Effects of Meiofauna and Cable Bacteria on Oxygen, Ph and Sulphide Dynamics in Baltic Sea Hypoxic Sediment

Effects of meiofauna and cable bacteria on oxygen, pH and sulphide dynamics in Baltic Sea hypoxic sediment

A sediment core experiment with focus on different abundances of meiofauna and meiofaunal bioturbation in hypoxic Baltic Sea sediments. Meiofaunal effect on redox biogeochemistry of the sediment was studied in co-existence with cable bacteria.

Johanna Hedberg

Department of Ecology, Environment and Plant sciences M.Sc. Thesis 60 ECTS credits Marine biology Master´s Programme in Marine Biology (120 ECTS credits) Spring term 2019 Supervisor: Stefano Bonaglia Effects of meiofauna and cable bacteria on oxygen, pH and sulphide dynamics in Baltic Sea hypoxic sediment

Johanna Hedberg

Abstract

Oxygen depleted areas in the Baltic Sea are wide spread and affecting sediment biogeochemistry leading to faunal migration by formation of toxic sulphide. In sediments where reoxygenation events occur, recolonization of meiofauna and cable bacteria are believed to enhance ideal conditions and facilitate recolonization of other fauna. In sediments world-wide most abundant are bioturbating meiofauna representing various phyla grouped by body size (>0.04 and <1 mm) and known to enhance bacterial activity. Meiofaunal abundance and meiofaunal bioturbation in sulfidic sediments and effects on bacterial community structure is currently poorly understood. As second thesis to find filament building sulphide oxidising cable bacteria in the Baltic Proper, their co-existence with meiofauna and effect on sediment were also studied. Alive meiofauna was added to otherwise intact cores creating gradients of abundance (CTR = unmanipulated cores, DEBRIS = debris addition, LM = low meiofauna abundance and HM = high meiofauna abundance, n = 3) and microsensor profiles of oxygen, pH and sulphide were measured weekly for three weeks to obtain effects. Cores were then sliced to confirm meiofaunal gradient and to obtain samples for cable bacteria presence and bacterial community structure. High meiofauna abundance and meiofaunal bioturbation increased oxygen penetration depths (OPDs) at week two (ANOVA: F2,6 = 6.395, p = 0.033; Tukey test: � = 0.613) and deepened sulphide apparent appearance boundaries (SAABs) at week one, though this difference was not statistically significant. Fluorescence in situ hybridisation (FISH) confirmed cable bacteria in all cores and insignificant different densities. Cable bacteria eventually deepened SAABs in CTR, DEBRIS and LM to same depth. Metabarcoding and sequencing revealed significant different microbial community structures between treatments, suggesting effects from meiofaunal abundances and meiofaunal bioturbation. The work in this thesis suggests that (1) cable bacteria and meiofauna coexist in sediments recently exposed to reoxygenation events, (2) high meiofauna abundance and meiofaunal bioturbation increase OPDs and (3) high meiofauna abundance and meiofaunal bioturbation changes bacterial community structure.

Keywords Meiofauna, Bioturbation, Cable bacteria, Sediment, Oxygen, pH, Sulphide

Sammanfattning

Syrefattiga områden i Östersjön är vitt spridda och påverkar sedimentets biogeokemi och resulterar i migration av fauna pga. toxisk sulfid. I sulfidiska sediment där återkommande syresättning förekommer, tror man att kolonisering av meiofauna och kabelbakterier förbättrar sedimentet och därmed underlättar för vidare fauna att återkolonisera. Mest förekommande i sediment jorden runt, är bioturberande meiofauna som är representerade från olika fyla, grupperade efter kroppsstorlek (>0.04 and <1 mm) och kända för att förstärka bakterieaktivitet. Förekomsten av meiofauna och dess bioturbation i sulfidiska sediment och påverkan på bakteriesamhällsstrukturen är mindre studerad. Som den andra som upptäckt filamentbildande sulfidoxiderande kabelbakterier i Egentliga Östersjön så studerades även deras samverkan med meiofauna. Levande meiofauna adderades till annars intakta sedimentproppar i ett försök att få en gradient av förekomst (CTR = ej manipulerade sedimentproppar, DEBRIS = addition av debris, LM = låg förekomst av meiofauna och HM = hög förekomst av meiofauna, n = 3) och microsensorprofiler av syre, pH och sulfid undersöktes varje vecka i tre veckor för att studera sedimentets påverkan. Sedimentpropparna skivades sedan för att konfirmera meiofauna gradienten och samla prover för förekomst av kabelbakterier och den bakteriella samhällsstrukturen. Hög förekomst av meiofauna och dess bioturbation ökade syrepenetrationen i vecka två (ANOVA: F2,6 = 6.395, p = 0.033; Tukey test: � = 0.613) och fördjupade sulfiduppkomstbarriärerna redan i vecka ett, men ej statistiskt signifikant. Fluorescerande in situ hybridisering (FISH) konfirmerade kabelbakterier i all sedimentproppar och ej signifikant skillnad mellan deras densitet. I slutet av experimentet fördjupande kabelbakterierna sulfiduppkomstbarriärerna i CTR, DEBRIS och LM till samma djup som i HM. Metabarkodning och sekvensering visade signifikant skillnad i den bakteriella samhällsstrukturen, vilket påvisar en möjlig effekt av meiofauna och dess bioturbation. Den här uppsatsen drar slutsatserna att (1) kabelbakterier och meiofauna samexisterar, (2) hög meiofauna förekomst och dess bioturbation ökar syrepenetrationen och (3) hög meiofauna förekomst och dess bioturbation påverkar den bakteriella samhällsstrukturen.

Nyckelord Meiofauna, Bioturbation, Kabelbakterier, Sediment, Syre, pH, Sulfid

Contents

INTRODUCTION ...... 1 AIM/RESEARCH QUESTIONS...... 6 METHODS ...... 7 RESULTS ...... 13 DISCUSSION ...... 20 ACKNOWLEDGEMENTS ...... 27 REFERENCES ...... 28 SUPPLEMENTARY INFORMATION...... 33

Introduction

Oxygen depleted areas in aquatic environments develops when the oxygen demand is higher than oxygen supply (Snoeijs-Leijonmalm and Andrén 2017) and caused by factors, such as eutrophication and climate change (Vahtera et al. 2007; Markus Meier, Eilola, and Almroth 2011). Eutrophication enhances primary production that causes higher oxygen consumption by the degradation of increased settled organic material (Vahtera et al. 2007). Ongoing climate change with warmer waters are projected by models to lower the solubility of oxygen in water and increase the oxygen consumption by enhancing aerobic respiration, leading to further oxygen depletion in aquatic systems (Markus Meier, Eilola, and Almroth 2011).

Figure 1. The positive feedback-loop. External load of nutrients increases the uptake by and the production of phytoplankton. Higher loads of organic material from the phytoplankton settle on the sediment and require more oxygen when aerobic bacteria degrade the settled organic material. As the aerobic degradation consumes oxygen, the oxygen concentration in the bottom water and sediment decreases and eventually affects the chemical and biological redox reactions causing benthic release of phosphorous. The bioavailable phosphorous eventually reaches the surface waters and increases the already high concentration of phosphorous that further enhances the primary production. The loop continues into a second round, and for every loop each step is enhanced (Vahtera et al. 2007). Illustration inspired by Vahtera et al. (2007) and created with the Courtesy of the Integration and Application Network, University of Maryland Center for Environmental Science (ian.umces.edu/symbols/).

In the debate about eutrophication and oxygen depleted areas the Baltic Sea is often mentioned due to the fact that the sea holds one of the largest areas of oxygen depletion. Oxygen depleted areas in the Baltic Sea are increasing (Carstensen et al. 2014), and this trend is caused by nutrient loading from anthropogenic actions (Hansson, Andersson, and Axe 2011) during the last two centuries (Noffke et al. 2016). Nutrients can reach the water mass in aquatic environments by two paths: (1) External loading where nutrients are coming from land, e.g. via run-off, precipitation or sewage outlets (Snoeijs- Leijonmalm and Andrén 2017), or (2) internal loading by benthic release due to changed sediment chemistry. Internal load of the macronutrient phosphorous (P) from oxygen depleted sediments is eight times higher than the external load (Noffke et al. 2016) and one important part in the positive feedback loop that slows down the recovery (Vahtera et al. 2007). The external loading of nutrients into the sea feeds a positive feedback-loop that increases the primary production and thus the areas of oxygen

1 depletion (Figure 1) (Vahtera et al. 2007; D.J. Conley 2012). Even though the external loading of nutrients has decreased by 40 % since the 1980s (D.J. Conley 2012), the recovery of the eutrophic conditions in the Baltic Sea is slow, and it might take up to decades (Gustafsson et al. 2012) or even a century (Savchuk and Wulff 2009) before effects of improvement are visible.

The slow recovery, of the eutrophic conditions in the Baltic Sea, is also caused by the still incoming load of nutrients together with the physical conditions of the Baltic Sea (Noffke et al. 2016). The nutrients arrive via the run-off from land inhabiting about 85 million people and the over 200 rivers (Sweitzer, Langaas, and Folke 1996; Snoeijs-Leijonmalm and Andrén 2017) in the large catchment (1.6 million km2) surrounding the semi-enclosed brackish sea (Figure 2) (Björck 1995). In the narrow outlet to the North Sea, between Denmark and Sweden, there are shallow sills preventing the deeper saline inflow of oxygenated water, which makes the renewal time of the Baltic Sea water slow (30 - 40 years). The inflow of lighter freshwater from the rivers and the inflow of heavier saline water forms a permanent halocline at ~80 m depth, separating the less saline surface water (5 - 8) from the more saline bottom water (9 - 13). The strong stratification by the halocline impedes vertical mixing of the two waterbodies resulting in different concentrations of nutrients and oxygen. The surface water has a high concentration of oxygen via diffusion from the atmosphere and primary producers lowers the nutrient concentration through assimilation. In the water below the halocline nutrient concentrations are higher because of the degradation of settled organic material from the surface water and from the internal release of nutrients from the sea floor due to oxygen depletion (Snoeijs-Leijonmalm and Andrén 2017). Whereas in oxic conditions animal bioturbation contributes with high nutrient fluxes from sediment to the overlying water due the flow-through of water in the burrows and enhancement of degradation (Reise 2002). The appearance of water column stratification sustains the oxygen depletion in the Baltic Sea sediments (Diaz 2001).

