Document [original]

Distribution of Dehalococcoidia in marine sediments and

strategies for their enrichment

vorgelegt von

Master of Science

Camelia Algora

geb. in Madrid, Spanien

von der Fakultät III – Prozesswissenschaften

der Technischen Universität Berlin

zur Erlangung des akademischen Grades

Doktor der Naturwissenschaften / Doctor rerum naturalium

-Dr. rer. nat.-

genehmigte Dissertation

Promotionsausschuss:

Vorsitzender: Prof. Dr. Juri Rappsilber

Gutachter: Prof. Dr. Peter Neubauer

Gutachter: PD Dr. Lorenz Adrian

Gutachter: Prof. Dr. Ricardo Amils

Tag der wissenschaftlichen Aussprache: 11. Juli 2016

Berlin 2016

III

“Life on earth is such a good story you cannot afford to miss the beginning.

Beneath our superficial differences we are all of us walking communities of bacteria.

The world shimmers a pointillist landscape made of tiny living beings”

Lynn Margulis

Dedicado a mis tíos,

Dr. José Luis Gallardo y Dra. María del Carmen López,

por plantar la semilla del interés por la Ciencia en mi adolescencia,

por su ejemplo y por todo su apoyo

DECLARATION OF INDEPENDENT WORK

Erklärung zur Dissertation

Ehrenwörtliche Erklärung zu meiner Dissertation mit dem Titel:

‘Distribution of Dehalococcoidia in marine sediments and strategies for their

enrichment’

hiermit erkläre ich, dass ich die beigefügte Dissertation selbstständig verfasst und keine

anderen als die angegebenen Hilfsmittel genutzt habe. Alle wörtlich oder inhaltlich

übernommenen Stellen habe ich als solche gekennzeichnet. Wörtlich oder inhaltlich Stellen,

die aus meinen persönlichen Publikationen stammen, wurden nicht zusätzlich gekennzeichnet.

Ich versichere, dass ich die beigefügte Dissertation nur in diesem und keinem anderen

Promotionsverfahren eingereicht habe und dass diesem Promotionsverfahren keine endgültig

gescheiterten Promotionsverfahren vorausgegangen sind.

_____________________ ______________________________

Ort, Datum Unterschrift

ACKNOWLEDGEMENTS

The present work was carried out within the research group of PD Dr. Lorenz Adrian at the

Department of Isotope Biogeochemistry (ISOBIO) at the Helmholtz Centre for Environmental

Research–UFZ in Leipzig, Germany, from the beginning of November 2008 to end of

February 2012, except for a period of 10 weeks on board the Polarstern, during the expedition

ARKXXV/3. The PhD work was done under the research project MICROFLEX (Proposal Nr.

202903), which was funded by the European Research Council.

In the first instance, I would like to deeply thank my supervisor, Dr. Lorenz Adrian, for giving

me the great opportunity to accomplish this PhD project, for supporting me throughout its

development, for innumerable ideas, and contagious enthusiasm. I thank him for sharing his

knowledge with numerous BBCs, inspiring discussions, and for providing excellent advice on

publications and this thesis.

I would like to thank Prof. Peter Neubauer for external supervision and review of my thesis

work, and Prof. Ricardo Amils for reviewing my dissertation work and for being a deeply

inspirational lecturer during my time at UAM (Universidad Autónoma de Madrid). I thank

Prof. Dr. Juri Rappsilber for being the head of the dissertation committee.

I am honoured to have worked with Dr. Kenneth Wasmund, whom I profoundly thank for

immense support, brilliant ideas, fabulous scientific discussions, and great friendship. This

work could not have been done without his help.

I am immensely grateful to Dr. Martin Krüger, for the opportunity to join the research

expedition on the Polarstern and for fruitful collaboration. Thanks to Friederike Gründger for

sharing field work and time on board the Polarstern and further collaboration in the BGR.

Thanks to all the colleagues at the expedition ARK XXV/3 and especially to Dr. Volkmar

Damm and Dr Thomas Pletsch.

I would like to thank Dr. Timothy G. Ferdelman and Dr. Casey Hubert, from the Max Planck

Institute for Marine Microbiology in Bremen, for kindly suppling sediment samples.

It has been an immense pleasure to me to collaborate with Dr. Sotirios Vasileiadis and Dr.

Edoardo Puglisi. I especially would like to thank them for their dedication and a fantastic

working atmosphere.

My most sincere gratitude goes to Dr. Paula Martínez, Dr. Petra Bombach, Dr. Anke Wagner,

and Dr. Kelly E. Fletcher, for their great scientific advice, suggestions, enthusiasm, support,

and friendship.

VII

It has been a pleasure to me to work with an international lab group, full of lovely and talented

microbiologists from all over the world. I especially want to thank Dr. Ernest Marco-Urrea,

Anja Kublik, Josefine Müller, Dr. Chang Ding, Katja Seidel, and Chao Yang, among many

others. I particularly would like to thank Myriel Cooper for helpful discussions and splendid

collaboration within our MICROFLEX project, and for her friendship. Special thanks go to

Christina Lachmann, for fabulous help with the first pioneering months setting up the lab and

the first cultures and experiments. Many thanks go to our technician, Benjamin Scheer, for his

support from the beginning until the end of my PhD project.

My very special thanks go to my students: Christina M. Ridley, Vaida Vaitkeviciute,

Sebastian Röther, and Sukwon Jang. I thank them for their hard work and dedication, their

eagerness to learn and enthusiasm.

I would like to thank all my colleagues at the ISOBIO department, and specially, I would like

to thank Dr. Ivonne Nijenhuis, our department leader Dr. Hans-Hermann Richnow, Dr. Sara

Herrero, Dr. Mònica Rosell, Dr. Felipe Bastida, Dr. Maria Luisa Feo, Marie Markantonis,

Kristina Hitzfeld, Jan Birkigt, and Julian Renpenning among many others.

I am honoured to have been part of the HIGRADE graduate school of the UFZ, and I would

like to thank Vera Bissinger and Barbara Timmel, among the many people who run it.

I very much would like to thank all the friends that have enriched my life throughout these

years in the UFZ, their names are too many to be written, but their support, love and

friendship was essential.

Mi mayor agradecimiento se lo dedico a mi familia, por su inmenso apoyo, comprensión y

amor, sin el cual hubiera sido totalmente imposible alcanzar la consecución de este trabajo de

tesis doctoral.

VIII

ABSTRACT

The marine subsurface is one of the largest microbial habitats on Earth. Microbial community

studies based on the 16S rRNA gene have revealed that bacteria of the class Dehalococcoidia,

phylum Chloroflexi, are one of the most widespread and abundant bacteria inhabiting the

marine subsurface. However, their physiology and ecological role are unknown. In this study,

marine sediments were cultivated with various potential electron acceptors to explore

respiration modes catalysed by Dehalococcoidia. The cultivation revealed that

Dehalococcoidia can be cultivated in an anoxic minimal medium amended with hydrogen and

acetate as potential electron donor and carbon source, respectively, at a temperature of 30ºC

and at atmospheric pressure, and thus Dehalococcoidia are not strict piezophilic bacteria. An

increase in Dehalococcoidia 16S rRNA gene copy numbers of one order of magnitude, as

evidenced by quantitative PCR with newly designed primers, could be observed in a time

span of months for several sediment cultures, indicating that their replication times are not in

geological times, i.e., thousands to millions of years. Dehalococcoidia 16S rRNA gene copy

numbers (in the range of 102–105 ml-1 culture) among the various sediment cultures inoculated

with diverse sediments and amended with different potential electron acceptors suggested no

specific respiratory mode to be preferred by subseafloor Dehalococcoidia. Additionally,

subseafloor Dehalococcoidia were not reliant on organohalide respiration under the

conditions tested, in contrast to cultivated Dehalococcoidia species such as Dehalococcoides

mccartyi. Interestingly, among the tested organohalide compounds, 1,2,3-trichlorobenzene

was transformed to 1,3-dichlorobenzene in sediment cultures. Inhibition studies using

antibiotics against Gram-positives and microbial community analyses by 454-pyrosequencing

of 16S rRNA genes, indicated members affiliated with the phylum Firmicutes were involved

in the transformation of 1,2,3-trichlorobenzene.

The presence, abundance, and distribution of Dehalococcoidia were investigated in sediments

of the Arctic Baffin Bay. Geochemical studies showed the shelf as an area rich in organic

matter, with indications for the presence of sulphate reduction (highest dsrA gene copies and a

decrease in sulphate concentration in its pore-waters). On the other hand, the basin area was

comparably poor in organic matter with high concentrations of iron(II) and manganese(II) in

its pore-waters. Dehalococcoidia were present at all investigated sites and depths in the range

of 1.1 x 103–8.4 x 105 16S rRNA gene copy numbers g-1 sediment. Dehalococcoidia copy

numbers were highest and generally stable with depth in shelf sites of the Baffin Bay

compared to any other area. In contrast, Dehalococcoidia copy numbers pronouncedly

decreased with depth at basin sites. Dehalococcoidia accounted for the highest proportion of

the total bacterial 16S rRNA genes when low bacterial copy numbers were found, mostly in

samples from deeper sediment layers, indicating that subseafloor Dehalococcoidia are

resilient to burial. Illumina sequencing of 16S rRNA genes gave further evidence that

Dehalococcoidia were mostly associated with shelf sediments. The relative abundance of

Dehalococcoidia and, most specifically, the clade GIF-9 within the class Dehalococcoidia,

positively correlated with organic matter content, and negatively correlated with sulphate and

manganese(II) pore-water concentrations. Thus, a potential fermenting metabolism is likely

for GIF-9 members. Apart from Dehalococcoidia, the Baffin Bay bacterial community was

dominated by members of the class Betaproteobacteria (with relative abundance of 38 to

64%), and specifically the order Burkholderiales, which strongly correlated to manganese(II)

pore-water concentration, suggesting a metal respiratory metabolism. In contrast to

Dehalococcoidia, the class Betaproteobacteria is not commonly found as widely distributed

and abundant bacterial group in the marine subseafloor, and its presence and high abundance

in sediments of the Baffin Bay may be the result of the glacial conditions in the area.

ZUSAMMENFASSUNG

Marine Sedimente sind einer der größten mikrobiellen Lebensräume der Erde. 16S rRNA-

basierte Studien der mikrobiellen Gemeinschaften haben gezeigt, dass Bakterien der Klasse

Dehalococcoidia des Phylums Chloroflexi ubiquitär und abundant in marinen Sedimenten

vorkommen. Die Physiologie und ökologische Funktion dieser Bakterien ist weitestgehend

unbekannt. In der vorliegenden Arbeit wurden marine Sedimente mit verschiedenen

potenziellen Elektronenakzeptoren versetzt, um die Stoffwechselcharakteristika der Klasse

Dehalococcoidia zu identifizieren. Die Kultivierungsexperimente zeigten, dass

Dehalococcoidia-Spezies in anoxischem Minimalmedium versetzt mit Wasserstoff und Acetat

als Elektronendonor und Kohlenstoffstoffquelle bei 30°C und atmosphärischem Druck

erfolgreich kultiviert werden können und dementsprechend nicht strikt barophil sind. Zur

Quantifizierung von Dehalococcoidia-Spezies wurde ein qPCR-Verfahren entwickelt.

