized Fe2+-rich conditions, while the coastal oceans tended towards euxinia (Lyons et al., 2014). One of the most intriguing questions is: Why was there a 1-2 billion years (Ga) long delay in the rise of O2 during the Proterozoic after the GOE? (section 1.1) This study does not directly add to our understanding of the mechanisms that sustained the apparent biogeochemical stasis. However, there is rising doubt that the Proterozoic was actually indeed the “boring billion”(Holland, 2006). Namely, the extent of euxinia, as well as the atmospheric O2 concentrations, might well have fluctuated. This idea arose from the fact that the geological records seem to be contradictory dependent on the time of deposition and distance from the shore (Lyons et al., 2014). Our data and model of the S-, C- and O-cycle in microbial mats comes with a possible scenario that might explain these fluctuations and the final collapse of euxinia towards the end of the Proterozoic. Johnston et al. (2009) proposed that significant contribution to primary production by AnoxyP might have sustained euxinia by increasing the export production of organic matter and thus perpetuating high sulfate reduction rates (section 1.1; Fig. 1.2). Furthermore, the authors propose a positive feedback effect of SR on AnoxyP. As the overall primary productivity is assumed to be nutrient limited, these feedback effects would have tempered OxyP. Let us now assume that the capability to perform AnoxyP was acquired late during the evolution of cyanobacteria, or that the metabolic regulation of the transition between OxyP and AnoxyP was slowly brought to perfection during the Proterozoic. The adjacent questions are: How would the rise of photosynthetically versatile cyanobacteria have impacted the S-, C- and O-cycling in the modelled euxinic oceans suggested by Johnston et al.? Would it have mattered if AnoxyP was performed by obligate anoxygenic phototrophs or by versatile cyanobacteria? As proposed in 8.1.4., the fate of S0, or any incompletely oxidized sulfur product, is the crucial factor that determines the effect of cyanobacterial AnoxyP on the S-, C- and O-cycle. Namely, if the intermediately reduced sulfur escapes further phototrophic oxidation, a negative feedback effect on the sulfide production rates arises. In microbial mats alternative sinks for the intermediate are plausibly mainly aerobic sulfur oxidation processes. In the water column of the Proterozoic oceans an additional sink could have 298 III. Perspective played a role: the sinking speed of S0 produced in deeper layers of the photic zone might have been sufficient to escape phototrophic oxidation. Johnston et al. already suggested that these sinking S0 particles might have enhanced pyritization rates once reaching the seafloor. Pyritization in our model represents an “external” sink of S0, which substantially enhances the negative feedback effect of versatility on sulfide production processes. Over geological time scales, shifts in the photosynthetic activity of cyanobacteria would have possibly impacted the distribution of euxinia and might have even lead to a collapse of euxinia once versatility was rendered to perfection. To date, it is, however, not clear if the redox cline of the Proterozoic oceans actually intruded into the photic zone, which is clearly the governing assumption of any model that involves AnoxyP. Also, it is not known when cyanobacteria acquired the genes needed to perform AnoxyP. If the assumptions made above are right, though, the perspective on the ecology of versatile cyanobacteria presented here comes with explanations for shifts in the global sulfide and O2 budgets through Earth’s history. 8.4. Outlook Rising attention is given to ecosystems that resemble our image of ancient environments (Voorhies et al., 2012; Sumner et al., 2015). What these analogue systems have in common is that the inhabitant cyanobacteria are well adapted to sulfidic conditions. This study has contributed to revealing the multiplicity of possible adaptations and provided further insights into the physiology of some cyanobacteria by relating photosynthetic rates to the dynamics of environmental parameters (Chapter 4, 6–7). Also, this study further supports the hypothesis that the photosynthetic versatility of cyanobacteria might have been an important process during the Earth’s history. A major link that would allow for an incontrovertible extrapolation of studies on the activity of extant cyanobacteria to ancient systems is, however, missing. Specifically, it is not known if the capability to perform AnoxyP is an ancient trait. There is a wide stretch of possible scenarios: AnoxyP might have been performed by the cyanobacterial ancestor even before it evolved the capability to perform OxyP but the enzymes required for AnoxyP might have also been acquired rather recently through horizontal gene transfer (Pham et al., 2008). This means that any models of ancient biogeochemical cycling based on conclusions drawn from the activity of extant cyanobacteria are not testable, so