Figure 2. The Baltic Sea with surrounding countries, connected lakes and rivers within the catchment (grey). Darker grey indicates higher salinity (Elmgren and Larsson 2001).

Conditions of oxygen depletion in sediments are often separated into three conditions depending on the −1 oxygen concentration in the water: (1) normoxic conditions (>2 mL O2 L ), (2) hypoxic conditions (>0 −1 −1 <2 mL O2 L ), (3) anoxic conditions (≤0 mL O2 L ) and (4) euxinia (Figure 3B) (Snoeijs-Leijonmalm and Andrén 2017). Decreasing oxygen concentrations affects the chemical and biological processes occurring in the water and the sediment, since it regulates the redox potential of present chemicals involved in redox reactions (Doods 2002a).

A B Figure 3. A) Usually occurring normoxic sediment zones: oxic zone (yellow); oxidised suboxic zone (brown) and sulphidic anoxic zone (black). B) Usually occurring zones within the sediment with varying oxygen concentration: normoxic conditions with oxic, suboxic and anoxic zones; hypoxic conditions with suboxic zone; anoxic conditions with a thin suboxic zone, and euxinic conditions with sulphide freely diffusing upwards to the overlying water (Snoeijs-Leijonmalm and Andrén 2017). Illustration: Johanna Hedberg.

2- 2+ In normoxic conditions in marine sediments, most phosphate (PO4 ) is bound to iron (Fe ), which makes the phosphate unavailable for assimilation by organisms and the sediments function as a sink. In oxygen depleted conditions phosphate is released and bioavailable again, and the sediment functions as a source of phosphate (Reed, Slomp, and Gustafsson 2011). In normoxic conditions biological processes concerning the degradation of organic material are performed by aerobic bacteria, using oxygen as oxidiser because of its high energy yield. When oxygen is depleted, anaerobic bacteria gradually take over the degradation of organic material. Anaerobic bacteria use other elements than oxygen as oxidisers - 4+ 2+ 3+ 2- that are consumed in an order following the decrease in energy yield: (O2), NO3 , Mn , Fe , Fe , SO4

, CO2 (Doods 2002b; Snoeijs-Leijonmalm and Andrén 2017). These oxidisers can be found vertically distributed in the sediment in this particular sequence. Oxygen is found in the oxic zone in the surface - 4+ 2+ 3+ 2- sediment, nitrate (NO3 ), manganese (Mn ) and iron (Fe , Fe ) in the suboxic zone, sulphate (SO4 ) in a third sulphidic anoxic zone (Figure 3A), followed by a fourth deep methane zone where carbon 4 dioxide (CO2) is formed into methane (CH ) (Figure 4A). The sediment zones are separated by boundaries: (1) the sediment water interface (SWI) separates the overlaying water from the sediment, (2) the oxygen penetration depth (OPD) separates the oxic zone from the suboxic zone and is the maximum depth where oxygen is detected, and (3) the sulphide apparent appearance boundary (SAAB) separates the suboxic zone from the sulphidic anoxic zone and this is where the sulphide concentration

3 appears (Figure 4B). Each of these zones varies in vertical expansion depending on oxygen concentration (Figure 3B) (Thamdrup, Fossing, and Jorgensen 1994; Schneider et al. 2017).

A B

- 4+ 3+ 4 Figure 4A) Vertical distributions of oxidants (O2, NO3 , Mn and Fe ) and reduction products (H2S and CH ) in the sediment. The x-axis shows the concentration and the y-axis shows the depth in the sediment starting with centimetres from the surface and then metres. The gradually changing colour show the depth expansion of the three sediment zones: oxic, suboxic and anoxic (Schneider et al. 2017). Illustration © Gregor Rehder 4B) The boundaries separating the zones within a normoxic sediment: sediment water interface (SWI) is found between the overlaying water and the sediment surface; oxygen penetration depth is found in the end of the oxic zone and sulphide apparent appearance boundary is found between the suboxic and anoxic zone. Illustration: Johanna Hedberg

Sediments exposed to long-term anoxic conditions can turn into euxinic conditions. In this state free sulphide (H2S) diffuses upwards in the water column from the sediment (Snoeijs-Leijonmalm and Andrén 2017) and causes migration of fauna due to its toxicity. Sulphide is toxic to most organisms since it diffuses freely across membranes and interfere with cellular processes, where it inhibits aerobic respiration enzymes in the mitochondria (Bagarinao 1992). Sulphur (S) is among the most abundant macronutrient on Earth and is found as sulphate in water which is its second largest reservoir (Muyzer and Stams 2008). The sulphur cycle includes combination of both chemical and biological processes (Jørgensen, Findlay, and Pellerin 2019). Sulphate is both assimilated and used as an energy source by organisms in redox reactions. In anoxic sediments sulphate-reducing bacteria are widespread and use sulphate as a terminal electron acceptor in the degradation of organic material, resulting in the production of sulphide (Muyzer and Stams 2008). In this reduction with several steps, HS- and S2- ions are also formed depending on the pH. Since S2- is a strong base it takes up hydrogen ions and the - formation of H2S increases. At a pH of 7, the H2S and HS ions are formed with proportions of about 30 % to 70 %, respectively (Schneider et al. 2017). The formation of the different sulphide species also depends on the temperature and salinity in the water and has to be accounted for together with pH, when - 2- calculating total hydrogen sulphide concentration ΣH2S =[H2S] + [HS ] + [S ] (Gerlach 1994 read in Jeroschewski, Steuckart, and Kühl 1996; Jeroschewski, Steuckart, and Kühl 1996).

However, there are other bacteria that can oxidise the sulphide back to sulphate when oxygen levels rise again (Jørgensen, Findlay, and Pellerin 2019), such as the recently discovered cable bacteria (Pfeffer et al. 2012). The aerobic cable bacteria belong to the Desulfobulbaceae family and are a multi-cellular filament building sulphide oxidiser (Pfeffer et al. 2012). The sulphide oxidation, occurring in the transition zone between the suboxic and anoxic zone, generates electrons

2- - + (1) 0.5 H2S + 2 H2O à 0.5 SO4 + 4 e + 5 H that can be passed internally through the vertical filament of the cable bacteria to the other end, in the transition between the suboxic and oxic zone,

- + (2) O2 + 4 e + 4 H à 2 H2O where they are consumed in the reduction of oxygen to generate water as end product (Figure 5). This consumption of protons generates a distinct pH peak in the transition from the oxic to the suboxic zone and is a strong indicator for the activity performed by cable bacteria (Nielsen et al. 2010; Meysman et al. 2015). Sending electrons through their filaments is advantageous to cable bacteria because it allows access to the major energy sources of oxygen in the sediment surface and sulphide in the subsurface sediment (Pfeffer et al. 2012). This vertical distribution of cable bacteria filaments creates a suboxic oxidised zone between the separated redox reactions in the sediment (Nielsen et al. 2010; Pfeffer et al. 2012; Seitaj et al. 2015). This zone is called suboxic because in oxic conditions it is found below the oxic zone (Nielsen et al. 2010) which partially is oxidised but still anoxic (Jørgensen, Findlay, and Pellerin 2019). Cable bacteria have been shown to expand this zone by the filaments growing further down in the sediment as they consume sulphide (Seitaj et al. 2015; Marzocchi et al. 2018). Thereby they delay the release of sulphide (Seitaj et al. 2015), which is thought to facilitate the recolonization of other organisms in reoxygenated sediments (Figure 5) (Seitaj et al. 2015; Marzocchi et al. 2018) such as bioturbating infauna.

Figure 5. From left to right: Oxygen depleted sediment with cultivating cable bacteria oxidising sulphide to sulphate in the transition zone between the anoxic and the suboxic zone, resulting in expansion of the suboxic zone that eventually might lead to recolonization of other organisms (Seitaj et al. 2015; Marzocchi et al. 2018). Illustration: Johanna Hedberg

As concluded in a review by Reise (2002), bioturbation by infauna is important since the walls created by animal activity in the sediment double the sediment surface towards the overlying oxygenated water

5 and thereby increases the diffusion of oxygen into the porewater. Increased diffusion of oxygen expands the oxic zone deeper into the sediment (Reise 2002) allowing aerobic respiration driven degradation of organic material (Snoeijs-Leijonmalm and Andrén 2017), and delaying the release of toxic sulphide (Bagarinao 1992). Effects of bioturbation by macrofauna on sediment biogeochemistry are relatively well understood (e.g. Bonaglia et al. 2013) while the knowledge regarding meiofaunal bioturbation is less understood, especially in the Baltic Sea (Janas et al. 2017).