Vereinzelt wurde in den Labormikrokosmenansätzen ein Anstieg der Dehalococcoidia-

spezifischen 16S rRNA-Genkopienzahl innerhalb der mehrmonatigen Inkubation verzeichnet.

Signifikante Änderungen der Dehalococcoidia-spezifischen 16S rRNA-Genkopienzahl in

Abhängigkeit von den verschiedenen Elektronenakzeptoren wurden nicht beobachtet und

ließen damit keine Rückschlüsse auf spezifische Stoffwechselprozesse zu. Weiterhin konnte

für marine Dehalococcoidia-Spezies im Gegensatz zu kultivierten Süßwasserspezies wie z.B.

Dehalococcoides mccartyi keine Abhängigkeit von Organohalid-Respiration nachgewiesen

werden. Interessanterweise wurde als einzige der getesten chlorierten Kohlenwasserstoffe

1,2,3-Trichlorbenzol zu 1,3-Dichlorbenzol umgesetzt. Studien zur Inhibierung von gram-

positiven Bakterien mit Antibiotika sowie 16S rRNA-basierte Analysen der mikrobiellen

Gemeinschaften mittels Pyrosequenzierung deuteten auf die Relevanz von Firmicutes-

verwandten Spezies zur Transformation von 1,2,3-Trichlorbenzol.

Das Vorkommen, die Abundanz und räumliche Verteilung von Dehalococcoidia wurde in

Sedimenten der nördlichen Baffin Bay untersucht. Geochemische Studien zeigen, dass große

Mengen an organischer Substanz im Schelf festgelegt sind. Hohe Kopienzahlen der

dissimilatorischen Sulfitreduktase (dsrA) verbunden mit einer Sulfatzehrung im Porenwasser

deuten auf die Relevanz von Sulfatreduktion hin. Das Baffinbecken ist dagegen

vergleichsweise arm an organischer Substanz mit hohen Eisen(II)- und Mn(II)-

Konzentrationen im Porenwasser. Das Markergen für Dehalococcoidia wurde in

Sedimentproben verschiedener Transekte und vertikaler Zonierungen mit einer Abundanz von

103 bis 105/g Sediment nachgewiesen. In den Schelfsedimenten trat das Dehalococcoidia-Gen

mit einer Abundanz von 105/g Sediment auf und zeigte keine signifikante Änderung entlang

der vertikalen Zonierung. Im Gegensatz dazu nahmen die Kopienzahlen des Markergens in

den Sedimentproben des Baffinbeckens mit zunehmender Tiefe rasch ab. Lagen bakterielle

Gene in geringen Kopienzahlen vor, was zumeist in den tieferen Sedimentschichten

beobachtet wurde, machten Dehalococcoidia-Gene den Großteil der bakteriellen 16S rRNA-

Gene aus. Illuminasequenzierung der 16S rRNA-Gene zeigte darüber hinaus, dass

Dehalococcoidia vorrangig in Schelfsedimenten vorkamen. Die relative Abundanz von

Dehalococcoidia, insbesondere der Gruppe GIF-9 innerhalb der Dehalococcoidia, korrelierte

positiv mit dem Gehalt an organischer Substanz und negativ mit der Sulfat- und Mangan(II)-

Konzentrationen des Porenwassers. Dementsprechend ist ein fermentativer Stoffwechsel für

Spezies der Gruppe GIF-9 in Betracht zu ziehen. Diese Hypothese wird durch die Ergebnisse

der Kultivierungsexperimente gestützt. Neben Dehalococcoidia wurde die mikrobielle Baffin-

Bay-Gemeinschaft von Spezies der Klasse Betaproteobacteria (mit einer relativen Abundanz

von 38 bis 64%), insbesondere der Ordnung Burkholderiales, dominiert. Das Vorkommen von

Burkholderiales-Spezies zeigte eine starke Korrelation mit den Eisen(II)- und Mangan(II)-

Konzentrationen und deutet auf eine mikrobielle Eisen- und Manganreduktion hin. Im

Gegensatz zur Klasse Dehalococcoidia kommen Betaproteobacteria nur vereinzelt in marinen

Sedimenten vor. Deren Vorkommen und hohe Abundanz in den Baffin Bay-Sedimenten

resultieren möglicherweise aus den eiszeitlichen Bedingungen.

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LIST OF PUBLICATIONS

Results from this thesis led to three peer-reviewed publications in scientific journals and to a

contribution to one scientific cruise report. This thesis is based on those original articles and

on unpublished data.

 PUBLICATION I

Algora C., Gründger F., Adrian L., Damm V., Richnow H.-H., Krüger M. (2013).

Geochemistry and microbial populations in sediments of the Northern Baffin Bay,

Arctic. Geomicrobiology Journal 30 (8), 690 – 705;

DOI:10.1080/01490451.2012.758195

Author contributions

Martin Krüger developed the concept and coordinated this study. The manuscript

concept was developed by Camelia Algora, Lorenz Adrian and Martin Krüger.

Volkmar Damm made possible and coordinated the ARK XXV/3 expedition and

supplied geological data. Hans-Hermann Richnow supplied ideas for the study

concept, field sampling and analysis of isotope data. Friederike Gründger and Camelia

Algora designed and performed the field work on board of the Polarstern and

produced the data in the laboratory after the expedition. Additionally, Friederike

Gründger and Martin Krüger supplied extra data on microcosms and coordinated the

geological analysis of sediment samples at BGR. Camelia Algora interpreted the

results and drafted the manuscript. Lorenz Adrian and Martin Krüger revised the

manuscript.

 PUBLICATION II

Wasmund K., Algora C., Müller J., Krüger M., Lloyd K. G., Reinhardt R., Adrian L.

(2015). Development and application of primers for the class Dehalococcoidia

(phylum Chloroflexi) enables deep insights into diversity and stratification of

subgroup in the marine subsurface. Environmental Microbiology.17:3540-3556;

DOI:10.1111/1462-2920.12510

Author contributions

Kenneth Wasmund and Lorenz Adrian developed the concept of this study. Kenneth

Wasmund designed the primers and performed alignments with several bioinformatics

software. The practical work in the laboratory was conducted by Kenneth Wasmund,

Camelia Algora and Josefine Müller. Karen G. Lloyd, Martin Krüger and Camelia

Algora designed, coordinated, and performed field sampling of sediments, as well as

XIII

the biogeochemical analysis of sediments. Richard Reinhardt performed the

pyrosequencing of the samples. Kenneth Wasmund performed the sequence

bioinformatics analysis, Unifrac analysis and produced the phylogenetic trees.

Kenneth Wasmund interpreted the results and drafted the manuscript. Lorenz Adrian

and Camelia Algora revised the manuscript.

 PUBLICATION III

Algora C., Vasileiadis S, Wasmund K, Trevisan M, Krüger M, Puglisi E, Adrian L.

(2015). Manganese and iron as structuring parameters of microbial communities in

subsurface sediments of the Arctic Baffin Bay. FEMS Microbiology Ecology.

91:fiv056; DOI: 10.1093/femsec/fiv056

Author contributions

Camelia Algora, Lorenz Adrian, and Kenneth Wasmund developed the concept of this

study, designed the laboratory experiments and coordinated the study. Sotirios

Vasileiadis, Marco Trevisan, and Edoardo Puglisi performed the Illumina sequencing.

Sotirios Vasileiadis performed the sequence bioinformatics analysis and the statistical

analysis. Camelia Algora and Martin Krüger supplied samples and designed the field

sampling. Camelia Algora performed the field sampling. The laboratory work was

conducted by Camelia Algora and Kenneth Wasmund. Camelia Algora was the main

responsible of the data interpretation. Camelia Algora drafted the manuscript. Lorenz

Adrian and Kenneth Wasmund revised the manuscript and supplied ideas.

 CRUISE REPORT

Gründger F. and Algora C. (2010), Chapter 13, Biogeochemistry and

Geomicrobiology, p. 136-139. In Damm V. (Editor), 2010. Cruise Report: The

expedition of the research vessel “Polarstern” to the Arctic in 2010 (ARK-XXV/3).

Reports on Polar- and Marine Research

ISSN 1866–3192; http://hdl.handle.net/10013/epic.36297.d001

Author contributions

Camelia Algora and Friederike Gründger designed and performed the field and

laboratory practical work on board of the Polarstern. The whole concept of the study

was designed by Martin Krüger together with Hans Hermann Richnow and Lorenz

Adrian. Camelia Algora wrote the manuscript, Friederike Gründger revised it.

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ABBREVIATIONS

bp base pairs

BGR Bundesanstalt für Geowissenschaften und Rohstoffe

BSA bovine serum albumin

cmbsf centimetres below surface

DCB dichlorobenzene

DNA deoxyribonucleic acid

GC-FID gas chromatography associated to a flame ionization detector

ICP-MS inductively coupled plasma mass spectrometry

Ma Megaannum, one million years

NW North-West

mbsf meters below surface

OTU operational taxonomic unit

PCR polymerase chain reaction

PCE tetrachloroethene

PVC polyvinyl chloride, synthetic plastic

qPCR quantitative PCR

RDase reductive dehalogenase

rdhA reductive dehalogenase-homologous gene

RNA ribonucleic acid

rpm rounds per minute

rRNA ribosomal ribonucleic acid

spp species

SW South-West

SD Standard Deviation

TCB trichlorobenzene

TCE trichloroethene

TOC total organic carbon

TC total carbon

UFZ Helmholtz-Zentrum für Umweltforschung

v/v volume per volume

VPDB Vienna PeeDee Belemnite standard

LIST OF CONTENTS

1 Introduction ......................................................................................................... 1

1.1 Marine sediments and the biogeochemical processes within ................................ 1

1.2 Microbial life in marine sediments ....................................................................... 3

1.3 Abundance and distribution of Chloroflexi in marine sediments .......................... 5

1.4 Organohalide respiration in marine sediments .................................................... 10

1.5 Study area: Baffin Bay ........................................................................................ 11

1.6 Aim of the study .................................................................................................. 15

2 Material & Methods .......................................................................................... 17

2.1 Chemicals ............................................................................................................ 17

2.2 Sediment samples ................................................................................................ 17

2.2.1 Core sampling ..................................................................................................... 20

2.2.2 Core subsampling ............................................................................................... 21