far. Future research on photosynthetically versatile cyanobacteria should therefore aim for the identification of 299 8. DISCUSSION signatures of cyanobacterial activity and additional molecular markers that might be suitable for “molecular clock” approaches. Among possible signatures that may have been left behind by cyanobacterial activity, isotope fractionations might be promising. Namely, significant sulfur isotope fractionation can be preserved in organic matter (Fike et al., 2009; Habicht et al., 2002). It is therefore worthwhile examining sulfur isotope fractionation as a function of cyanobacterial metabolism. Furthermore, we have shown in Chapter 6 that the C:N ratio of the cyanobacteria varies dependent on photosynthetic mode and it would be interesting to study if this is a general pattern among versatile cyanobacteria. Also, the effect of photosynthetic activity on carbonate precipitation in lithifying microbial mats might become of interest. To be able to identify possible enzymes or enzyme complexes that are of use to make deflections on the evolutionary timing of AnoxyP, our understanding of the physiology and biochemistry of versatile cyanobacteria has to be deepened. So far, sulfide:quinone:oxidoreductase (SQR) is the only enzyme identified to be directly involved in cyanobacterial AnoxyP. It is, however, likely that other enzymes are, at least indirectly, involved. For instance, the cyanobacteria need to “sense” sulfide and light to be able to induce the synthesis of SQR. A regulation based on the redox state of the PQ pool can be excluded as the universal mechanism. This is because some cyanobacteria have a sulfide-resistant PSII and can continue OxyP unaffected by the presence of sulfide, which means that also the redox state of the PQ pool is expected to not be affected. Therefore, other mechanisms of redox (or direct H2S-) sensing are likely involved. Another interesting approach is to study how cyanobacteria deal with the simultaneous presence of sulfide and O2 in the cells as this might substantially enhance the generation of reactive oxygen species (e.g., Hoffman et al., 2012). Yet another problem that needs to be solved by cyanobacteria is the transport of S0 out of the cells. This is because, in analogy to other bacteria, a localization of SQR at the luminal side of the thylakoid membrane is likely, which would complicate the disposal of S0 via the periplasm (Griesbeck et al., 2000). If right, S0 transport would therefore be expected to be a mechanism that is very specific to cyanobacteria and its study might be promising in the search for molecules that can be used as “clocks”. 300 III. Perspective 8.5. References Alberty, R. (2003). Thermodynamics of biochemical reactions. Hoboken, NJ, USA: John Wiley & Sons Inc. Anbar, A. D., Duan, Y., Lyons, T. W., Arnold, G. L., Kendall, B., Creaser, R. A., Kaufman, A. J., Gordon, G. W., Scott, C., Garvin, J., et al. (2007). A whiff of oxygen before the great oxidation event? Science 317, 1903–6. doi:10.1126/science.1140325. Boyd, E. S., Fecteau, K. M., Havig, J. R., Shock, E. L., and Peters, J. W. (2012). 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Al-Najjar1, Dirk de Beer1 1Max Planck Institute for Marine Microbiology, Celsiusstr.1, 28359 Bremen, Germany Spectrally resolved imaging was applied to study the growth dynamics of phototrophic biofilms comprizing a mixture of one cyanobacterial and one diatom species. Linear spectral unmixing was combined with liquid chromatography to quantitatively discriminate the areal biomass densities of the two populations. The grown biofilms exhibited highly heterogeneous distribution with patches of 1-2 mm in size, although the conditions provided for growth, including substrate roughness, illumination and flow of the overlying water, were homogeneous. The biomass was initially dominated by cyanobacteria, which exhibited an exponential-like growth phase during days 2-7. Their population declined during days 9-17, which coincided with the growth phase of the diatom population. By allowing non-invasive and real-time measurements and data evaluation, the spectral imaging approach constitutes a useful tool for microbial ecologists. Published in: First Workshop on Hyperspectral Image and Signal Processing: Evolution in Remote Sensing, WHISPERS 2009, p. 1 – 4 304 IV. Contributed work SPATIAL PATTERNS AND LINKS BETWEEN MICROBIAL COMMUNITY COMPOSITION AND FUNCTION IN CYANOBACTERIAL MATS Mohammad A. A. Al-Najjar1,2, Alban Ramette1, Michael Kühl3,4, Waleed Hamza5, Judith M. Klatt1, Lubos Polerecky1,6 1Max Planck Institute for Marine Microbiology, Celsiusstr.1, 28359 Bremen, Germany 2Marine Microbial Ecology Group, Red Sea Research Center, KAUST, Thuwal, Saudi Arabia 3Marine Biological Section, Department of Biology, University of Copenhagen, Helsingør, Denmark 4Plant Functional Biology and Climate Change Cluster, University of Technology Sydney, Sydney, NSW, Australia 5Biology Department, UAE University, Al-Ain, UAE 6Department of Earth Sciences – Geochemistry, Faculty of Geosciences, Utrecht