Meiofauna is the most diverse and abundant group of organisms, and generally consists of nematodes (60 % - 90 %) and copepods (10 % - 40 %) (Coull 1999). Meiofauna is grouped by their body size (>0.04 and <1 mm) and the group comprises taxa from various phyla (Hulings and Gray 1971). Most of the meiofauna are active in the oxic zone in the sediment (Coull 1999), but some can be found in the suboxic zone in association to burrows from macrofauna (Meyers, Powell, and Fossing 1988). A number of Nematoda taxa seem to be relatively tolerant to sulphide and can be found close to the sulfidic anoxic zone in the sediment (Bagarinao 1992). Tietjen (1980 read in a review by Coull 1999) suggests four meiofaunal activities that can enhance bacterial growth and thereby stimulate organic matter mineralisation: (1) meiofauna changes the structure of organic material which makes it more accessible for the bacteria and thus increases their activity, (2) meiofauna excretes nutrients directly accessible for the use of bacteria, (3) meiofauna produces mucus which attracts and sustains the growth of bacteria and (4) meiofauna acts as a vertical transporter between the overlying water and the sediment via their bioturbation. Meiofauna presence has been shown to increase the diffusion of solutes and reactions within the oxic zone (Aller and Aller 1992). Even though several studies (e.g. Nascimento, Näslund, and Elmgren 2012; Piot, Nozais, and Archambault 2014; Bonaglia et al. 2014) and reviews (e.g. Coull 1999; Schratzberger and Ingels 2018) suggests that meiofauna has an important role in the aquatic ecosystems, meiofaunal role in Baltic Sea biogeochemistry is poorly understood (Janas et al. 2017). More recent studies have also shown that meiofaunal presence enhances the bacterial remineralisation in several ways (Nascimento, Näslund, and Elmgren 2012; Bonaglia et al. 2014), however no studies have investigated the effect of meiofauna on the activity and presence of cable bacteria.

Aim and research questions The poor knowledge of meiofaunal activity and their effect on bacterial community structure in reoxygenated sulfidic sediments together with the presence of cable bacteria, the aim was to answer the following research questions: (1) Will different abundances of meiofauna and bioturbation expand the oxic and suboxic zone? (2) Are meiofauna and cable bacteria co-existing in recently reoxygenated sediments? (3) Will different abundances of meiofauna and bioturbation effect the bacterial community structure?

Methods

Sampling site and bottom water conditions Sampling was conducted in Tvären which is located in the western side of the Baltic Proper in proximity of the Askö Marine Laboratory about 70 km south east of Stockholm, Sweden (Figure 6A). Tvären can be defined as an inshore embayment and is a ~80 m deep crater formed by a meteoritic impact 450 million years ago (Lindström et al. 1994). The topography of Tvären make the sediments in the centre to accumulation, which usually have high accumulation rates of organic materials, both vertically from the water surface and laterally from the surrounding inclining transport bottoms (Figure 6B). Tvären is also suffering from oxygen depletion due to degradation of organic material from elevated algal blooms because of an excess of nutrient loading mainly from the outer sea (VISS 2019).

A B Figure 6. A) Sampling site locations. 1) overview of the Baltic Sea and location of Tvären (red point) on the western side of the Baltic Proper, 2) showing Tvären as an embayment with its inshore position, 3) Tvären with the two sampling sites at A) 77 m depth (58 46.3116 N, 017 25.8471 E) and B) 38 m depth (58 47.0643 N, 017 24.6370 E). Google Maps. B) Bathymetric map of the embayment Tvären. The black lines connect areas with equivalent depth and the numbers show the depth of that area (Flodén, Tunander, and Wickman 1986).

To answer the research questions in this thesis high sulphide concentration in the sediment and low oxygen concentration in both sediment and bottom water was desired. Therefore, Tvären was chosen since it suffers from eutrophication due to overload of nutrients from surrounding landmass and the open sea (VISS 2019). The surplus of nutrients leads to algal blooms and as a consequence the bottom water oxygen is consumed by the degradation of settled organic material, which results in sediment areas of oxygen depletion (VISS 2019). Since a number of previous works on meiofauna abundance and composition has been conducted in the area between Tvären and Askö Marine Laboratory (Olafsson and Elmgren 1997; Nascimento, Näslund, and Elmgren 2012), meiofauna was also chosen to be collected in Tvären to be able to compare results.

Sampling was performed 10 of October 2018 onboard R/V Limanda, Askö Marine Laboratory, and the aim was to sample in the deepest part in centre of Tvären to collect hypoxic sulfidic sediment. Bottom water parameters at this station were carefully measured with a digital multi-meter in the cores right after sampling. Salinity was 7 and bottom water oxygen ranged from 9.0 mg L-1 to 9.9 mg L-1, which was higher than expected. The bottom water temperature ranged from 6.8 °C to 8 °C, while the surface water was 5 °C. However, the obvious smell and black colour revealed the presence of sulphide in the sediment.

Sediment collected for experimental setup Sediment cores were collected from 77 meters depth (58 46.3116 N, 017 25.8471 E) in PVC core liners using a multi corer (surface area of 63.6 cm2) and one smaller core (surface area of 16.6 cm2) was used to sub-sampled each multi core. A total of 15 smaller cores were collected where 12 were aimed for the four treatments, one for sediment porosity and total organic carbon content and two extras (for practical practising and if one core would be lost in some way). The cores were capped on both ends and isolated from daylight in a large Styrofoam box with cool packs, while sampling meiofauna.

Sampling for meiofauna individuals Meiofauna was sampled from the same basin but at a shallower depth of 38 m (58 47.0643 N, 017 24.6370 E) using the multi corer with PVC core liners (surface area of 63.6 cm2). Sediment from the top two centimetre from six cores were sliced off. The slices were covered with bottom water and stored in capped plastic containers together with the cores until arriving at Stockholm University later that day. Ahead of the transportation back to Stockholm University, filtered bottom water from 20 m depth with a salinity of 7 was collected for required laboratory work for the experiment at Askö Marine Laboratory.

At Stockholm University, all cores were placed in a large incubation tank, filled with ~ 20 L of collected in situ bottom water, in a thermo-constant chamber set to 8 °C. Before placed in the incubation tank, the cores were rinsed from left-over sediments on the outside to eliminate contamination of nutrients. To avoid resuspension of the sediment surface, the cores were carefully uncapped only after being completely submerged in in situ bottom water with a salinity of 7. A pump for circulation and aeration was added and oxygen levels in the water during the experiment was 10.9 mg L-1.

Sediment properties The core sampled for sediment properties was equally treated as the other cores before slicing for sediment properties. The core was sliced every 0.5 cm while avoiding contamination between layers and intact sediment of 4 ml was subsampled using a cut off syringe. The sediment was placed on an aluminium tray and left to dry in the oven at 75 ºC for about 24 hours and then placed in the furnace at 450 ºC for about 15 hours. The properties analysed were sediment density, porosity, vertical distribution of organic matter and sediment water content.

Table 1. Experiments, methods used and section where to find method details. Experiment Aim Method See section 1 OPD, SAAB, Sediment 1) Micro sensor profiling 1) Sediment microprofiling for O2, pH and ΣH2S characteristics 2) Sediment core slicing 2) Sediment properties 2 Cable bacteria densities FISH, Inverted epi-fluorescence Cable bacteria analyses and counting microscopy 3 1) Meiofauna extraction, 1) Levasil extraction 1) Meiofauna extraction and addition to sediment 2) counting and identification 2) Stereomicroscopy cores 2) Experiment termination, sediment sampling for bacterial analyses and meiofauna counting 4 Bacterial diversity DNA extraction, sequencing Bacteria community composition analyses

Meiofauna extraction and addition to sediment cores Two days after sampling, meiofauna extractions were done following the protocol by Nascimento, Näslund, and Elmgren 2012 by sieving the collected meiofauna sediment through a 40-µm sieve. The meiofauna and sediment retained on this sieve were then submerged for 5 min in a 7 % solution of

MgCl2, prepared by mixing 250 ml of in situ bottom water and 18.5 ml of MgCl2. The MgCl2 solution have an anaesthetic effect on the meiofauna, which detaches them from sediment particles and prevents them from swimming. The meiofauna and sediment in the sieve was then rinsed with in situ bottom water and washed into an E-flask (300 ml for 0.5 slice and 500 ml for 1 slice of sediment) with Levasil® 200A 40 % colloidal silica solution (H. C. Starck SilicaSol GmbH) with a density of 1.21, as described in (Nascimento et al. 2012). The E-flask covered with parafilm was turned upside down several times to mix the meiofauna with the Levasil® and was then left to settle for 5 min. The higher density of the Levasil® solution, makes the meiofauna float to the surface while the heavier sediment particles will sink to the bottom. After 5 min, the top 3 to 4 cm of Levasil® solution was poured onto a 40-µm sieve, the retained meiofauna and sediment were rinsed with in situ bottom water to remove the Levasil®, and washed into a 50 ml Falcon tube. The sieved Levasil® was then poured back into the E-flask with the remaining sediment and meiofauna and the procedure was then repeated twice, the last repetition with a 20 min settling time. The remaining Levasil® solution in the E-flask after the last extraction was poured through a 250-µm sieve on top of a 165-µm sieve, to get rid of bigger particles and to gather Ostracods. The sediment and meiofauna retained in the 165-µm sieve, was rinsed with in situ bottom water and washed into a 50 ml Falcon tube. Until all slices were sieved the extracted meiofauna were stored in the climate chamber at 8 °C.

When extracting meiofauna, a small amount of fine sediment particles and organic matter also floats because of the higher density in the Levasil® and become extracted with the targeted meiofauna. Therefore, the DEBRIS treatment was included to control if the added amount of debris could have an effect on the sediment, like changing the rate of oxygen diffusion. The six slices of sediment collected for the meiofauna treatments were divided to get an approximately two times higher abundance in LM

(0.5 slice) and six times higher abundance in HM (1.5 slice) than in the control cores (CTR and DEBRIS). Before adding the treatment to the sediment cores, sieved sediment (debris) was collected to the DEBRIS treatment from the sediment left-overs after extracting the meiofauna. The collected amount of debris corresponded to the amount of extracted meiofauna, ~5 ml. Water was removed to accommodate the volume of debris and meiofauna that was carefully poured into the corresponding cores (see below). The cores were left to settle before placing them back into the incubation tank.