2.3 Sediment geochemical analysis........................................................................... 25

2.4 Cultivation of microorganisms in sediment cultures .......................................... 26

2.5 Isolation of pure strains by cultivation in deep-agarose dilution tubes ............... 29

2.5.1 Media, solutions and preparation procedure...................................................... 29

2.5.2 Picking and transferring of colonies ................................................................... 30

2.6 Analytical methods.............................................................................................. 31

2.6.1 Analyses of trichlorobenzenes by gas chromatography ...................................... 31

2.6.2 Analyses of chlorophenols by gas chromatography............................................ 31

2.6.3 Cell visualization by epifluorescence microscopy .............................................. 32

2.7 Molecular biology methods................................................................................. 32

2.7.1 DNA isolation from sediments, cultures and colonies ........................................ 32

2.7.2 Determination of DNA concentration ................................................................. 33

2.7.3 Quantification of the 16S rRNA gene by qPCR .................................................. 34

2.7.4 Quantification of the functional genes mcrA and dsrA by qPCR ........................ 38

2.7.5 Amplification of reductive dehalogenase genes .................................................. 39

2.7.6 Amplification of 16S rRNA gene from colonies .................................................. 40

2.7.7 Amplification of 16S rRNA gene for 454-pyrosequencing .................................. 40

2.7.8 Amplification of the 16S rRNA gene for Illumina sequencing ............................ 41

2.7.9 Agarose gel electrophoresis and amplicon purification ..................................... 42

2.7.10 Elution of amplicons from agarose gels.............................................................. 42

2.7.11 Cloning of 16S rRNA gene amplicons from Dehalococcoidia qPCR

amplifications ...................................................................................................... 42

2.7.12 454-pyrosequencing, Illumina sequencing and subsequent data analysis .......... 43

2.7.13 Sanger sequencing and subsequent data analysis .............................................. 44

3 Results ................................................................................................................ 46

3.1 Approaches to cultivate Dehalococcoidia from various marine sediments ........ 46

3.1.1 Dehalococcoidia abundance under various terminal electron acceptors

in sediment cultures ............................................................................................. 46

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3.1.2 Total Bacteria abundance under various terminal electron acceptors in

sediment cultures ................................................................................................. 54

3.1.3 Colony formation in deep-agarose dilution tubes under various

terminal electron acceptors................................................................................. 55

3.1.4 Colony identification after 16S rRNA gene sequencing ...................................... 57

3.2 Organohalide transformation in marine sediment cultures ................................. 60

3.2.1 Transformation of 1,2,3-TCB to 1,3-DCB in marine sediment cultures ............. 60

3.2.2 Enrichment of trichlorobenzene dechlorinating bacteria from sediment

cultures ................................................................................................................ 64

3.2.3 Quantification of total Bacteria and known organohalide-respiring

bacteria by qPCR ................................................................................................ 70

3.2.4 Bacterial community changes in enrichment cultures studied by 454-

pyrosequencing ................................................................................................... 73

3.2.5 Microbial community study with universal primers in the enrichment

cultures using 454-pyrosequencing..................................................................... 81

3.3 Baffin Bay geochemistry and microbial communities ........................................ 84

3.3.1 Geochemistry of Baffin Bay cores ....................................................................... 85

3.3.2 Microbial ecology of Baffin Bay cores................................................................ 93

4 Discussion ......................................................................................................... 112

4.1 In situ abundance of Dehalococcoidia .............................................................. 113

4.2 Electron acceptors in the Baffin Bay ................................................................ 117

4.2.1 Sulphate reduction and methanogenesis in shelf sediments ............................. 118

4.2.2 Importance of metal biogeochemistry in the central Baffin Bay ....................... 119

4.3 Burkholderiales dominate in Baffin Bay sediments ......................................... 121

4.4 Dehalococcoidia cultivation ............................................................................. 123

4.5 Dehalogenation of organohalides in sediment cultures .................................... 126

4.6 Firmicutes are common cultured bacteria from marine sediments ................... 130

5 Conclusion ........................................................................................................ 132

6 Appendix .......................................................................................................... 135

6.1 Appendix 1 ........................................................................................................ 135

6.1.1 Mineral solution “Widdel solution” ................................................................. 135

6.1.2 Trace metal solution “SL-9 solution” ............................................................... 135

6.1.3 Bicarbonate solution ......................................................................................... 135

6.1.4 Vitamin 7 solution ............................................................................................. 136

6.1.5 Titanium (III) citrate solution as reducing agent .............................................. 136

6.1.6 Iron sulphide solution as reducing agent .......................................................... 136

6.1.7 Medium composition and concentration ........................................................... 137

6.2 Appendix 2 ........................................................................................................ 138

6.3 Appendix 3 ........................................................................................................ 143

7 References ........................................................................................................ 144

Introduction

1 INTRODUCTION

Microorganisms are almost ubiquitously distributed throughout our planet, independently of

how high or low the temperature or the pressure or the osmotic pressure or the pH is. Deep

marine sediments are a vast underexplored environment on Earth, where we are currently

starting to get an idea how and to which extend life is thriving there. In the last decades,

scientific expeditions have drilled into the marine subsurface and tried to give some answers to

those questions. We now know that microorganisms are present and active down to depths of

2,458 meters below seafloor (mbsf) (Inagaki et al 2015). Furthermore, microorganisms are

present in marine sediments sometimes in massive amounts, which range from 106 cells per

cm3 at depths of 1,000 mbsf to 109 cells per cm3 at surface sediments (Parkes et al 1994,

Parkes et al 2000, Parkes et al 2014). We also know that these marine sediment

microorganisms belong to diverse microbial groups. Nevertheless, many more scientific efforts

are needed to unravel microbial life underneath the seafloor, such as: i) which metabolisms do

the various microbes have, i.e., what are their carbon and energy sources?; How do they

conserve energy?; Which compounds do they use as terminal electron acceptors in their

respiratory chain?; Are all of them actively thriving or in stationary phase?, and/or: ii) what

geochemical parameters from the sediments affect microbial diversity?; Does microbial

diversity change with depth and within geochemical gradients? Studying life in the subseafloor

enlarges our understanding of the concept of life, especially in energy terms, since very little

nutrients are present in deep marine sediments, where light and phototrophic production is

absent. The present study is a contribution to provide more knowledge on microbial life in

deep marine sediments, with focus on a microbial group that is one of the most abundant and

widespread in marine sediments: the class Dehalococcoidia within the phylum Chloroflexi.

1.1 MARINE SEDIMENTS AND THE BIOGEOCHEMICAL PROCESSES WITHIN

Marine sediments are deposits of eroded rocks and soil particles transported by rivers, currents

and the wind together with material produced in oceanic processes, i.e., submarine volcano

products, seawater precipitates, products from or/and remains of marine organisms that deposit

at the seafloor and accumulate over years (Schulz and Zabel 2006). Marine sediments occupy

the surface underneath the oceans, which is close to 70% of the Earth’s surface and with depths

that vary depending on the location and that can reach 10 km (Parkes et al 2000, Parkes et al

2014). This is a vast volume of our planet, so any process happening in the marine sediments

may have critical consequences globally.

Marine sediments are naturally heterogeneous in their conditions, including: variable

temperatures that range from 4°C at the seafloor to 100–150°C in areas affected by

thermogenic processes, variable pressures that can reach up to 1,100 bar depending on the

Introduction

depth, variable mineral composition and organic matter content (Jørgensen and Boetius 2007).

Many of these conditions influence others, for example i) sediment porosity is influenced by

pressure at variable sediment depths, and ii) the quantity and presence of oxidants or electron

acceptors are influenced by the mineral composition of the marine sediments. In this line,

organic matter content depends on the primary productivity within the overlying water column

(which in turn depends on the proximity to the coast for input of fertilizing minerals and sea-

ice free period in Polar Regions), water depth, sediment depth, distance to land, and latitude

(Biddle et al 2006, Franco et al 2007, Hamdan et al 2013, Jørgensen et al 2012, Lipp et al

2008). Thus, there are oligotrophic areas such as the South Pacific Gyre, where the sediment

cover is thin (Jørgensen and Boetius 2007, Parkes et al 2014), and organic-rich areas such as

coastal sediments at continental margins (Parkes et al 2014).

It has been estimated that 5 to 10 billion tons of organic matter is present in oceanic waters,

sinking to the sediments continuously (Jørgensen 1982). The bulk of this organic matter is

oxidized during sedimentation in the water column and within the first cm of the marine

sediments. The rest of it (about 15,000 x 1018 g C; (Hedges and Keil 1995)) accumulates in the

sediment, representing the major reservoir of organic carbon on the planet. Organic matter is

oxidized with oxygen via aerobic respiration of marine biota, which are chiefly

microorganisms. In marine sediments, oxygen is consumed usually within the first near-surface

mm, sometimes within the first metre depending on the sediment type, i.e., productivity in

overlying water column (Cai and Sayles 1996, Jørgensen and Boetius 2007, Wenzhöfer and

Glud 2002). Once oxygen is depleted, organic matter is oxidised by other oxidants via

anaerobic respiration. These anaerobic oxidants are subsequently used by microorganisms in a

higher to lower standard free energy-yields order as follows: nitrate, manganese(VI), iron(III)

minerals, sulphate, and bicarbonate (Froelich et al 1979). All these reactions establish a

zonation in sediments (D'Hondt et al 2004, Froelich et al 1979, Nealson 1997) and are

enzymatically mediated by microorganisms.

Evidence of microbial processes in deep sediments of hundreds of mbsf was provided by pore-

water analysis of oxidant (i.e., oxygen, nitrate, sulphate, etc.) concentrations and of released

dissolved products such as sulphide and methane in depth profiles (D'Hondt et al 2002,

D'Hondt et al 2004). Sulphate reduction and methanogenesis have been intensively studied in

marine sediments worldwide (D'Hondt et al 2002). Sulphate reduction is one of the main

processes of organic matter degradation in marine sediments due to the high content of

sulphate ions in seawater (about 27 mM in average), which diffuses to the marine sediments

(D'Hondt et al 2002, Froelich et al 1979). Once sulphate is depleted in the sediment, the main

known microbial process in marine sediments is methanogenesis (D'Hondt et al 2002).

Although sulphate reduction is usually the dominant pathway for the anaerobic oxidation of

Introduction

organic carbon in marine sediments worldwide, other anaerobic pathways may play significant

roles in the oxidation of organic carbon in specific marine sediments. For example, in Arctic

near-surface sediments with high amounts of iron and manganese oxides, iron and manganese

reduction are the dominant processes of organic carbon oxidation accounting between 50% and

90% of anaerobic oxidation of organic carbon (Nickel et al 2008, Vandieken et al 2006,

Vandieken et al 2014). Iron reduction accounted for 25% of the total annual organic carbon

oxidation in Arctic sediments of Northeast Greenland, where nitrate reduction accounted for

4%, manganese reduction below 1%, sulphate reduction for 33%, and oxygen respiration for

38% (Rysgaard et al 1998). In this way, the microbial metabolic activity within the marine

sediments contributes substantially to Earth’s biogeochemical cycles, especially over

geological timescales (D'Hondt et al 2002, D'Hondt et al 2004, Wellsbury et al 2002).