University, Princetonplein 9, 3584 CC, Utrecht, The Netherlands We imaged reflectance and variable fluorescence in 25 cyanobacterial mats from four distant sites around the globe to assess, at different scales of resolution, spatial variabilities in the physiological parameters characterizing their photosynthetic capacity, including the absorptivity by chlorophyll a (Achl), maximum quantum yield of photosynthesis (Ymax), and light acclimation irradiance (Ik). Generally, these parameters significantly varied within individual mats on a sub-millimeter scale, with about 2-fold higher variability in the vertical than in the horizontal direction. The average vertical profiles of Ymax and Ik decreased with depth in the mat, while Achl exhibited a sub-surface maximum. The within-mat variability was comparable to, but often larger than, the between-sites variability, whereas the within-site variabilities (i.e., between samples from the same site) were generally lowest. When compared based on averaged values of their photosynthetic parameters, mats clustered according to their site of origin. Similar clustering was found when the community composition of the mats' cyanobacterial layers were compared by automated ribosomal intergenic spacer analysis (ARISA), indicating a significant link between the microbial community composition and function. Although this link is likely the result of community adaptation to the prevailing site-specific environmental conditions, our present data is insufficient to identify the main factors determining these patterns. Nevertheless, this study demonstrates that the spatial variability in the photosynthetic capacity and light acclimation of benthic phototrophic microbial communities is at least as large on a sub-millimeter scale as it is on a global scale, and suggests that this pattern of variability scaling is similar for the microbial community composition. Published in: Frontiers in Microbiology; 5(604). DOI: 10.3389/fmicb.2014.00406 305 IV. Contributed work EFFECT OF HIGH ELECTRON DONOR SUPPLY ON DISSIMILATORY NITRATE REDUCTION PATHWAYS IN A BIOREACTOR FOR NITRATE REMOVAL Anna Behrendt1, Sheldon Tarre2, Michael Beliavski2, Michale Green2, Judith M. Klatt1, Dirk de Beer1, Peter Stief1,3 1Max Planck Institute for Marine Microbiology, Celsiusstr.1, 28359 Bremen, Germany 2Faculty of Civil and Environmental Engineering, Technion, Haifa, Israel 3University of Southern Denmark, Department of Biology, NordCEE, Odense, Denmark í The possible shift of a bioreactor for NO3 removal from predominantly denitrification (DEN) to dissimilatory nitrate reduction to ammonium (DNRA) by elevated electron í donor supply was investigated. By increasing the C/NO3 ratio in one of two initially identical reactors, the production of high sulfide concentrations was induced. The í response of the dissimilatory NO3 reduction processes to the increased availability of organic carbon and sulfide was monitored in a batch incubation system. The expected shift from a DEN- towards a DNRA-dominated bioreactor was not observed, also not under conditions where DNRA would be thermodynamically favorable. Remarkably, the í microbial community exposed to a high C/NO3 ratio and sulfide concentration did not use the most energy-gaining process. Published in: Bioresource Technology; 171C:291-297.DOI: 10.1016/j.biortech.2014.08.073 306 IV. Contributed work LOW-LIGHT ANOXYGENIC PHOTOSYNTHESIS AND Fe-S-BIOGEOCHEMISTRY IN A MICROBIAL MAT Sebastian Haas1, Jennifer L. Macalady2, Artur Fink1, Judith M. Klatt1, Volker Meyer1, Trinity L. Hamilton2, Brian Kakuk, Dirk de Beer1 1Max Planck Institute for Marine Microbiology, Celsiusstr.1, 28359 Bremen, Germany 2 Pennsylvania State University, University Park, Pennsylvania, USA We report extreme low-light anoxygenic photosynthesis on the surface of thick microbial mats from 30 m depth at the cave wall of Magical Blue Hole (MBH) situated on Abaco Island (Bahamas), which present one of the most light-limited habitats for phototrophs known to date. In situ irradiance on a sunny December day was between 0.021 and 0.084 μmol photons m-2 s-1. We directly demonstrated light-dependent carbon uptake under slightly increased irradiance (0.27 μmol photons m-2 s-1) and estimated a photoautotrophic carbon fixation rate of 14.5 nmol C cm-2 d-1. A 16S rRNA clone library of the green surface mat layer was dominated (74%) by clones affiliated with Green Sulfur Bacteria (GSB). Typical pigments of brown GSB, Bacteriochlorophyll e and (ȕ-)isorenieratene, were detected. Their absorption properties are well-adapted to harvest the most abundant part of the photosynthetically active radiation spectrum in situ (475-530 and 350-400 nm). Hydrogen sulfide (3.4-8.3 μM) is available as electron donor for the mat community from below halo-chemocline at 25 m water depth. Despite the high abundance of į- Proteobacteria (40%) in our clone library obtained from a thick orange layer underneath the phototrophic surface layer, incubations with 35S-labeled sulfate showed low sulfate reduction