Experimental design The experiment had four treatments: (1) extracted meiofauna with low abundance (LM, n = 3), (2) extracted meiofauna with high abundance (HM, n = 3), (3) one intact control (CTR, n = 3) and (4) one control with addition of sediment extracted from meiofauna (DEBRIS, n = 3). The unmanipulated CTR treatment was included as an experimental control. During the first three weeks microprofiles for O2, pH and H2S was conducted to examine how sediment chemistry was affected by the different abundances of meiofauna. The experiment started two days after sediment sampling when meiofauna was added to the cores and ended after 29 days with the slicing of all cores for further analyses (meiofauna counting and identification, bacterial composition, cable bacteria counting and sediment properties).

Sediment microprofiling for O2, pH and �H2S The sediment cores were measured for O2, pH and ΣH2S profiles once a week for three weeks (W1, W2 and W3) with three replicate profiles in each core per measurement. The first week O2, pH and ΣH2S was measured separately, while the two following weeks O2 was measured separately and pH and ΣH2S together by combining the probes. Profiling was performed using an automatic microprofiling unit (O2:

OX-500, pH: pH-100, H2S: H2S -100; Unisense, Denmark) attached to a computer-controlled stand with place for a water filled and temperature-controlled aquarium. Each day of measurement the microprofiling unit was calibrated for each substance measured according to the product manual. One sediment core at the time was placed in the aquarium where an air-pump carefully circulated the overlying water to create a steady diffusive boundary layer (DBL) on the surface of the sediment. O2 was vertically measured in the sediment every 100 µm and pH and ΣH2S every 250 µm.

Oxygen penetration depth (OPD) was determined by the oxygenated sediment turning into anoxic -1 conditions ([O2] < 1 µmol L ). The sulphide apparent appearance boundary (SAAB) was determined by -1 locating the layer where the sediment became sulfidic ([H2S] > 1 µmol L ).

Experiment termination, sediment sampling for bacterial analyses and meiofauna counting To prepare the cores for slicing they were emptied for water and the last 2 cm of water was carefully removed with a syringe and poured in a 40-µm sieve until preservation. The cores were sliced into two slices (0 - 1 cm and 1 - 2 cm) avoiding contamination between slices by cleaning the equipment used

10 with Ethanol (70 %). The top slice of sediment was gently homogenised and from each core, 2 ml of sediment was sampled for the bacterial community composition analyses and 0.5 ml of sediment was sampled and mixed with 0.5 ml Ethanol (99.5 %) for cable bacteria analyses. Precautions were taken to avoid contamination and cross-contamination and samples were kept frozen at - 20°C until further processing. Each slice was preserved separately with 5 % Formaldehyde and was kept in climate chamber at 8 °C until counting of meiofauna. The Formaldehyde contained Rose Bengal which stains the live meiofauna in the sediment in a bright pink colour. This helps locating the meiofauna in the debris during counting and identification.

Extraction for counting and identification of the meiofauna from preserved sediment slices was performed as described in the Meiofauna extraction and addition to sediment cores section. One slice at the time was extracted and kept at 4 °C. Extracted meiofauna was counted using a 50x binocular stereomicroscope and identified to group level. Counted meiofauna was preserved in 5 % Formaldehyde containing Rose Bengal and stored at 8 °C for the time being.

Bacteria community composition analyses Total environmental DNA were extracted from 0.25 g of sediment samples using a DNeasy PowerSoil Kit (QIAGEN) with included protocol and purity was controlled using a Nanodrop.

Library preparations started with that one library aimed for the 16S ribosomal RNA gene was arranged according to the combined methods from Creer and Sinniger (2012) and Herlemann et al. (2016). Primer pair 341F/805R was used to amplify regions of the 16S rRNA gene (Herlemann et al. 2011) (Table 2). Polymerase chain reaction (PCR) round one was performed using primers adjusted with an Illumina Adapter Sequence in an initial PCR to amplify the 16S gene aimed for. Conditions during the thermocycling round one was: 30 s at 98°C, followed by 12 cycles of 10 s at 98°C, 30 s at 50°C, 30 s at 72°C. Q5 HS High-Fidelity Master Mix (New England BioLabs) was used to prepare the library following the manufacturer´s protocol and PCR reactions was performed in a BioRad T100 Thermal Cycler (BioRad Laboratories). The amplicons from the first round were cleaned with the addition of 0.1 µL Exonuclease 1 (New England BioLabs) and 0.2 µL Thermosensitive Alkaline Phosphtase (Promega), and to finalise the reaction the amplicons were incubated for 15 min at 37°C followed by 15 min at 74°C. PCR round two was performed with the use of indexing primers as described by Herlemann et al. (2016) so that each sample was prepared with forward and reverse index sequences in a unique combination. Conditions during the thermocycling round two was: 3 min at 95°C, 15 cycles of 30 s at 95°C, 30 s at 55°C, 30 s at 72°C, and 5 min at 72°C. Cleaning of the final amplicons was performed using Agencourt AMPure XP magnetic beads (Beckman Coulter) and to measure the concentrations of the amplicons a Qubit 2.0 Fluorometer (dsDNA BR Assay Kit, Invitrogen) was used. Then the samples were standardized, pooled together and sent for sequencing.

Table 2. Primers used in this study during bacteria community structure analysis.

Primer Sequence (5′→3′) Target gene Reference Round one forward adapter ACACTCTTTCCCTACACGACGCT Indexing overhang Illumina Inc. 2019 CTTCCGATCT NNNNN Round one reverse adapter GTGACTGGAGTTCAGACGTGTG Indexing overhang CTCTTCCGATCT 341F CCTACGGGNGGCWGCAG Bacterial 16S r RNA Herlemann et al. 2011 805R GGACTACHVGGGTWTCTAAT Bacterial 16S r RNA

The sequencing was performed by the National Genomics Infrastructure (NGI) in Stockholm, Sweden (SciLifeLab, Stockholm) using an Illumina MiSeq instrument (v3). 16S rRNA sequence reads were demultiplexed by NGI, and further processing was carried out using the DADA2 pipeline (version 1.10.1, Callahan et al 2016) implemented in R (version 3.5.1). DADA2 was used to trim raw sequences to remove low quality bases and filtered using the following parameters: truncLen = c (290,210), maxEE = c (2,2), trimLeft = c (8,8), minFoldParentOverAbundance = 4 and allowoneoff = TRUE. This procedure followed by merging the paired-ends and removal of chimers from the dataset. The SILVA database (r132, Quast et al. 2013) and the DECIPHER package (v 2.10.2; Wright 2016) was used to carry out taxonomic assignments on the sequence variants on the 16S rRNA amplicon. To detect differences in bacterial community composition between treatments a Non-metric multidimensional scaling (NMDS) ordination was done after loading of bacteria amplicon sequence variants data into the R package phyloseq 1.24.2 (McMurdie and Holmes 2013). NMDS plots of Bray-Curtis dissimilarity, based on the sequence variants relative abundance were constructed using the ‘ordination’ and ‘plot.ordination’ function in phyloseq.

Cable bacteria analyses and counting Fluorescence in situ hybridisation (FISH) was conducted to estimate the abundance of cable bacteria, at Aarhus University, Aarhus, Denmark at the Department of Bioscience – Microbiology. Counting of duplicates was performed at DEEP, Stockholm University.

To homogenise the samples and separating particles from each other, a mild ultrasonic treatment of ~30 % power was used with 3 cycles x 20 s with 5-10 s between cycles. Subsamples of 100 µl was added to a mixture of 880 µl of sodium pyrophosphate buffer and 20 µl of agarose 1 %. FISH protocol was then completed as in previously published protocols (Pernthaler et al. 2001). The positive control and the samples were marked with two probes: (1) EUB 338mix (I, II, III), positive control for all bacteria, (2) DSB 706 (Loy et al. 2002) positive for all sulphate-reducing bacteria, the negative control was added with a solution called Non, with no probe. All samples were also stained with DAPI for DNA in general. For every FISH protocol started, at least one slide had one negative and one positive control to be able

12 to discover possible mistakes while following the FISH-protocol. Cable bacteria analysis at Aarhus University was done in an Axiovert 200 inverted microscope for transmitted light and epifluorescence (Carl Zeiss, Göttingen, Germany) using a 40x lens (with 10x in the binocular equals a total magnification of 400 times), while analysis at Stockholm University was done in a Nikon Inverted Microscope Eclipse Ti-U. To estimate the filament densities of cable bacteria within the sediment (meters of filaments per cubic centimetre of sediment (m cm-3)), a previously used method for cable bacteria (Pfeffer et al. 2012) was used based on the line-intersection method (Newman 1966). One line was selected from the grid in the eyepiece of the microscope and used for counting every filament crossing while travelling from one side of the well to the other, both horizontally and vertically, a total of ten times.

Statistical analyses Differences between meiofauna abundances, OPDs, SAABs, ΣH2S concentrations, O2 consumption, methane fluxes and cable bacteria densities were tested using one-way analysis of variance (ANOVA) with an initial normality test (Shapiro-Wilk test) and following pairwise comparisons were tested with a post hoc test (Tukey test). SigmaPlot 11.0.Ink was used performing statistical analyses. Detection of statistical differences in bacteria community composition was done using PERMANOVA tests (9999permutations) with the adonis function in the vegan package (Oksanen et al. 2018).

Results

Sampling conditions −1 The expected hypoxic conditions (>0 <2 mL O2 L ) in the bottom water turned out to be normoxic, −1 since the oxygen measured from 6.3 to 6.93 mL O2 L in the collected cores. The also unexpected higher temperature of about 3 °C in the bottom water than in the surface water were most likely explained by the occurrences of stormy periods in the weeks before sampling day that mixed the water masses (SMHI 2019).