1.2 MICROBIAL LIFE IN MARINE SEDIMENTS

Although deep marine sediments were for a long time considered without life due to the

absence of energy supply, the high hydrostatic pressure and low temperatures, the Danish

Galathaea deep-sea expedition already in 1951 showed that deep marine sediments harboured

microbial life. Abundance of 104 to 106 cells per ml of viable bacteria in marine sediments of

the Philippine Trench exceeding 10,000 m water depth was observed by microscopy and

cultivation (Morita and Zobell 1955, Zobell and Morita 1959). Later on, detailed microbial

investigations under the Ocean Drilling Program identified active metabolic processes and the

presence of microorganisms in every single drilled location (Jørgensen and Boetius 2007,

Parkes et al 2000, Parkes and Sass 2009). This gave further evidence for the presence of

microorganisms and microbially mediated processes in deep marine sediments. Since then,

various studies have demonstrated the existence of prokaryotes in all investigated deep marine

sediments. Evidence for the presence of living bacterial cells was shown in terms of the

presence of intact cells, visualized with fluorescent microscopy (Parkes et al 1994), or intact

cell molecular constituents, such as membrane lipids, DNA and RNA, isolated with molecular

biology techniques (Biddle et al 2006, Krüger et al 2002, Lipp et al 2008, Schippers et al

2005). Microscopic cell-counting after staining with the fluorescent DNA stain acridine orange

demonstrated that deep marine sediments (even deeper than 500 mbsf) harbour significant

numbers of microbial cells, with maximum cell numbers in surface sediments of around

109 cells cm-3, which decreased logarithmically to 107 at 518 mbsf in Pacific Ocean sediments

(Parkes et al 1994). Quantitative-PCR (qPCR) studies corroborated the microscopic cell

counting results, showing highest 16S rRNA gene copy numbers of 108 cells of prokaryotes

cm-3 in near-surface sediments on the Peru continental margin and numbers decreasing to

106 gene copies cm-3 by 40 mbsf (Schippers and Neretin 2006). More importantly, catalysed

reporter deposition-fluorescence in situ hybridization (CARD-FISH) targeting ribosomal RNA

Introduction

(rRNA) within the cells revealed that high numbers (up to 107 cells of prokaryotes cm-3) of

deeply buried (>400 mbsf) bacterial cells are alive as they do have ribosomes (Schippers et al

2005).

DNA-targeted molecular biological techniques allowed phylogenetic studies by partial

sequencing of the 16S rRNA genes after generation of clone libraries that identified the most

abundant microbial groups (Durbin and Teske 2011, Inagaki et al 2003, Inagaki et al 2006,

Kormas et al 2003, Reed et al 2002, Webster et al 2006). Microorganisms inhabiting marine

sediments predominantly belong to the phyla Proteobacteria, Chloroflexi, and candidate

phylum JS1 within the Bacteria, and the crenarchaeotal groups of Miscellaneous

Crenarchaeotal Group (MCG) and Marine Bentic Group B (MBG-B) within the Archaea (Fry

et al 2008, Parkes et al 2014). Many of these microbial phyla, such as the candidate phylum

JS1 (Webster et al 2004), have no isolated members, and others, such as the phylum

Chloroflexi, have isolated members, however very distantly related to the sequences retrieved

from marine sediments.

Cultivation efforts of marine sediment bacteria have yielded isolates belonging to the phyla

Proteobacteria, Firmicutes, Actinobacteria and Bacteroidetes (D'Hondt et al 2004). Firmicutes

and Actinobacteria bacteria are fast growing microorganisms that form spores, which may be

reasons for their successful cultivation in laboratory media. A likely explanation for the low

cultivation efficiencies (lower than 0.1% generally (Parkes and Sass 2009)) of the dominant

microbial phyla, i.e., Chloroflexi and JS1, may be partly due to the energy limitation conditions

in marine sediments, which when starving microorganisms are exposed to the rich in nutrients

laboratory media, may experience a substrate-shock, possibly as a result of uncoupling of

metabolic reactions (Parkes and Sass 2009). No isolate from the widely distributed marine

sediment Chloroflexi or candidate phylum JS1 has been obtained so far. Members from the

phylum Chloroflexi were maintained in anoxic medium (Köpke et al 2005, Lysnes et al 2004,

Webster et al 2011), however never isolated. The isolation of a Chloroflexi from marine

sediments may lead us to a better understanding of their physiology, metabolism and

ecological role in marine sediments.

Due to the widespread presence and high abundance of Chloroflexi in marine sediments

worldwide, Chloroflexi may play a critical role in the subseafloor ecosystem. Up to date,

scientific surveys in marine sediments have focused firstly on biogeochemical processes, and

secondly, on the microorganisms associated with these processes. However, microorganisms

associated with biogeochemical processes identified until now did not belong to the dominant

bacterial groups identified with DNA-based techniques, e.g., Chloroflexi, candidate phylum

JS1. Thus, very little is known regarding the dominant bacterial groups such as Chloroflexi and

Introduction

an in depth study on them may contribute to better understanding of the marine sediment

ecosystem.

1.3 ABUNDANCE AND DISTRIBUTION OF CHLOROFLEXI IN MARINE

SEDIMENTS

Bacterial members affiliated with the phylum Chloroflexi (formerly described as the ‘Green

non-sulphur’ Bacteria) are almost ubiquitously found in marine sediments (Biddle et al 2006,

Coolen et al 2002, Fry et al 2008, Inagaki et al 2006, Nunoura et al 2009, Parkes et al 2005,

Parkes et al 2014). Indeed, sequences affiliated to Chloroflexi are reported to be present in 66%

of the bacterial 16S rRNA gene clone libraries from marine sediments deeper than 2 mbsf (205

prokaryotic clone libraries analysed by (Parkes et al 2014)). Moreover, those 16S rRNA clone

libraries pointed out Chloroflexi as one of the most abundant, if not the most abundant,

bacterial phylum in marine sediments, with an average abundance of 25.5% (Parkes et al

2014). Sometimes, sequences affiliated to Chloroflexi add up to 80% of the total bacterial 16S

rRNA genes recovered in some marine sediment sites (Parkes et al 2005). In many occasions,

members of the Chloroflexi are associated with high organic matter contents (Parkes et al

2014), and are therefore thought to be heterotrophic and have a role in sedimentary organic

matter mineralization (Parkes et al 2014).

The phylum Chloroflexi is deeply branching within the domain Bacteria and some of its first

isolated members such as Chloroflexus, Herpetosiphon and Thermomicrobium are

thermophiles, chemoorganotrophs or/and conserve energy from light (Garrity and Holt 2001,

Oyaizu et al 1987). Members of the Chloroflexi have in common a unique, single-layer

membrane and are therefore monoderms, whereby the great majority stain Gram-negative

(Gupta et al 2013, Sutcliffe 2010, Sutcliffe 2011). The phylum currently encompasses six

named classes as well as candidate classes consisting of sequences derived from uncultured

organisms (Figure 1), which are metabolically diverse bacteria, ranging from aerobic

thermophiles, anoxygenic phototrophs to organohalide-respiring bacteria (Gupta et al 2013,

Hugenholtz and Stackebrandt 2004, Löffler et al 2013, Sekiguchi et al 2003, Yabe et al 2010,

Yamada et al 2006). In the last years, the isolation of a novel species of a nitrite-oxidizer and a

nitrate- and iron-reducers evidenced the expanded metabolic diversity still to be discovered

within the phylum Chloroflexi (Kawaichi et al 2013, Sorokin et al 2012).

Chloroflexi sequences retrieved from marine sediments are usually affiliated to the classes

Anaerolineae, Caldilineae (Blazejak and Schippers 2010) and Dehalococcoidia (Löffler et al

2013), as well as to other groups not classified as classes and comprising exclusively

environmental sequences such as the formerly known subphylum IV (Parkes et al 2014).

Classes Anaerolineae and Caldilineae were formerly known as subphylum I (Hugenholtz and

Introduction

Stackebrandt 2004, Yamada and Sekiguchi 2009) and their cultured members are

organoheterotrophs (Sekiguchi et al 2003, Yamada et al 2006). Class Dehalococcoidia was the

formerly known subphylum II (Hugenholtz and Stackebrandt 2004) with several cultured

members known for their ability to respire with organohalides (Bowman et al 2013, Löffler et

al 2013, Moe et al 2009). Among these three Chloroflexi classes, the most frequently detected

phylotypes retrieved from marine sediments affiliate to the Dehalococcoidia (Durbin and

Teske 2011, Inagaki et al 2003, Inagaki et al 2006, Jørgensen et al 2012, Webster et al 2006,

Wilms et al 2006). Moreover, PCR-independent metagenomic studies identified both

Dehalococcoidia affiliated 16S rRNA gene sequences and other DNA sequences most similar

to known Dehalococcoidia (Biddle et al 2008, Biddle et al 2011). Each of these three classes

may have different niches in marine sediments, as suggested by microbial community surveys

of South Pacific abyssal marine sediments, which evidenced a change in Chloroflexi

abundance and in Chloroflexi diversity and composition with depth (Durbin and Teske 2011).

In this study, sequences retrieved from oxic upper layer sediments (60-70 cmbsf) contained

13% of Chloroflexi of the so-termed “Chloroflexi VIb” (Durbin and Teske 2011) and the class

Anaerolineae (Yamada et al 2006). However, sequences retrieved from the anoxic deeper

sediment part increased proportionally with depth and belonged mainly to the class

Dehalococcoidia (Durbin and Teske 2011).

Introduction

Figure 1. Phylogenetic tree of the phylum Chloroflexi as classified by the SILVA 100 16S rRNA gene database.

The phylum Chloroflexi includes six named classes: Anaerolineae, Caldilineae, Dehalococcoidia,

Ktedonobacteria, Chloroflexi, and Thermomicrobia. Among the many groups, the class Dehaloccocoidia includes

the organohalide-respiring bacteria: Dehalococcoides mccartyi (Löffler et al 2013), Dehalobium chlorocoercia

DF-1 (May et al 2008) and Dehalogenimonas lykanthroporepellens strains BL-DC-8 & BL-DC-9 (Moe et al

2009), and sequences retrieved from marine sediments.