rates (0-99.2 nmol (cm porewater)-3 d-1). High amounts of chromium reducible 0 -3 sulfur (S + FeS2; 1.3-43.1 μmol (cm mat) ) as well as light-independent sulfide depletion suggest that the mats function as net sulfide sinks. Surprisingly abundant dithionite reactive Fe species (4.3-22.2 μmol (cm mat)-3) may be the major source of oxidizing power. Abiotic Fe-S-reactions, possibly in combination with microbially catalyzed Fe-reduction, may play the main role in pyrite formation. This first biogeochemical study of a microbial mat in a blue hole describes an energy-limited environment with low-light-adapted anoxygenic photosynthesis and intriguing links between sulfur, carbon and iron biogeochemistry. In preparation 307 Acknowledgements First, I would like to thank my supervisor Lubos for introducing me into the crazy world of photosynthesis and microsensors. Thank you for all the freedom and special thanks for the “illuminating discussions” and taking me along lines of thought off the mainstream (even without some “vicious wine”). Dirk, thanks for letting me be part of the wonderful microsensor group, and for being there in the right moments to encourage me and disentangle my nods in the neurons. Thanks to Prof. Widdel for being my reviewer and for the surpassing moral support! Thank you Nicole and Astrid who agreed to be on my committee! Thanks to the Microsensor group; I really loved working with you in the last years! Especially to our fantastic technicians - What would I have done without your patience and flexibility! Thanks also to my wonderful office mates; Peter with his calming aura that has often counterbalanced hyped-up me; Duygu and Anna who have always had an open ear for me and believed in me. Thanks for all the laughter and looniness. I am so incredibly happy that we finally managed to sit in one office! Thanks to Anja K. who always gave me the feeling that I can manage everything I want. Thank you Arjun, especially for your support during proposal writing! Thank you Mohammad. I so much enjoyed working with you and highly appreciate you as a scientist and person! Thanks to the Frasassi stinky springs team! Stefan, it was really great fun to work and hang out with you in Coldigioco! Thanks also to the Dan for his support in the field. The same goes to Jenn who has additionally been of great support once we left Coldigioco. Thank you Sandro and Paula for the incomparable atmosphere in Coldigioco! Steffi, you have been just amazing. I wish we could “relax” again in Lindwurm listening to the calming voice of the generator. Thank you Kathleen for passing by in Frasassi during your holidays! You are really a fantastic friend and scientist! Thank you Petra for being you – and my all-in-one-collaborator-babysitter-and-friend. Thank you Hannah for “holding my hand” during thesis writing, for amazing science discussions and for being a wonderful friend. Thanks to the electronic and mechanical workshops who helped me make my setup dreams come true! Thanks also to the Christiane-Nüsslein-Volhard foundation that has made my life so much easier. Special thanks to Jens – It all started with you and the Planktomycetes! 308 Danke Herr Keppler. Sie waren ein fantastischer Lehrer und ich weiß nicht, ob ich ohne Ihre Chemiestunden und Ihren Enthusiasmus jetzt da wäre wo ich bin. Es sollte mehr Lehrer wie Sie geben. Last but definitely not least, ein unglaublich großes Danke an meine Familie Danke Ingrid und Franz für eure Zeit und Unterstützung in jeder Situation. Danke Mama; dafür, dass du immer meine beste Freundin und Beraterin in jeder Lebenslage warst, dass du mich Stärke gelehrt hast und dafür, dass du immer bedingungslos an mich glaubst. Alles wird gut. Danke Reimar; dafür, dass du mein sowohl mein Freund, als auch Lebenspartner und wunderbarer Vater bist. Danke für deine Geduld und deine grenzenlose Unterstützung; im Alltag, beim Zelten an stinkigen Quellen im Regen und wenn ich verrückte Pläne schmiede, die in kein Konzept passen wollen. Danke, dass du mich einfach so akzeptierst wie ich bin. Mara und Reimar – Danke, dass ihr so seid wie ihr seid. 309 Anlage 1 zur Promotionsarbeit Versicherung an Eides Statt Ich, Judith Klatt, versichere an Eides Statt durch meine Unterschrift, dass ich die vorstehende Arbeit selbstständig und ohne fremde Hilfe angefertigt und alle Stellen, die ich wörtlich dem Sinne nach einer Veröffentlichung entnommen habe, als solche kenntlich gemacht habe, mich auch keiner anderen als der angegebenen Literatur oder sonstiger Hilfsmittel bedient habe. Ich versichere an Eides Statt, dass ich die vorgenannten Angaben nach bestem Wissen und Gewissen gemacht habe und dass die Angaben der Wahrheit ensprechen und ich nichts verschwiegen habe. Die Strafbarkeit einer falschen eidesstattlichen Versicherung ist mir bekannt, namentlich die Strafandrohung gemäß § 156 StGB bis zu drei Jahren Freiheitsstrafe oder Geldstrafe bei vorsätzlicher Begehung der Tat bzw. gemäß § 161 Abs. 1 StGB bis zu einem Jahr Freiheitsstrafe oder Geldstrafe bei fahrlässiger Begehung. ______Judith Klatt