The core aimed for sediment properties was sliced down to three centimetres depth every half centimetre. The first 0.5 cm had a dry weight organic matter content of 19.0 %, at 0.5 - 1 cm and 1 – 1.5 cm depth the dry weight organic matter content was 16.9 % and 17.0 % and from 1.5 - 3 cm depth the organic content stabilised around 15.4 % (Table 3). The density in the first 0.5 cm was 1.04 g cm-3 and decreased to 0.97 g cm-3 between 0.5 - 1 cm depth and then stabilised at a density of ~1.0 g cm-3 from 1 - 3 cm depth. The porosity followed the same pattern, higher porosity of 0.93 φ in the first 0.5 cm,

13 decreased to 0.85 φ between 0.5 - 1 cm and then stabilised at ~0.90 φ from 1 - 3 cm depth. These properties can be seen as start conditions, as the sediment was not exposed to any manipulation.

Table 3. Sediment properties analysed every 0,5 cm: dry weight organic matter content (%), change of dry weight organic matter content between each layer of 0.5 cm (%), density (g cm-3), porosity (φ) and water content (%). Sediment depth Org. material Org. material Density Porosity Water (cm) (%) (change in %) (g cm-3) (φ) (%) 0 - 0.5 19.0 1.04 0.93 89.1 0.5 - 1 16.9 - 12 % 0.97 0.85 88.5 1 - 1.5 17.0 + 0.5 % 1.03 0.91 88.5 1.5 - 2 15.4 - 10 % 1.01 0.89 87.8 2 - 2.5 15.3 - 0.5 % 1.01 0.90 89.3 2.5 - 3 15.5 + 0.1 % 1.04 0.91 87.6

Meiofauna abundance Extracted meiofauna from the top centimetre (0 – 1 cm) of sediment at the end of the experiment showed that a gradient of individuals was added to the cores in the different treatments (Table 4, Table S1 in supplementary info 2 for sediment layer 1 - 2 cm). The difference in meiofauna abundance among treatments significantly differed (ANOVA, F3,8 = 6.088, p = 0.018), the CTR (Tukey HSD: p = 0.023) and DEBRIS (Tukey HSD: p = 0.027) significantly differed from HM, while LM did not (Tukey HSD: p = 0.159). The added amount of meiofauna was about 15 times higher in HM, 6 times higher in LM and 1.5 times higher in DEBRIS than in the CTR treatment.

Table 4. Meiofauna abundance in individuals per square meter (ind. 10-3 m2) in the first centimetre (0 - 1cm) of sediment in each treatment; CTR, DEBRIS, LM and HM. Values are mean ± standard deviation (n = 3). * Most of the organisms were identified to the genus Bosmina. CTR DEBRIS LM HM Meiofauna (ind. 10-3 m-2) (ind. 10-3 m-2) (ind. 10-3 m-2) (ind. 10-3 m-2) Nematods 77.8 ± 31.3 142.0 ± 60.0 661.8 ± 340.1 1793.0 ± 397.1 Ostracods 0 0 15.8 ± 22.7 9.3 ± 4.3 Copepods 16.5 ± 8.2 17.4 ± 5.7 16.5 ± 7.6 21.4 ± 13.2 Kinorhyncha 0.5 ± 0.4 0.5 ± 0.4 40.2 ± 25.7 145.9 ± 53.4 Cladocerans* 33.0 ± 10.8 27.2 ± 3.0 90.2 ± 72.0 56.7 ± 28.5 Halacaridae 0.5 ± 0.8 4.0 ± 3.1 3.0 ± 4.6 3.7 ± 4.3 Total 128.3 ± 45.7 191.0 ± 65.5 827.5 ± 431.6 2030.0 ± 500.6

Figure 7. Pictures taken during identification and counting of meiofauna through the stereomicroscope eyepiece. From left: one square centimetre (1 cm2, in blue) of debris and meiofauna (red circle around one stained Copepod); Halacaridae; Cladoceran; Kinorhyncha; Copepod carrying eggs; Ostracod; Nematode. Pictures are not to scale.

Nematodes were the most abundant phylum in all treatments, comprising on average of 61 % ± 24 %, 74 % ± 31 %, 80 % ± 41 % and 88 % ± 20 % of the total abundance in CTR, DEBRIS, LM and HM, respectively (mean ± standard deviation (SD), n = 3). The second most abundant species in HM and LM was Kinorhyncha standing for 7 % ± 3 % and 5 % ± 6 % respectively, whereas in the CTR and DEBRIS Copepods were the second most abundant species standing for 13 % ± 18 % and 9 % ± 9 % respectively. The extracted meiofaunal community found in this experiment, correspond to previously reported compositions from the same depth in the Askö area, e.g. Olafsson and Elmgren 1997.

Even though Cladocerans not belong to the group meiofauna they were counted because unusually high numbers were observed in the sediment. Cladocerans were standing for 26 % ± 24 %, 14 % ± 5 %, 11 % ± 17 % and 3 % ± 6 % of the total abundance in CTR, DEBRIS, LM and HM, respectively.

Sediment profiles of O2. pH and �H2S. OPDs. There was an effect of treatment on OPDs in each of the three weeks (ANOVA: W1: F3,8 =

18.429, p = < 0.001; W2: F3,8 = 8.427, p = 0.007; W3: F3,8 = 9.760, p = 0.005) (Figure 8 for OPDs and

Figure 11 for sediment profiles of O2). At W1 the OPDs in the CTR (Tukey HSD: p = 0.020) and LM (Tukey HSD: p = 0.031) significantly differed from HM while DEBRIS (Tukey HSD: p = 0.990) did not. Same effect was seen at W3, CTR (Tukey HSD: p = 0.003) and LM (Tukey HSD: p = 0.029) significantly differed from HM while DEBRIS (Tukey HSD: p = 0.056) did not. While at W2 CTR (Tukey HSD: p = 0.007) and DEBRIS (Tukey HSD: p = 0.022) significantly differed from HM while LM (Tukey HSD: p = 0.057) did not.

Microprofiling of the sediment showed that the OPDs significantly expanded deeper over the weeks within the CTR, LM and HM treatments (ANOVA: CTR: F2,6 = 11.096, p = 0.010; F2,6 = 5.403, p =

0.046; HM: F2,6 = 6.395, p = 0.033) but not in the DEBRIS (ANOVA: DEBRIS: F2,6 = 0.970, p = 0.431). At W1 the deepest OPD was found in the LM treatment (2.2 ± 0.3 mm, n = 3) while the HM treatment had the deepest OPDs at W2 (2.3 ± 0.4 mm, n = 3) and at W3 (2.3 ± 0.2 mm, n = 3).

Week 1 Week 2 Week 3

CTR DEBRIS LM HM CTR DEBRIS LM HM CTR DEBRIS LM HM 0 0 0

1 1 1

2 2 2 Sediment depth (mm) 3 3 3

Figure 8. Oxygen penetration depths in the four treatments; CTR (green), DEBRIS (blue), LM (yellow), HM (red) at the three points of measure; week 1, week 2 and week 3. Zero at the y-axis represents the surface water interface and the bars -1 show how deep the oxygen (O2>1 µmol L ) penetrates the sediment in mm (mean ± standard error, n = 3).

SAABs. There was no effect of the treatment on the SAABs in each of the week (ANOVA: W1: F3,8 =

3.433, p = 0.072; W2: F3,8 = 1.522, p = 0.282; W3: F3,8 = 1.010, p = 0.437) or within treatments over the weeks (ANOVA: CTR: F2,6 = 1.043, p = 0.409; DEBRIS: F2,6 = 1.983, p = 0.218; LM: F2,6 = 2.971, p = 0.127; HM: F2,6 = 1.016, p = 0.417). Still the deepest SAABs were found in the HM treatment in all weeks (Figure 9 for SAABs and Figure 11 for sediment profiles of ΣH2S).

Week 1 Week 2 Week 3

CTR DEBRIS LM HM CTR DEBRIS LM HM CTR DEBRIS LM HM 0 0 0

2 2 2

4 4 4

6 6 6 Sediment depth (mm) 8 8 8

Figure 9. Sulphide apparent appearance boundary in the four treatments; CTR (green), DEBRIS (blue), LM (yellow), HM (red) at the three points of measure; week 1, week 2 and week 3. Zero at the y-axis represents the surface water interface and -1 the bars shows the depth of sulfidic free sediment (ΣH2S<1 µmol L ) in mm (mean ± standard error, n = 3).

�H2S. There was no significant difference in ΣH2S concentration within treatments in each week at both

5 mm (ANOVA: F3,8 = > 2.263, p = > 0.05) and 7.5 mm (ANOVA: F3,8 = > 3.025, p = > 0.05) (Figure

10 and Figure 11 for sediment profiles of ΣH2S). There was no significant difference in ΣH2S concentration between weeks in the CTR, DEBRIS and LM at both 5 mm (ANOVA: F2,6 = > 1.954, p

= > 0.05) and 7.5 mm (ANOVA: F3,8 = > 4.345, p = > 0.05) but the ΣH2S concentration significantly differed in the DEBRIS at both 5 mm (ANOVA: F3,8 = 5.666, p = 0.041) and 7.5 mm (ANOVA: F3,8 =

18.986, p = 0.003). Even though the DEBRIS treatment showed a significant decrease in ΣH2S concentration from W1 to W3 and the other treatments did not, there is an overall trend. The trend indicates that the HM treatment lowers the ΣH2S concentrations in the sediment already at W1 and that the CTR, DEBRIS and LM treatments also, but more slowly, lowers the ΣH2S concentrations until W3. pH. While microprofiling the cores pH peaks just beneath the sediment surface were observed in the HM treatment at W1 at 0.75 mm depth (8.3 ± 0.2, n = 3). Also, at W3 pH peaked in the CTR treatment at 0.5 mm depth (8.0 ± 0.3, n = 3) and in the DEBRIS treatment at 0.75 mm depth (8.0 ± 0.1, n = 3) (Figure 11). These pH peaks are indications of activity by cable bacteria releasing electrons as they reduce oxygen in the transition zone between the oxic and suboxic zone (Nielsen et al. 2010; Meysman et al. 2015).