Introduction

But what do we know about bacteria of the class Dehalococcoidia? The class Dehalococcoidia

includes to date only a few isolated species: Dehalococcoides mccartyi (various strains isolated

(Löffler et al 2013)), Dehalogenimonas lykanthroporepellens (Moe et al 2009),

Dehalogenimonas alkenigignens (Bowman et al 2013), and Dehalobium chlorocoercia strain

DF-1 (May et al 2008). All of them are phylogenetically relatively closely related and all of

them are known for exclusively growing by respiring organohalides, a process called

“organohalide respiration” (Hug et al 2013b, Taş et al 2010b). In addition to the isolates, the

mixed cultures KB-1 (Duhamel et al 2004) and ANAS (Richardson et al 2002, West et al

2008) are known to contain organohalide-respiring Dehalococcoidia from the genera

Dehalococcoides. Moreover, several studies of mixed cultures from marine sediments enriched

with organohalides have detected Dehalococcoidia 16S rRNA gene sequences (Bedard et al

2007, Fagervold et al 2005, Fagervold et al 2007, Kittelmann and Friedrich 2008a, Kittelmann

and Friedrich 2008b, Watts et al 2005). All these organohalide-respiring Dehalococcoidia are

phylogenetically close to each other, forming a cluster within the class when 16S rRNA genes

are analysed (Figure 2). Indeed, Dehalococcoidia 16S rRNA gene sequences retrieved from

marine sediments are in most cases, relatively distantly related to cultivated organohalide-

respiring Dehalococcoidia, forming a so-called “sister lineage”, as well as several other deeper

branching clades (Durbin and Teske 2011, Inagaki et al 2006). Therefore, organohalide

respiration may be a possible metabolism for marine Dehalococcoidia, and evidence for

organohalide respiration in the marine subsurface was found when amplifying and sequencing

the gene of the key enzyme for organohalide respiration, i.e., reductive dehalogenase

(Futagami et al 2009). However, it cannot be concluded that all marine sediments containing

Dehalococcoidia have a metabolism based on organohalide respiration, since marine sediment

Dehalococcoidia and organohalide-respiring Dehalococcoidia form separate phylogenetic

clusters. In fact, attempts to enrich and isolate marine sediment Dehalococcoidia have been

unsuccessful so far (Köpke et al 2005, Süβ et al 2004). Getting to know marine sediment

Dehalococcoidia is an important task because they form a huge proportion of the microbial

sub-seafloor biosphere and may play key roles for the functioning of the planet.

Introduction

Figure 2. Phylogenetic tree of the class Dehalococcoidia based on 16S rRNA genes from the SILVA 100 database representing the different clusters of Dehalococcoidia

sequences retrieved from uncultured organisms, mostly from the subseafloor. Organohalide-respiring Dehalococcoidia members (isolated spp. or from enrichments or stable

isotope probing studies) are shown in green. 16S rRNA gene sequences from published single cell genomes of the Dehalococcoidia are shown in blue. A 16S rRNA gene from an

unpublished single cell genome is shown in red. The tree was calculated using the maximum likelihood algorithm, courtesy of Dr. Kenneth Wasmund, UFZ–Leipzig.

Introduction

1.4 ORGANOHALIDE RESPIRATION IN MARINE SEDIMENTS

Cultivated members of the Dehalococcoidia are highly specialized on organohalide

respiration for growth, i.e., they exclusively use organic compounds carrying halogen

substituents as terminal electron acceptors in their respiratory chain, coupled to hydrogen as

the sole electron donor (Hug et al 2013b, Richardson 2013, Taş et al 2010b). In this way, the

organohalide compound is being reduced and a halogen is removed from the organohalide

compound. This process has been extensively studied as some halogenated compounds, such

as tetrachloroethene (PCE) and trichloroethene (TCE), for example, are major groundwater

pollutants worldwide. PCE and TCE are transformed via organohalide respiration into the

dichloroethenes 1,2 cis-DCE, 1,2 trans-DCE or 1,1 DCE by a wide spectrum of

microorganisms, such as the proteobacterial Sulfurospirillum multivorans, Desulfuromonas

spp., Geobacter lovleyi, and the Firmicutes Desulfitobacterium spp. and Dehalobacter

restrictus (Christiansen and Ahring 1996, Gerritse et al 1996, Holliger et al 1998, Utkin et al

1994). However, only members of Dehalococcoides mccartyi within the Dehalococcoidia are

known to completely metabolically respire PCE to the non-toxic ethene (Maymó-Gatell et al

1997). Various Dehalococcoides mccartyi strains have been observed to grow not only with

chlorinated aliphatics but also with many other aromatic organohalides. For instance,

Dehalococcoides mccartyi strain CBDB1 is able to respire a broad spectrum of chlorinated

and brominated aromatic compounds including chlorobenzenes, bromobenzenes,

chlorophenols, dioxins and polychlorinated biphenyls (PCBs) (Adrian et al 2007, Bunge et al

2003, Cooper et al 2015, Wagner et al 2012, Yang et al 2015).

Organohalide-respiring microorganisms represent a bioremediation option for contaminated

sites because the majority of organohalides that have been extensively used in agriculture and

industry are highly toxic and persistent in the environment (Braeckevelt et al 2011, Cichocka

et al 2010, Imfeld et al 2011, Löffler and Edwards 2006, Maphosa et al 2012, Mészáros et al

2013, Narihiro et al 2010, Pérez-de-Mora et al 2014, Taş et al 2010a, Taş et al 2011, Verce et

al 2015). Despite being mainly known for their anthropogenic origin, organohalides are also

naturally produced by a wide array of biological and chemical processes in the environment,

and oceans are the largest source of biologically produced halogenated organic compounds on

Earth (Gribble 2003, Häggblom and Bossert 2003). Regarding the biological processes, an

immense number of marine organisms including algae, sponges, corals, invertebrates and

bacteria, among others, produce halogenated compounds, the polybrominated ones being the

most common ones (Ashworth and Cormier 1967, Baker and Duke 1973, Herrera-Rodriguez

et al 2011, Lira et al 2011, Pauletti et al 2010, Pedersén et al 1974, Utkina et al 2001, White

and Hager 1977). Chemical processes include i) geothermal processes at high temperature and

Introduction

pressure, e.g., volcanic eruptions, and ii) halogenation during the degradation and diagenesis

of organic matter (Häggblom and Bossert 2003, Keppler et al 2000). Chemically halogenated

organic matter buried in marine sediments can therefore be used for respiration by

organohalide-respiring bacteria. Thus, organohalide-respiring bacteria have long evolved to

thrive on natural organohalogens during Earth’s history. In this respect, halogenation and

dehalogenation are part of the halogen cycle on Earth. Evidence of dehalogenation processes

mediated by microorganisms in marine and estuarine environments already exists (Ahn et al

2003, Futagami et al 2013, Häggblom et al 2003, King 1988, Monserrate and Häggblom

1997). In fact, reductive dehalogenase-homologous (rdhA) genes have been amplified from

various marine sediment locations within the Nankai Trough plate-subduction area in Japan

(Futagami et al 2009), suggesting the occurrence of organohalide respiration in marine

sediments. Sediment slurries incubated over 200 days showed activity with halophenols

(bromophenols, chlorophenols, iodophenols) in shallow sediments and, after RNA studies,

Desulfuromonadales bacteria were identified as predominant (Futagami et al 2013). In

terrestrial uncontaminated soils, Dehalococcoidia correlated positively with organochlorine

concentration and organic carbon content. In addition, Dehalococcoidia numbers increased

when cultured with enzymatically produced organochlorines concomitant to chlorine

accumulation (Krzmarzick et al 2012), indicating that some Dehalococcoidia identified in

pristine sites may have an organohalide-respiring metabolism. The metabolism of marine

subsurface Dehalococcoidia from pristine sediments that fall outside of the clade containing

cultured species (i.e., the predominant groups of Dehalococcoidia found in the marine

subsurface (Kittelmann and Friedrich 2008a)), is completely unknown, i.e., whether they

respire either organohalides or other known anaerobic compounds, e.g., sulphate, nitrate, iron,

manganese, possibly even fermentation. For this reason, enrichments and isolates of

Dehalococcoidia from the marine subsurface may substantially aid in understanding the

widely distributed, abundant and enigmatic marine subsurface Dehalococcoidia. Additionally,

more information on the specific ecological distributions and their relationships with the

geochemistry of the marine subsurface will also greatly aid in our understanding of these

microorganisms.

1.5 STUDY AREA: BAFFIN BAY

During this thesis work, focused sampling of marine sediment cores from the Baffin Bay was

undertaken and studied for their geochemistry and microbial communities, and therefore this

section aims to provide a background into the characteristics of the Baffin Bay and its

subsurface. The Baffin Bay is a relatively isolated sea located between the Eastern coast of

Northern Canada and Greenland (Figure 3). From North to South, the Baffin Bay connects the

Arctic Ocean to the Labrador Sea and the North Atlantic Ocean. It is thought that in the late

Introduction

Paleogene, the Baffin Bay formed the main link for surface waters exchange between the

Arctic and the Atlantic Ocean (Srivastava et al 1989). Bathymetrically, the Baffin Bay is an

ocean basin formed of a large and deep plain in the centre of the bay with depths that exceed

2,300 m and two continental shelves, one shelf on the side of Greenland, and the other shelf

on the side of the Baffin Island, in Canada. The continental shelf on the side of Greenland is

much wider than the continental shelf at the Baffin Island. In both shelves, and running across

them, there are submarine canyons that cut the shelves (Tang et al 2004). The current

circulation is cyclonic (counter-clockwise) with the formation of an eddy at the northern part

of the Baffin Bay. Two main currents are present in the Baffin Bay: the subsurface, salty and

warm West Greenland Current going northward along the coast of Greenland and the near-

surface Baffin Island Current with fresh and cold waters coming from the Arctic Ocean,

which enters the Baffin Bay through the Kane Basin and the Smith Sound and flows

southwards along the Western coast of Canada to the Labrador Sea (Münchow et al 2015,

Tang et al 2004). Three water masses are present in the Baffin Bay although not in all areas.

The most surficial water mass, present in the upper 100–300 m is an Arctic water mass,

however not present in the southeast of the Baffin Bay. An intermediate water mass at 300–

800 m, named West Greenland Intermediate water, is present in the central area of the Baffin

Bay. Below 1,200 m, there is a deep Baffin Bay water mass, which covers all areas of high

depths (Tang et al 2004).

The Baffin Bay, as any other Arctic region, experiences contrasted sun irradiance periods that

turn into contrasted seasonal periods of daylight and temperature along the year. Extreme low

air temperatures in winter result in sea-ice formation on the surface of the ocean waters,

which restricts the primary production performed by photosynthetic planktonic organisms and

algae (Hulth et al 1996). Although some algae are photosynthetically active under the sea-ice

cover, this algal primary production of biomass is also restricted to the summer months, as it

depends on the sea-ice thickness, snow cover and the water column stratification (Boetius et

al 2013). Consequently, to the restriction of the primary production in the Arctic, the organic

matter supply to Arctic sediments is reduced (Hulth et al 1996). The Baffin Bay is sea-ice

covered most of the year. It is completely free of sea-ice only in the months of August and

September (Tang et al 2004). Sea-ice strongly restricts access to this area for navigation and

of course, for sampling. This partially explains why very few microbiological studies have

been performed in the Baffin Bay.