CTR DEBRIS LM HM -1 -1 H S (µmol L ) -1 -1 H2S (µmol L ) 2 H2S (µmol L ) H2S (µmol L ) 0 50 100 150 200 250 0 50 100 150 200 250 0 50 100 150 200 250 0 50 100 150 200 250

5 mm Week 1 7.5 mm

0 50 100 150 200 250 0 50 100 150 200 250 0 50 100 150 200 250 0 50 100 150 200 250

5 mm Week 2 7,5 mm

0 50 100 150 200 250 0 50 100 150 200 250 0 50 100 150 200 250 0 50 100 150 200 250

5 mm Week 3 7,5 mm

Figure 10. ΣH2S concentration at 5 mm and 7.5 mm depth in the four treatments; CTR, DEBRIS, LM and HM (from left to right) at the three points of measure; week 1, week 2 and week 3 (from top to bottom) (mean ± standard error, n = 3).

pH was steady at 7.8 ± 0.0 (n = 12) in the overlying water 0.5 mm above the sediment surface throughout the experiment (Figure 11). pH started to decrease beneath the surface (or after the pH peaks) and eventually stabilised further down in the sediment. The range of pH stabilisation depth decreased in the different treatments from W1 to W3, ranging approximately from 2 mm to 7 mm at W1 and from 4 mm to 6 mm at W2, while at W3 pH stabilised around 6 mm depth. As for the stabilisation range of pH, at W1 pH ranged approximately from 6.5 to 7.2, in W2 from 5.5 to 6.2 and in W3 pH stabilised around 5.5. In the DEBRIS and HM treatment pH already stabilised at W2 around 5.5. The confined stabilisation depth and acidic environment in the sediment are also indications of activity by cable bacteria, because at this depth in the sediment cable bacteria release protons as they oxidise sulphide to sulphate (Nielsen et al. 2010; Meysman et al. 2015).

CTR DEBRIS LM HM

-1 0 100 200 300 400 0 100 200 300 400 0 100 200 300 400 0 100 200 300 400 O2 (µmol L ) 5 6 7 8 9 5 6 7 8 9 5 6 7 8 9 5 6 7 8 9 pH 0 100 200 300 0 100 200 300 0 100 200 300 0 100 200 300 -1 H2S (µmol L ) 0 0 0 0

2 2 2 2

4 4 4 4

6 6 6 6 Week 1 Depth (mm) 8 8 8 8

10 10 10 10

12 12 12 12

-1 0 100 200 300 400 0 100 200 300 400 0 100 200 300 400 0 100 200 300 400 O2 (µmol L ) 5 6 7 8 9 5 6 7 8 9 5 6 7 8 9 5 6 7 8 9 pH -1 0 100 200 300 0 100 200 300 0 100 200 300 0 100 200 300 H2S (µmol L ) 0 0 0 0

2 2 2 2

4 4 4 4

6 6 6 6 Week 2 Depth (mm) 8 8 8 8

10 10 10 10

12 12 12 12

2 2 2 2

4 4 4 4

6 6 6 6 Week 3 Depth (mm) 8 8 8 8 H2S 10 10 10 10 O2 pH 12 12 12 12

Figure 11. Sediment profiles of O2 (blue), ΣH2S (black) and pH (yellow) in the four treatments; CTR, DEBRIS, LM and HM (from left to right) at the three points of measure; week 1, week 2 and week 3 (from top to bottom). Zero at the y-axis represents the surface water interface and the top x-axis show oxygen concentration in micromole per litre (µmol L-1), middle -1 -1 x-axis shows pH and bottom x-axis show ΣH2S (µmol L ) concentration in micromole per litre (µmol L ) (mean ± standard error, n = 3).

Cable bacteria analyses The FISH analysis of the DNA extraction from the sediment samples showed positive results for presence of cable bacteria (Table 5). However, there were no significant difference in cable bacteria filament densities between the treatments (ANOVA: F3,8 = 0.964, p = 0.455). The insignificant different densities can be seen as positive results, because the equal densities of cable bacteria will not interfere with the results from the gradient of meiofauna.

Table 5. Average total length of cable bacteria filaments in meter per cubic centimetre sediment (m cm-3) in each treatment (CTR, DEBRIS, LM, HM). Values are in mean ± standard deviation (n = 3). Cable bacteria density Treatment m cm-3 CTR 118.6 ± 20.7 DEBRIS 111.6 ± 12.7 LM 96.3 ± 14.9 HM 112.8 ± 18.0

Bacterial community structure The gene pool of 16S rRNA characterised by the T-RFLP analysis showed that the bacterial community structure in the sediment was significantly different between the treatments (PERMANOVA: F2,8 = 2.8, p = 0.001), which suggest an effect from the treatment as shown by the completely separated clusters in figure 12. The large distance between the CTR and HM clusters indicates the most differentiated bacterial community structures, which probably is caused by the large difference in meiofauna abundance and meiofaunal bioturbation. The CTR and HM treatment, also, have the most diverse bacterial community structures as can be seen by the large distances of the dots within each cluster, followed by the LM treatment whereas the nearness of the dots within the LM cluster indicates less diverse bacterial community structure.

Figure 12. Non-metric multidimensional scaling showing the bacterial composition in the four treatments; CTR (red), DERBIS (green), LM (turquoise) and HM (purple). The distance between the treatment groups in the figure show the difference in bacterial composition. No overlap means no similarity. The distance between the dots within each treatment group show the difference in bacterial composition. Small spaces between the dots mean high similarity.

Discussion

Sampling site and sediment conditions Sediment in the collected cores were sulphidic even though bottom water was reoxygenated in the deep centre of Tvären. The sediment cores were black except for a thin grey layer at the sediment surface, which indicates that anaerobic bacteria were reducing sulphate into sulphide almost to the surface of the sediment core in the lack of oxygen (Muyzer and Stams 2008). The shallow sulphide appearances in the sediments detected by microprofiling in the beginning of the experiment are also signs that Tvären suffers from longer periods of oxygen depletion in the bottom water (Daniel J Conley et al. 2007; VISS 2019) impeding deepening of the oxic and suboxic zones. However, it seems like the reoxygenation and inverted temperature in the water can be explained by autumn storms (SMHI 2019) which previously been recorded in the area around Tvären (e.g. Olafsson and Elmgren 1997) and as a repetitive phenomenon in other stratified water bodies (e.g. Goodrich et al. 1987) affecting sediment biogeochemistry.

Sediment analyses show that the organic matter content decreased from the top to the second and from the third to the fourth half centimetre with about 12 % and 10 % in respective layer and with a layer of unchanged organic matter content in-between. This pattern might be an indication that meiofaunal bioturbation activates the aerobic bacterial degradation of the organic material in the top of the sediment layer (Coull 1999). This is supported by the higher porosity found in the top sediment layer, indicating higher meiofaunal bioturbation in the top layer of sediment, even in the unmanipulated cores. The second decrease of organic matter content occurred in the third sulphidic anoxic layer of sediment, where the sulphate reducing bacteria presumably are responsible for the degradation of the organic material (Aller 1994; Muyzer and Stams 2008). One thing to have in mind is that the core used for these analyses were unmanipulated and may not fully represent the sediment properties in the cores in the end of the experiment where meiofauna abundances were manipulated. However, when analysing the sediment properties, the organic carbon content was about 20 to 50 % higher in the centre of Tvären compared to the central Gotland Basin, where Leipe et al. (2011) describe that the “POC contents are exceptionally high”. Further, the water content in the sediments of Tvären (~90 %) correspond to what Leipe et al. (2011) estimated in the central Gotland Basins (>90%). The combination of high carbon content and high water content, makes the sediments fluffy and are characteristic for accumulation bottoms (Duplisea 2000; Leipe et al. 2011) which Tvären could be described as because of its topography. This type of sediment properties has been described to inhabit high numbers of meiofauna (Coull 1999), but in the sulphidic sediments in Tvären the abundance of fauna is highly reduced as can be seen by the meiofauna abundance in the CTR treatment.

Effects of meiofauna on porewater conditions The composition of meiofauna identified in this thesis is in line with what other studies have reported (Olafsson and Elmgren 1997; Nascimento, Näslund, and Elmgren 2012) but the abundance of meiofauna, even in the high meiofauna treatment, is under previous recorded number of individuals in the surrounding area (Ankar and Elmgren 1978; Olafsson and Elmgren 1997). As an example, Ankar and Elmgren (1978) sampled at the same depth (37 m) in the outer normoxic archipelago in proximity to Tvären and the Askö Marine Laboratory and found about three- and ten-times higher abundance compared to the abundance in the HM and LM treatment, respectively. Further they found about 20 times higher abundance in their deepest sampling site (50 m) compared to the unmanipulated control treatment in this thesis sampled at 77 m depth. Since Tvären embayment suffers from oxygen depletion (VISS 2019) the oxic and suboxic zone is reduced which probably forces the oxygen dependent meiofauna into the upper oxic zone (Coull 1999) away from the sulphidic anoxic toxic zone (Bagarinao 1992) known to reduce the abundance of Nematodes (Van Colen et al. 2009). This large difference in meiofaunal abundance is probably due to the unlike conditions at the sampling sites.

The abundance of meiofauna was found to be 1.5 time higher in the DEBRIS than in the CTR treatment which could be a consequence of natural variation at the sampling site. But, since the increase mainly consisted of Nematodes, they were most likely coming from the debris collected from the extracted sediment that was added to the DEBRIS treatment. Smaller individuals of Nematodes could have slipped through the sieve during extraction. This has been observed in previous extractions when trying to eliminate meiofauna from sediments (Nascimento, Näslund, and Elmgren 2012).