Introduction

Figure 3. Baffin Bay map modified from (Tang et al 2004). K: Kane Basin, S: Smith Sound. Baffin Island and

Ellesmere Island belong to Canada. Darker blue colour corresponds to deeper bathymetry.

In 1985, the Ocean Drilling Program (ODP) Leg 105 sampled the southern Baffin Bay at the

site 645, which is located on the slope off Baffin Island, for geological and geochemical

purposes. This study also included two further sites in the Labrador Sea (Srivastava et al

1989). During ODP Leg 105, cores down to 1,147 mbsf of depth were sampled at site 645,

with a water depth of 2,018 m in the only study ever done in Baffin Bay sediments so far. Site

645 had high sedimentation rates, averaging to 60 m Ma-1, strongly influenced by terrigenous

input of clay, silt, sand, and abundant dropstones coming from nearby continental lands and

minor biogenic components (Srivastava et al 1989, Stein et al 1989). Geochemical

characterization of core pore-waters at site 645 indicated a decrease of sulphate

concentrations in the upper 25 mbsf from 22 mM to less than 1 mM, and complete sulphate

depletion at 35 mbsf (Zachos and Cederberg 1989). This sulphate depletion corresponds to

Pliocene-Pleistocene hemipelagic sediment layers with high sedimentation rates above 120 m

Ma-1 (Zachos and Cederberg 1989). The decrease in pore-water sulphate concentrations is

Introduction

most likely the result of microbial sulphate reduction. Sulphate reduction products are

sulphide and bicarbonate. Sulphide may lead to the formation of pyrite, which is a common

mineral in the upper sediment layers of site 645 where sulphate is depleted. In addition,

bicarbonate may cause a decrease of pore-water calcium and magnesium ions in pore-water

depth profiles, which was the case in upper sediment layers, and may be linked to carbonate

precipitation of calcite and dolomite minerals (Zachos and Cederberg 1989). Organic carbon

content at site 645 was relatively high and increased with depth with values that ranged

between 0.5% to almost 3% (Stein et al 1989). The composition of this organic matter is

mostly terrigenous, and thus originating by nearby terrestrial lands (Stein et al 1989).

In 2007, as part of a study for the international Polar Year, Galand and co-workers took one

sample of the Baffin Bay intermediate water mass at the southern Baffin Bay at a depth of

1,000 m in the water column for microbiological studies. Galand and co-workers investigated

if there were patterns of microbial biogeographical distribution in various Arctic water bodies,

and found that it was explained by the water circulation and hydrography of the Arctic Ocean

(Galand et al 2010).

The Baffin Bay is an excellent location to conduct studies of microbial communities in

sediments for several reasons: i) the nature of its sediments, particularly the quantity and

composition of its mostly terrestrially derived organic matter ii) topographically, it is a

relatively isolated ocean basin with lower influence from global ocean circulation. This

relative isolation may therefore have reduced ubiquitous bacterial dispersal, and selected for

bacteria highly adapted to the processes within this area, e.g., glacial processes, and iii) it

combines an ocean basin with continental shelves and deep abyssal plain so that the influence

of gradients of environmental parameters (e.g., water column depth, sediment depth, distance

to land) on the distribution of microorganisms can be studied.

For these reasons, the Baffin Bay was selected as the study site. Emphasis was laid on

investigating the presence, distribution, and diversity of bacteria of the class Dehalococcoidia

in Baffin Bay sediments.

Introduction

1.6 AIM OF THE STUDY

The overall aim of this study was to understand the ecological role of Dehalococcoidia in

marine sediments. As a step towards this overall aim, it was tried to identify physiological

characteristics of marine Dehalococcoidia. To accomplish the aim, two main experimental

approaches were chosen:

Approach 1. Use of cultivation techniques to describe the physiology of marine

Dehalococcoidia, and in particular, their respiration mode. The cultivation relied on batch

cultures containing marine sediment bacteria as inoculum. The medium was based on the

growth medium described for Dehalococcoides mccartyi, which are phylogenetically the

closest and best known cultivated members to marine Dehalococcoidia. As a tool to evaluate

growth of Dehalococcoidia, qPCR and chemical analysis of metabolic products was chosen.

Several tasks within this approach were defined:

a) Study if Dehalococcoidia could be anaerobically cultivated in the laboratory at

atmospheric pressure, within a reasonable growing timespan, i.e., months.

b) Study if Dehalococcoidia physiology may be similar to Dehalococcoides

mccartyi in that they use organohalides as a terminal electron acceptors in their

respiratory chain.

c) Study of other types of respiratory modes, e.g., sulphate, iron, manganese or

humic acids.

Approach 2. Description of the natural occurrence of Dehalococcoidia, their distribution and

abundance in marine sediments associated to biotic (i.e., other bacterial groups), and abiotic

(i.e., sediment geochemistry, geographical location, depth) parameters in marine sediments, to

identify natural conditions promoting their growth. Such an investigation was performed with

an in situ approach in sediments of the Baffin Bay in the Arctic. To accomplish this second

approach, three specific tasks were defined:

a) Description of the presence, abundance and distribution of Dehalococcoidia in

marine sediments at different sites and depths with a quantitative approach

using qPCR.

b) Description of the microbial diversity in a geochemical gradient of organic

matter contents and biogeochemical parameters within a transect from the shelf

and into the basin of the Baffin Bay, including also other bacterial groups

associated with Dehalococcoidia.

Introduction

c) Identification of correlations of Dehalococcoidia abundance with

biogeochemical parameters to obtain indications of preferable growth

conditions to extrapolate to Dehalococcoidia physiology.

Material & Methods

2 MATERIAL & METHODS

2.1 CHEMICALS

All chemicals were purchased from Sigma-Aldrich (MO, USA), Merck KGaA (Darmstadt,

Germany) or AppliChem (Darmstadt, Germany) at highest purity available. The gases

“Biogon” (80% nitrogen, 20% carbon dioxide; v/v), hydrogen, and nitrogen were purchased

from Air Liquide (Paris, France). Humic acids were bought from Carl Roth (Karlsruhe,

Germany).

2.2 SEDIMENT SAMPLES

Sediments from various sources were used. These sediment sources include the Greenlandic

side of the Baffin Bay, the continental shelf at the central off Chile, Århus Bay in Denmark,

and the continental shelf at the South-West of Ireland (Table 1). Sediments from the Baffin

Bay were collected during the ARK XXV/3 expedition on board of the research vessel

Polarstern in August–October 2010. Other sediments were collected during other expeditions

and kindly supplied by Dr. Timothy G. Ferdelman from the Max Planck Institute for Marine

Microbiology in Bremen, Germany.

Baffin Bay sediments were cored at 34 sites of the Baffin Bay, which extended from the coast

of Greenland to the central area of the Baffin Bay (Damm 2010). Due to the geology of the

area, with abundant dropstones brought by glaciers, some of the sites yielded unsuccessful

recovery of sediments. From those cores with successful recoveries, ten cores were selected

based on recovery length (longest cores), intact state of recovered sediments, and location of

the site. Selected sediments were used for two biogeochemical and microbiological studies: i)

involving three geographical distinct areas “Northern Greenlandic shelf” (sites 363 and 371 in

Figure 4), “central deep basin” (sites 389, 391, and 453 in Figure 4), and “Southern slope”

(sites 486 and 488 in Figure 4), (Algora et al 2013), and ii) involving seven sites along a

North-to-South, shelf-to-basin transect (Figure 5) (Algora et al 2015).

Material & Methods

Table 1. Oceanographic details from the sites where the sediment samples used in this study were cored.

Sample location Cruise Collection

date

Site Coordinates

(latitude, longitude)

Water

depth (m)

Sediment

depth (mbsf)

References

Århus Bay, Denmark – 2008 Århus M1 GC

A/2

not known not known 4.44a from Dr. T.G. Ferdelman,

MPI Bremen. Metrol project:

http://metrol.mpi-bremen.de

Eastern slope, Porcupine Seabight,

SW continental margin of Ireland

IODP Leg 307 2005 U1318 51°26.16'N, 11°33.0'W 423 23.15a (Webster et al 2009)

NW-shoulder, Challenger Mound,

Porcupine Seabight,

SW continental margin of Ireland

IODP Leg 307 2005 U1317 51°22.8'N, 11°43.1'W 781–815 227a (Webster et al 2009)

Central off Chile continental margin SO-156/3 2001 GeoB 7155-4 34°35.00'S, 72°53.11'W 2,744 4.37–4.42a (Treude et al 2005)

Central off Chile continental margin SO-156/3 2001 GeoB 7165-2 36°32.32' S, 73°40.02' W 799 6.35–6.40a (Treude et al 2005)

Baffin Bay, Greenland ARK-XXV/3 2010 363 76° 52.92' N, 71° 34.01' W 938 4.69b (Algora et al 2013)

Baffin Bay, Greenland ARK-XXV/3 2010 365 76° 39.04' N, 71° 18.79' W 658 3.67b (Algora et al 2015)

Baffin Bay, Greenland ARK-XXV/3 2010 371 75º 58.24' N, 70° 34.86' W 598 4.05b (Algora et al 2013)

Baffin Bay, Greenland ARK-XXV/3 2010 383 75º 17.69' N, 69º 53.75' W 674 2.32 b (Algora et al 2015)

Baffin Bay, Greenland ARK-XXV/3 2010 387 74° 50.42' N, 69º 27.14' W 1,300 3.32b (Algora et al 2015)

Baffin Bay, Greenland ARK-XXV/3 2010 389 74° 37.05' N, 69º 13.75' W 1,716 4.24b (Algora et al 2013)

Baffin Bay, Greenland ARK-XXV/3 2010 391 74° 23.36' N, 69º 01.22' W 1,864 4.27b (Algora et al 2013)

Baffin Bay, Greenland ARK-XXV/3 2010 453 73° 19.37' N, 64° 58.11' W 2,300 4.69b (Algora et al 2013)

Baffin Bay, Greenland ARK-XXV/3 2010 486 72° 24.51' N, 60° 48.85' W 645 4.69b (Algora et al 2013)

Baffin Bay, Greenland ARK-XXV/3 2010 488 72° 08.80' N, 60° 58.86' W 1,493 4.69b (Algora et al 2013)

Sediment depth: a refers to the depth that was used for inoculum for sediment cultures, b refers to the deepest part of the core that was retrieved, from seafloor until the indicated

depth. For this later case, the entire core, from 0 mbsf to the indicated depths, was used for various analyses. SW–South-West, NW–North-West.

Material & Methods

Figure 4. Map of the Baffin Bay indicating investigated sites from three distinct geographical areas.

These areas were: “Northern Greenlandic shelf” (triangle symbol; sites 363 and 371), “central deep basin”

(circle symbol; sites 389, 391, and 453), and “Southern slope” (square symbol; sites 486 and 488). These

sites were studied for differences in geochemical parameters and for the presence, distribution, and

abundance of specific microbial groups. The symbol and colour code are consistently used for each site in

all figures of this thesis. Modified from (Algora et al 2013).

Material & Methods

Figure 5. Map of the Baffin Bay showing the location of the sediment sampling sites investigated for a

study focused on the microbial diversity associated to geochemical parameters along a North-to-South,

shelf-to-basin transect; a) locations from the seven sites within the Baffin Bay that comprise the transect;

and b) sediment cut-section scheme of the seven sites investigated. Modified from (Algora et al 2015).

2.2.1 Core sampling

Core sampling and sediment processing for those sediments that were donated by other

groups are described elsewhere (see Table 1 for references). Sediments from the Baffin

Bay were sampled within this work with a gravity corer equipped with a 4.70 m core

barrel (Rehau AG & Co.) and a 90 mm (outside diameter) PVC liner as described

(Algora et al 2013, Damm 2010). Recovered cores were divided into one metre sections

on board using a PVC tube cutter and a clean masonry spatula, capped and sealed with

electrical tape. All cores were kept at 4°C until further subsampling (Figure 6).

Material & Methods

Figure 6. Core sampling at the Baffin Bay by means of a gravity corer during the expedition ARK

XXV/3.

2.2.2 Core subsampling

Core subsampling consisted of retrieving five different types of samples from each core:

i) sediment for free gases (not adsorbed to sediments, i.e., methane and carbon dioxide)

analyses, ii) pore-water for dissolved ion concentration analyses, iii) sediment for

anaerobic cultivation of indigenous sediment microorganisms iv) sediment for DNA-

based analyses v) sediment for organic carbon content, elemental composition, and

mineralogy.

Samples for gases, pore-water, and DNA-based analysis were collected at specific

depth-sampling points, every 50 cm along the core and starting at 25 cmbsf for all cores

(see scheme in Figure 7).

Material & Methods

For gases, 5 ml sediment was sampled with sterile needle-adapter-cut syringes

immediately after drilling a hole on the PVC liner at each depth-sampling point (Figure

7). Once sampled, liner holes were immediately sealed with electrical tape and the

sampled sediment was placed in 50 ml glass serum vials which were previously filled

with 20 ml of a 1 M NaOH solution in order to stop any microbial activity (Figure 7).

Filled vials were instantly sealed with gas-tight rubber stoppers and aluminium crimp

caps and mixed until the sediment was completely suspended in the solution. Vial

storage was at 4°C until vial headspace analysis was performed by Gas

Chromatography (GC).

Figure 7. Scheme of depth-sampling points for every core (left), and sediment sampling for gas analyses

(right).

Pore-water samples were taken immediately after gas sediment sampling (at the same

core positions). Between 8 to 10 ml pore-water was recovered from all cores at every

depth-sampling point using a rhizon sampler (CSS-F, 5 cm or 10 cm porous length, 2.5

mm tip diameter, Rhizosphere Research Products, Wageningen, NL). Rhizon samplers

were fine plastic tubes with a porous membrane on one end, which was introduced into

the sediment, and a syringe adaptor on the other end of the tube (Figure 8). The pore-

water sampling was done at 4°C inside a cool room.

Material & Methods

Rhizons used for pore-water sampling were previously dipped in double distilled water

for 1–2 h before use, checked for proper functioning (Figure 8), and introduced into

each depth-sampling point inside each core section using the holes already made from

the gas sampling in a 4°C room (Figure 8). Rhizons where connected to a sterile syringe

which had the plunger partly out to create vacuum and facilitate pore-water extraction

from sediment (Figure 8). Through the rhizon porous membrane, the pore-water

penetrated and flowed to the syringe barrel where accumulated. The rhizon was left

inside the sediment until the syringe was filled and, in any case, for at least 4 hours. The

pore-water samples were transferred to polypropylene vials, preserved with 1% (v/v) of

a 65% (v/v) concentrated nitric acid solution, and stored at 4°C until analysed (Figure

8). The polypropylene vials were previously washed with 1% (v/v) nitric acid solution

and rinsed with double-distilled water. Rhizons were cleaned and rinsed with double-

distilled water and further re-used.

Figure 8. Pore-water sampling procedure, which was performed using rhizon samplers. See text for

further details.

For retrieval of the remaining sample types, each core section was sliced in two halves.

For that, the PVC liner was laterally cut with two sledge-mounted vibrating saws

(Figure 9). The two sediment halves were separated with a clean stainless steel thread

and a clean masonry spatula. One half was used for stratigraphic studies and stored at

the core archive of the Bundesanstalt für Geowissenschaften und Rohstoffe (BGR) in

Hannover, Germany, after the expedition. The other half was used for sediment

sampling on board, which was done immediately after core slicing. Before sediment

sampling, the sediment sliced area was removed to avoid contamination with a sterilized

masonry spatula and sterile scalpels. All sediment samples taken for microbiological

purposes were strictly sampled from the untouched and uncontaminated interior of

every core.

Material & Methods

Figure 9. Slicing of one-metre core sections into two halves, which was executed for further sampling of

sediment for microbiological purposes.

Sediment samples for anaerobic cultivation of microorganisms were taken with sterile

needle-adapter-cut 5 ml syringes immediately after the core was sliced. Sediment

samples of 3–20 ml volume were inoculated into glass serum bottles that contained

various volumes (27, 45, 75, or 90 ml) of sterile anoxic minimal media. The minimal

media components, which are detailed in Appendix 1 (section 6.1), contained in some

cases amendments of yeast extract (0.02%), selenium (6 µg l-1) and tungsten (8 µg l-1),

and acetate (5 mM). The medium was buffered and reduced with various reducing

agents (either titanium (III) citrate, or iron sulphide, or sodium sulphide together with L-

cysteine; see Appendix 1) at least a day prior to sediment inoculation and had a

headspace atmosphere of “Biogon” (consisting of 80% nitrogen and 20% carbon

dioxide; v/v) which occupied ~50% of the glass serum bottle volume. Glass serum

bottles were sealed with rubber or Teflon-lined butyl rubber stoppers and aluminium

crimps. Prior to inoculation, glass serum bottles containing the reduced and sterile

medium were de-crimped, de-capped, and inoculated with sediment contained in the

needle-adapter-cut syringe and with help of the plunger. Once inoculated, the glass

serum bottle was partly capped and purged with a Biogon stream which was supplied

with a sterile needle connected to a 0.2 µm filter to the Biogon rubber pipe source for

the headspace exchange (Figure 10, right). Afterwards, the glass serum bottle was

capped, crimped and gently mixed.

Material & Methods

Figure 10. Sediment sampling for inoculation in glass serum bottles (left). Headspace gas was purged

with the mixture gas Biogon (80% N2, 20% CO2; v/v) to eliminate the air which may have entered during

the inoculation procedure (right).

Sediment samples for DNA-based analyses were retrieved with sterile needle-adapter-

cut 5 ml syringes at every depth-sampling point, or as close as possible, in an effort to

avoid possible contaminated sediment after the gases or pore-water sampling at each

sampling point. Sediment samples were placed in sterile 15 ml Falcon tubes, labelled

and immediately frozen at -80°C (Figure 11). Sediment samples were stored at -80°C

until analysed.

Figure 11. Sediment sampling for further DNA-based analyses.

Sediment samples for analysis of organic carbon content, elemental composition, and

mineralogy were collected at various intervals of 50–100 cm along the core. Sediment

samples were placed in sterile Schott glass bottles of 100 or 250 ml (Schott Duran),

which were closed with butyl septa and plastic screw caps under a nitrogen atmosphere

and stored at 4°C until measured.

2.3 SEDIMENT GEOCHEMICAL ANALYSIS

The determination of methane concentrations from the headspace of the sediment

sample vials was performed using a GC-FID equipped with a 6’ Hayesep D column

(SRI 8610C, SRI Instruments) at 60°C. This was done by collaborators at the BGR as

described elsewhere (Algora et al 2013).

Material & Methods

Pore-water samples were analysed to obtain ion compositions using an inductively

coupled plasma mass spectrometry (ICP-MS) instrument (Perkin Elmer Sciex Elan

5000). This was performed by collaborators in the BGR as previously described (Algora

et al 2013).

Sediment carbon content was analysed as total organic carbon (TOC) and total carbon

(TC) from the sediment samples frozen at -80ºC, after the sediment sample for DNA

was already analysed, from cores 363, 365, 371, 383, 387, 389, and 391 (Figure 5). For

that, a sample from the frozen sediment from each core and depth was thawed, dried,

homogenized with a mortar, and grinded. One gram of the dry sediment was analysed in

a LECO RC-412 carbon determinator (LECO Corporation) with a temperature program

of 80°C min-1 from 100°C to 530°C for the measurement of TOC, followed by

100°C min-1 from 530°C to 1000°C for the measurement of total inorganic carbon. All

measurements were done in duplicate and values are reported in weight percentages as

mean of duplicate measurements.

Sediment samples from cores 363, 389, and 486 (Figure 4) were examined for sediment

mineralogy by X-Ray Powder Diffraction (XRD) patterns, for sediment elemental

composition by wavelength dispersive x-ray fluorescence spectrometry (WD-XRFS) in

their mineral material, and for TOC and TC. These analyses were performed by

collaborators at the BGR as described elsewhere (Algora et al 2013).

In addition, stable isotope composition of the sediment organic content (organic carbon

13C) for cores 363, 389, and 486 was determined in order to obtain an indication of the

sediment organic carbon origin, either terrestrial or marine. For this, 5 mg of sediment,

which was previously freeze-dried and de-carbonated with hydrochloric acid, was

wrapped in tin caps (3.5 x 5 mm, HEKAtech) and analysed in an Elemental Analyzer

(EuroEA3000, Euro Vector) coupled via a ConFlow III (Thermo Fisher Scientific) to a

MAT 253 isotope ratio mass spectrometer (Thermo Fisher Scientific) as previously

described (Algora et al 2013). Triplicate 13C values are expressed in delta notation

(δ13C) relative to the Vienna PeeDee Belemnite (VPDB) standard and reported in per

mil (‰) (Coplen 2011).