The significant differences between the OPDs in the CTR, LM and HM treatments seem to be mediated by the different abundances of meiofauna and meiofaunal bioturbation. The insignificant difference in cable bacteria densities and thus equal activity from the cable bacteria probably did not interfere with the effect from the meiofauna treatment. The increase in OPDs in the cores corresponds to the increasing abundance of meiofauna and bioturbation in the different treatment and the maintained OPDs at W2 and W3 suggests that the different meiofauna abundances and meiofaunal bioturbation affects the OPDs. This result is in line with other studies that show that presence of meiofauna in sediment in laboratory experiment (Aller and Aller 1992) and different abundances of meiofaunal bioturbation measured in in situ sediments (Rysgaard et al. 2000) increase the solute transport, like oxygen and thereby increases the OPDs.

Meiofauna are thought to be the first colonisers after bacteria in reoxygenated sediment due to their small body size which enables farther translocation with bottom water currents, compared to macrofauna (Van Colen et al. 2009). Nematodes are probably the first to recolonize as they, as a group, are more tolerant to sulphide (Bagarinao 1992) and since they are the most abundant taxonomic group. Meiofauna

21 seem to affect the sediment chemistry rapidly by their bioturbation compared to cable bacteria as seen in this thesis. As they are bioturbating the sediment they expand the oxic zone which creates a deepening of the different redox layers within the sediment that mediates the same facilitation of recolonization and delay of sulphide release as the cable bacteria.

The increase in OPDs with 1 mm in the unmanipulated CTR treatment from W1 to W2 and W3 was unexpected but could be an effect from the ~15 % elevated oxygen level in the thermal constant room, as reoxygenated in situ conditions were strived for. OPDs in the DEBRIS was expected to follow the development of the OPDs in the CTR treatment but possibly with a slightly expanded OPD due to the addition of debris from oxygenated conditions. The addition of oxygenated debris could also be the explanation for the deeper OPDs at W1 in the LM treatment. At W2 the OPDs decreases in depth in the LM treatment which could be a result from the increasing aerobic degradation due to high content of organic material per meiofauna, as discussed by Coull (1999).

There was no significant difference in the SAABs over time or within treatments, nevertheless, the SAABs in the HM treatment were the deepest during the three weeks which indicate a potential effect from the higher abundance of meiofauna and meiofaunal bioturbation. Maybe a significant result could have been achieved if the abundance of meiofauna in the HM treatment had been closer to natural abundances as recorded by Ankar and Elmgren (1978) or Olafsson and Elmgren (1997).

Cable bacteria on porewater conditions Cable bacteria filament densities found in this thesis are about 4 – 50 times higher when comparing densities found by Marzocchi et al. (2018) in the eastern Gotland Basin after a reoxygenation event in 2014. The presence of cable bacteria, in both Tvären and the eastern Gotland Basin, could be explained by cable bacteria normally being the first colonisers after reoxygenation events in sulfidic sediments (Seitaj et al. 2015; Marzocchi et al. 2018). This because of the high co-existence of their energy sources sulphate and oxygen (Pfeffer et al. 2012) and uninhabited environment. Since this is the second study finding cable bacteria in the northern Baltic Proper, conclusions regarding the densities are difficult to make as most of the previous experiments on cable bacteria densities have been conducted in laboratories with manipulated sediments. In a laboratory study by Schauer et al. (2014) cable bacteria filaments were estimated to over 2 km cm-3 and in a field study conducted in the southern North Sea by Malkin et al. (2014) in situ densities were 82 to 123 m cm-3, which, however, correspond to the densities in this thesis.

The insignificant differences in SAABs at W3 and despite the significant differences in OPDs at W3 together with the equal filament densities of cable bacteria, are suggesting an effect from the cable bacteria. Even though OPDs were shallower in the CTR, DEBRIS and LM treatment the SAABS

22 reached the same depth at W3, which suggests that cable bacteria alone are efficient in expanding the suboxic zone over time. Cable bacteria initially develop in the transition between the oxic and sulphidic anoxic zone (the OPD) and from there they grow downwards as they deepen the SAABs (Schauer et al. 2014). The zone of development (the OPD) was shallower in the CTR treatment than in the HM treatment, which might be an explanation why the CTR treatment reached the same depth of SAABs as in the HM treatment later.

The results of the ΣH2S concentrations are closely linked to the SAABs as both measurements are the concentration of ΣH2S. The results with similar ΣH2S concentrations at W3 can therefore lean on the same interpretation: that cable bacteria alone are as efficient in expanding the suboxic zone as their co- existence with meiofauna. The expansion of the suboxic zone also enables the growth of other sulphur oxidising bacteria that decreases the ΣH2S concentrations (Jørgensen, Findlay, and Pellerin 2019). However, to confirm this trend future studies with different densities of cable bacteria are needed to understand their effect on the ΣH2S concentrations and expansion of SAABs in co-existence with meiofauna.

Even though there was no significant difference in the ΣH2S concentrations at the two sediment depths between treatments or within weeks (except for the LM treatment at 7,5 mm depth), there was an obvious decrease in ΣH2S concentrations from W1 to W3 in all treatments. The low ΣH2S concentrations in the HM treatment already at W1 probably is a cause from the higher abundance of meiofauna and meiofaunal bioturbation rapidly oxygenating the sediment. The DEBRIS and LM treatment reach the same low level of ΣH2S concentration as the HM treatment at W2, which might be explained by the lower abundance of meiofauna and bioturbation having a slower effect on the ΣH2S concentration. At

W3 the CTR treatment reaches the same low level of ΣH2S concentration as the other treatments. This slow development could be an aftermath of the slower vertical diffusion of oxygen across the unmanipulated sediment in the CTR treatment mediating a slower expansion and less deep expansion of the oxic zone, compared to the more rapid three-dimensional diffusion occurring at higher rates in bioturbated sediments (Coull 1999) such as in the HM treatment.

Microprofiles of pH in this thesis further explain the effects on the sediment geochemistry by the presence of cable bacteria (e.g. Nielsen et al. 2010, Pfeffer et al. 2012, Meysman et al. 2015). As the cable bacteria successively expanded deeper into the sediment during the experiment they lowered the pH in all treatments as they oxidise the sulphide in the deeper part of the suboxic zone, resulting in net gain of protons (Seitaj et al. 2015). The pH and stabilisation depths fluctuate at W1 and W2, whereas at W3 the pH stabilises around 5.5 at the depth of the SAABs which are confined around 5.5 mm depth. These acidic conditions were not measured in situ in the Eastern Gotland Basin (Marzocchi et al. 2018) or in Aarhus bay (Schauer et al. 2014) in the Baltic Sea and not in the southern North Sea (Malkin et al.

2014; Seitaj et al. 2015). Neither have these low pH values been measured in laboratory studies (Pfeffer et al. 2012; Risgaard-Petersen et al. 2012).

The distinct pH peak characteristic for presence of cable bacteria (Pfeffer et al. 2012) found in the HM treatment at W1 reveals the activity and presence by cable bacteria. Cable bacteria activity was probably enhanced by the higher abundance of meiofauna rapidly oxygenating the sediment in the advantage for the cable bacteria. This might also explain the deep SAAB found at W1. At W2 and W3 in the HM treatment, the bioturbation from the meiofauna seem to homogenise the sediment so no distinct pH peak could be seen, however small pH peaks could be seen at individual microprofiles. At W3 the most distinct pH peak was found in the CTR treatment where the highest densities of cable bacteria filaments were found and the lowest abundance of meiofauna and bioturbation were found, suggesting an ideal environment.

Also, in the CTR treatment the largest suboxic zone was measured. Even though cable bacteria were assumed to have better access to oxygen by the higher abundance of bioturbating meiofauna in the HM treatment, the suboxic zone expanded 0,4 mm shorter in the HM than in the CTR treatment. It might be the lack of the suggested destructive bioturbation (Malkin et al. 2014) in the CTR treatment, that facilitated the growth of the cable bacteria filaments further down in the sediment, while in the HM treatment bioturbation might have hindered the expansion of the suboxic zone.

Microbial community structure The bacterial community structure in the four treatments was significantly different, probably due to the different abundances of meiofauna and meiofaunal bioturbation. Presence of meiofauna and their different effect on bacteria have been studied for decades and are known to increase bacterial growth (e.g. Hubas et al. 2010), increase bacterial mineralisation (e.g. Coull 1999, Bonaglia et al. 2014) and increase solute transport over the SWI from bacterial degradation (Aller and Aller 1992; Rysgaard et al. 2000). These different effects might have been triggered by the different meiofauna abundances and bioturbation in this thesis at different levels and thereby form different sediment environments generating diverse bacterial community structures. The effect of different abundances of meiofauna and bioturbation on bacterial community structures have been reported before, e.g. in Näslund, Nascimento, and Gunnarsson (2010) and Nascimento, Näslund, and Elmgren (2012). Näslund, Nascimento, and Gunnarsson (2010) saw that higher abundance of meiofauna affected the bacterial community structure significantly by grazing on bacteria. Nascimento, Näslund, and Elmgren (2012) found that the mineralisation by bacteria increased with increased abundance of meiofauna and bioturbation, but also significantly affected the bacterial community structure. The former mentioned studies have most likely disturbed the natural occurring processes by homogenising the sediment when extracting the meiofauna to control the meiofaunal abundances and presence of macrofauna. In this thesis, the sediment cores

24 were intact when adding the meiofauna gradients to the different treatments and might simulate more realistic conditions for the development of the bacterial community structure in the presence of different meiofauna abundances. Still, the bacterial community structures were significantly different by the presence of meiofauna and meiofaunal bioturbation in Näslund, Nascimento, and Gunnarsson (2010), Nascimento, Näslund, and Elmgren (2012) and in this thesis, which indicates a strong effect by the presence of meiofauna. Nevertheless, further studies are needed to understand what mechanisms conducted by meiofauna are affecting the diversity of the bacterial community structures.

Conclusions and outlook The expansion of the oxic and suboxic zone from both meiofauna and cable bacteria are important as it results in the same effect: enables recolonization of other organisms and delays the release of sulphide during oxygen depletion events.

Cable bacteria may play an important role as they are fast colonisers and thereby faster changes the sediment chemistry in sediments with recurrent oxygen depletion events. The fast colonisation of cable bacteria is probably due to a resting seed population within the sediment, ready to cultivate when the conditions are favourable (Malkin et al. 2014; Schauer et al. 2014). When cable bacteria colonise reoxygenated sulfidic sediments, they expand the suboxic layer deeper into the sediment (Nielsen et al. 2010) and prevent free sulphide to diffuse up to the sediment surface and the overlying water (Seitaj et al. 2015) that is toxic to most animals (Bagarinao 1992). The cable bacteria are also important as they create this deeper suboxic zone faster than the normal diffusion in defaunated sediments (Nielsen et al. 2010), which could enable a faster recolonization of other organisms between recurrent oxygen depletion events (Seitaj et al. 2015), such as meiofauna.

In sediments during oxygen depletion events, phosphorous is released chemically when anaerobic respiration uses iron as an oxidiser instead of oxygen and biologically when phosphorous stored within the aerobic bacteria is released (Aller 1994). When the sediments eventually get reoxygenated, the fast recolonization by the likely seed population (Malkin et al. 2014; Schauer et al. 2014) of cable bacteria or the bottom water current transported meiofauna could play an important role in decreasing the total release of benthic bound phosphorous. A study by Aller (1994) showed that oxygen depleted sediments exposed to reoxygenation events binds more phosphorous than sediments unexposed to reoxygenation events and may even store phosphorous in sediments with longer periods of oxygen depletion. Aller (1994) also showed that the deeper the suboxic zone penetrates into the sediment the better the zone could withstand recurrent oxygen depleted events, which is why the fast colonisation of meiofauna also are important.

Since areas of temporarily and permanent oxygen depletion increases (Hansson, Andersson, and Axe 2011) cable bacteria might function as a nomadic ecosystem engineer in these areas, moving from place to place, taking advantage of the energy sources and at the same time quickly transform the toxic sediment to a more suitable place for colonization of other organisms. Thereafter the meiofauna might colonise as a settler reworking the sediment. This could result in shortened time for sulphide and nutrient release in the Baltic Sea and deeper precipitation within the sediment.

The recent discovery of the cable bacteria makes the available field of research wide with a lot of fundamental questions in the need of answers. Even though meiofauna has been studied for decades in the Baltic Sea (e.g. Scheibel 1974) there are still articles requesting for more worldwide research (e.g. Schratzberger and Ingels 2018). This thesis only touches upon questions regarding the co-existence of cable bacteria and meiofauna. With the large gap in this field of science, more studies are needed to confirm results but also to answer the supplementary questions that arises.

Acknowledgements

I would like to thank my supervisor Stefano Bonaglia for excellent supervision, time spent and patience, my co-supervisor Francisco Nascimento for great supervision, Ugo Marzocchi for the opportunity to have a fun and educative visit at Aarhus University and for great feedback, Per Hedberg for giving me his day of sampling, Séréna Albert for provision of equipment and knowledge, Sven Iburg for the guidance of laboratory work, Ronny Mario Baaske for the guidance of laboratory work and patience, Nellie Stjärnkvist for the assistance on the day of sampling, Elias Broman for lending me equipment, Helena Höglander for the time spent on microscope introduction and Rachel Foster for time spent on diverse questions. I also would like to thank the DEEP laboratory staff for assistance with nutrient analyses and Francisco Nascimento for assistance with bacterial analyses and interpretation. Last but not least I would like to thank Ph. D. student Johan Wikström, M. Sc. Andrea De Cervo and Ph. D. student Nannie Persson for the support.

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Supplementary information

Supplementary information 1

Methods

Sediment core incubation for O2 consumption and methane fluxes

An incubation was performed the last week of the experiment to estimate the total O2 consumption in the cores, which correspond to the activity in the sediment. The day before, the incubation tank and cores were carefully cleaned, and water was changed with new in situ bottom water to avoid possible effects from developing biofilm and resuspended particles. Also, a test-incubation of two hours was performed to estimate the time needed to get a sufficient O2 consumption of ~20% (Bonaglia et al. -1 2014). The test incubation showed an O2 consumption of ~5 % h , which resulted in a six-hour incubation the following day. During the incubation, the sediment cores were capped without gas bubbles in the water phase and equipped with magnetic stirrers to mediate a well-mixed water column and stabilised DBL.

Measurements for O2 concentrations and water samples for methane fluxes were taken at four random places in the incubation tank at the start and in each core at the end of the incubation. For the measurements of O2 concentrations a pre-calibrated microsensor (OX-500; Unisense, Denmark) connected to a Microsensor Multimeter (Unisense, Denmark) was used. The water samples for methane fluxes were transferred to two 12 ml Exetainer vials (Labco Scientific) from each core. To stop present processes in the vials 0.1 ml of zinkcloride (ZnCl2) was added. Head space analyses of methane was done in a gas chromatograph (Shimadzu GS-8A).

Fluxes of both O2 and methane across the sediment water interface (SWI) during the incubation were calculated from the difference in start and end concentrations in the water column, accounting for the individual amount of water in each core

Results

O2 consumption

The negative fluxes show that O2 was consumed in the cores during the six-hour incubation (Figure S1).

There was no significant difference in average total O2 consumption in the water column between

33 treatments (ANOVA, p = > 0,05). The lowest O2 consumption was found in core LM1 (4.3 %) and the highest in core LM2 (18.0 %) (Figure S1A).

A CTR DEBRIS LM HM B AVERAGE 1 2 3 1 2 3 1 2 3 1 2 3 CTR DEBRIS LM HM 0 0 0 0 0

-5 -5 -5 -5 -5

-10 -10 -10 -10 -10

decrease (%) -15 -15 -15 -15 -15 2 O -20 -20 -20 -20 -20

Figure S1. Total oxygen consumption in the water column in percentage (%) for six hours. A) Total oxygen consumption in each replicate core; 1,2 and 3per treatment; CTR, DEBRIS, LM and HM. B) Average total O2 consumption per treatment; CTR, DEBRIS, LM and HM (mean ± standard error, n = 3).

Methane fluxes

The negative fluxes indicate that CH4 consumption occurred and the positive fluxes indicates that CH4 production occurred in the cores (Figure S2A). The highest production of CH4 could be found in the LM treatment in core 2 (38.1 % increase) and there was also a small production of CH4 in the LM treatment core 3 (1.5 %) and in the CTR treatment in core 1 (0.7 %). The highest consumption of CH4 was found the HM treatment in core 3 (97.7 %). Still, there was no significant difference in average flux of CH4 between treatments (ANOVA, p = > 0,05) (Figure S2B).

A CTR DEBRIS LM HM B AVERAGE 1 2 3 1 2 3 1 2 3 1 2 3 CTR DEBRIS LM HM 50 50 50 50 50

0 0 0 0 0 flux (%) flux 4 -50 -50 -50 -50 -50 CH

-100 -100 -100 -100 -100

Figure S2. Difference in concentration of CH4 in the water column in percentage (%) compared to the concentration at the start of the six-hour incubation. A) Difference in concentration of CH4 in each replicate core; 1,2 and 3 per treatment; CTR, DEBRIS, LM and HM. B) Average difference in concentration of CH4 per treatment; CTR, DEBRIS, LM and HM (mean ± standard error, n = 3).

Bacterial community structure In the core LM2 there was a peak of bacterial diversity with over 2500 different species, still the similarity of the diversity in bacterial composition is the highest in the LM treatment.

Figure S3. Non-metric multidimensional scaling showing the bacterial composition in the four treatments; CTR (red), DERBIS (green), LM (turquoise) and HM (purple). The distance between the treatment groups in the figure show the difference in bacterial composition. No overlap means no similarity. The distance between the dots within each treatment group show the difference in bacterial composition. Small spaces between the dots mean high similarity.

Discussion Bacterial peak Possible fish bones (gill cover and spikes from a Stickleback, genus Gasterosteus) were observed during meiofauna counting and pictures were sent to the Institute of Coastal Research at Swedish University of Agricultural Sciences for identification (still no confirmation at the time of writing). This potential explanation could be supported by a peak in total O2 consumption (18 %) (Figure S1), probably caused by higher abundance from the increased diversity of bacteria. With the imaginable intense degradation and oxygen consumption within the sediment by the presence of the fish carcases, it is possible that the oxic and suboxic zones are retracted closer to the sediment surface together with the underlying methanogenesis resulting in methane release (51,1 ± 1,4 µmol L-1, n = 2) (Figure S2).

Supplementary information 2

Results Meiofauna abundances In the second centimetre of sediment (1 – 2 cm) nematodes were still the most abundant phyla while no Ostracods were found at all.

Table S1. Meiofauna abundance in individuals per square meter (ind. 103 m2) in the second centimetre (1 – 2 cm) of sediment in each treatment; CTR, DEBRIS, LM and HM. Values are mean ± standard deviation (n = 3). * Most of the organisms were identified to the genus Bosmina. CTR DEBRIS LM HM Meiofauna (ind. 103 m-2) (ind. 103 m-2) (ind. 103 m-2) (ind. 103 m-2) Nematods 5.8 ± 2.8 16.3 ± 16.1 18.6 ± 8.5 84.6 ± 28.1 Ostracods 0 0 0 0 Copepods 1.2 ± 1.5 0.2 ± 0.4 0 0.9 ± 1.6 Kinorhyncha 0 0 0.9 ± 1.6 1.9 ± 1.6 Cladocerans* 0.2 ± 0.4 0.2 ± 0.4 3.3 ± 4.5 9.3 ± 11.6 Halacaridae 0.5 ± 0.8 0.2 ± 0.4 0 0 Total 7.7 ± 5.3 17.0 ± 17.3 22.8 ± 6.9 96.7 ± 42.9

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