δ13C = ((13C/12C)sample / (13C/12C)VPDB-standard)-1

2.4 CULTIVATION OF MICROORGANISMS IN SEDIMENT CULTURES

For the cultivation of microorganisms indigenous to the sediments, a defined anoxic

minimal liquid medium was used. The medium composition is given in Appendix 1,

section 6.1, and was based on the medium used for the cultivation of Dehalococcoides

Material & Methods

mccartyi strain CBDB1 as described by L. Adrian (Adrian 1999). The medium was

prepared by adding a mineral solution (Widdel 1980), a trace metal solution “SL-9”

(Tschech and Pfennig 1984), sodium acetate (5 mM) as carbon source, and resazurin as

redox indicator to Milli-Q water (see Appendix 1, section 6.1, for specific components

and concentrations in the solutions). In order to eliminate oxygen from the medium,

dissolved air was removed by applying vacuum to a one- or two-litre glass Schott bottle

filled with the medium for at least one hour. Subsequently, the medium was purged with

pure nitrogen gas for 30 min and further transferred to an anaerobic glovebox where it

was dispensed in aliquots to glass serum bottles. Aliquot volumes were 15 ml, 30 ml, 60

ml or 90 ml, leaving a headspace of 40–55% volume in the glass serum bottle. Filled

glass serum bottles were left inside the anaerobic glovebox for fifteen minutes to one

hour, to exchange the gas phase to the glovebox atmosphere composed of nitrogen and

2–3% hydrogen. Glass serum bottles were then sealed with Teflon-lined butyl rubber

stoppers and aluminium crimp caps. The bottles were autoclaved, and once they reached

ambient temperature, buffered with bicarbonate to pH 7.0–7.3 (see Appendix 1, section

6.1.3, for bicarbonate solution). Afterwards, reducing agents (Table 2, see Appendix 1,

sections 6.1.5 and 6.1.6 for preparation of the solutions) were added. Reducing agents

were allowed to equilibrate for at least 12 h, to overnight. Vitamins were added from a

filtered stock solution (Vitamin 7; (Adrian 1999)).

Table 2. Reducing agents combinations used in this study.

Mixture

Composition

Concentration

Titanium (III) citrate 15% (0.1 M Ti(III), 0.2 M citrate)

1.3%

Iron sulphide solution

1.6%

III

Sodium sulphide

0.3 mM

L-cysteine

0.2 mM

For inoculation, 9–10% (v/v; wet sediment) of sediment (various origins, Table 1) was

transferred to the glass serum bottles containing the sterile medium with sterile scalpels

inside an anaerobic glove box, except sediments from the Baffin Bay which were

inoculated on board as previously described. The sediment inocula were always taken

from inner undisturbed core parts, avoiding contamination with the liner or any other

recipient holding core intervals.

Material & Methods

The headspace gas phase from the glass serum bottles was interchanged to the gas

mixture Biogon by five cycles of vacuum and gas mixture addition, and further

pressurized to 0.2 bar overpressure. In addition, hydrogen was added up to a pressure of

0.3 bar, which was supplied as a potential electron donor for microorganisms. Various

compounds (Table 3) were individually supplied as potential electron acceptors through

the septum with a syringe from sterile stock solutions. Some compounds were added as

solids, which were autoclaved inside the glass serum bottle with the medium, except for

4-bromo-3,5-dimethoxy benzoic acid and 2,4,6-tribromoresorcinol. Sediment cultures

were set up at least in duplicates. Controls without sediment, without electron acceptors,

and with autoclaved sediments were always prepared, to evaluate possible abiotic

processes within the sediment or medium and microbial growth without electron

acceptors. Antibiotics against Gram-positives (vancomycin and ampicillin) and the

inhibitor against methanogens (2-bromoethanesulfonate) were used to detect a possible

effect on compound transformation and obtain an indication of the type of

microorganism performing the transformation. Sediment cultures were statically

incubated at 30ºC in the dark.

Table 3. Compounds added as potential electron acceptors to the sediment cultures. Concentration refers

to final concentration in sediment cultures.

Electron acceptor

Type

Name

Stock solution details

Concentration

Halogenated 1,2,3-Trichlorobenzene (1,2,3-TCB) Crystals solved in 1,2,4-

TCB in a 1:1 solution

15 µM

1,2,4-Trichlorobenzene (1,2,4-TCB) Liquid. In a 1:1 solution

with 1,2,3-TCB

15 µM

2-Monochlorophenol (2-MCP)

Solved in methanol

50 µM

2,3-Dichlorophenol (2,3-DCP)

Solved in methanol

50 µM

Tetrachloroethene (PCE)

Solved in acetone

90 µM

4-Bromo-3,5-dimethoxy benzoic acid

Added as crystals

~ 0.1 g l

-1

2,4,6-Tribromoresorcinol

Added as crystals

~ 0.1 g l

-1

Hexachlorobenzene

Added as crystals

~ 2 g l

-1

Organic

Humic acids

Added as solid

~ 0.3 g l

-1

Inorganic Ferric iron, Fe(OH)3 Added from an anaerobic

sterile solution

5.3 mM

Manganese oxides, MnO2 Added from an anaerobic

sterile solution

10 mM

Sulphate, Na2SO4 Added from an anaerobic

sterile solution

20 mM

Material & Methods

Sediment cultures were sampled at different time points of incubation to monitor the

16S rRNA gene copy numbers of Dehalococcoidia with molecular biology methods

(see section 2.7). These time points were within 8 and 17 months of incubation for

Århus sediment cultures, and 4, 10, and 12 months of incubation for Chile sediment

cultures. The sampling consisted of withdrawing 1 ml (or eventually 1.5 ml) of culture

after gentle homogenization of the deposited sediment particles. The sample withdrawal

was performed through the septum with sterile needles and syringes. Samples were

stored in 2 ml Eppendorf tubes at -80ºC.

2.5 ISOLATION OF PURE STRAINS BY CULTIVATION IN DEEP-

AGAROSE DILUTION TUBES

2.5.1 Media, solutions and preparation procedure

Growth of bacteria in semisolid media was obtained by preparing deep-agarose dilution

tubes. For this, glass tubes of 16 ml volume were filled with 2 ml of 2% (final

concentration in the tube of 0.3%) low-melting agarose (SeaPlaque®, Biozym

Scientific), autoclaved and allowed to cool down at ambient temperature. Sterile

agarose in the tubes was melted in a 80 ºC preheated heatblock, and cooled down at

60ºC before further addition of 10 ml of sterile anoxic medium, which was buffered

with sodium bicarbonate and warmed to 35ºC in a water bath. Further, the tubes were

closed with rubber caps and maintained inside a 32–35 ºC water bath. Addition of

0.2 ml titanium (III) citrate and electron acceptors in identical concentrations to liquid

sediment cultures was performed. Tubes were sealed with butyl rubber stoppers and

plastic screw caps. The headspace atmosphere was exchanged to the Biogon gas

mixture through a series of seven vacuum and Biogon gas exchange steps. Tubes were

mixed by turning them up and down and maintained at 35ºC for at least one hour to

allow equilibration of the reducing conditions. Inoculation was done through the rubber

stopper using the previously set up sediment cultures when the sediment cultures were

one week old.

A series of dilutions of four deep-agarose dilution tubes per sediment culture was

established aiming to obtain single colonies (Figure 12). For first dilution of the deep-

agarose tubes, 0.5 ml liquid sediment culture was added as inoculum. Second and third

dilution deep-agarose tubes were inoculated with 0.2 ml from the previous dilution tube

and the forth dilution tube was inoculated with 0.5 ml from the third dilution tube.

Deep-agarose dilution tubes without inoculum were set up as medium blanks.

Overpressure of 0.1 bar hydrogen gas was applied to each tube headspace as an

Material & Methods

available potential electron donor. Tubes were mixed by two times turned up and down

for mixing and solidified in an ice water bath. Incubation was performed in the dark at

30ºC. Tubes were visually observed periodically to monitor colony formation.

Figure 12. Deep-agarose dilution tubes were inoculated with 0.5 ml from a liquid sediment culture and a

dilution series was established aiming to obtain single colonies.

2.5.2 Picking and transferring of colonies

Selected single colonies formed in the tubes were picked inside an anaerobic glovebox

and transferred into 10 ml sterile anoxic medium. The 10 ml medium was pH buffered

and reduced with titanium (III) citrate prior to colony transfer. The picking process

involved using a sterile Pasteur pipette or a sterile 0.60 mm diameter x 80 mm length

needle connected to a syringe, which already contained a small volume (1–2 ml) of

sterile medium reduced with and titanium (III) citrate. Once the colony was inoculated

into the 10 ml liquid medium, the electron acceptor that the colony was exposed to in

the deep-agarose tube was added in the same concentration. The glass serum bottle

headspace was flushed with Biogon to an overpressure of 0.2 bar. Hydrogen was added

to an overpressure of 0.3 bar. Incubation was done statically in the dark at 30 ºC.

New deep-agarose dilution tubes were prepared in duplicate from the cultures

inoculated from single colonies to observe the morphology of colonies and to compare

it to the former original colony. The agarose tube preparation was done after two weeks

of incubation of the cultures inoculated from colonies.

A selection of colonies, either directly picked from the deep-agarose tubes, or 1.5 ml

from liquid sub-cultured colonies was sampled for later DNA isolation and 16S rRNA

gene amplification, and stored at -20 ºC until analysis. The picking and transferring

procedure was done inside the glovebox and with the help of some anoxic medium

(same as used for the liquid cultivation of the sediment cultures) reduced with titanium

(III) citrate.

Material & Methods

2.6 ANALYTICAL METHODS

2.6.1 Analyses of trichlorobenzenes by gas chromatography

The concentrations of tri- and di-chlorobenzenes in sediment cultures were periodically

monitored. For that, 0.5 ml culture aliquots were taken with sterile plastic syringes and

needles through the culture septum after homogenization of deposited sediment

particles. Aliquots were transferred into a 20 ml headspace vial, mixed with 0.5 ml

aqueous 1 M NaCl solution to increase volatilization of organic compounds.

Subsequently, vials were sealed with Teflon-coated stoppers and aluminium crimps.

Vials were preconditioned for 30 min shaking at 70°C in a HP 7694 auto-sampler

(Agilent Technologies) prior to the automatic injection into the Gas Chromatograph

(GC) for determination of the concentration of chlorinated benzenes. The GC was a HP

6890 GC system (Agilent Technologies) equipped with a flame ionization detector

(FID). The capillary column was a HP 5 (30 m length, 0.32 mm diameter, 0.25 μm film

thickness). Helium was the carrier gas. The GC temperature program (‘DCB_TCB

method’) was as follows: initial temperature of 55°C for 1 min; increase at a rate of

10°C per min until 90°C was reached; increase at a rate of 6°C per min until 130°C was

reached; increase of 30°C per min until 220°C were reached. The injector and detector

temperatures were 250°C and nitrogen was the makeup gas, set at a flow of 20 ml min-1.

Standards of pure 1,2,3-TCB, 1,2-DCB and 1,3-DCB were prepared for peak

identification (based on the respective compound retention time) and quantitation. Five-

point standard curves were prepared by spiking different volumes of 5–100 μM 1,2,3-

TCB, and 1,3-DCB and 1,2-DCB in cultivation serum bottles filled with 50 and 45 ml

sterile anoxic medium, and were measured in triplicate. Standard curves were associated

to linear regression lines for calibration. Detection limits were 1 μM for

trichlorobenzenes and 5 μM for dichlorobenzenes.

2.6.2 Analyses of chlorophenols by gas chromatography

The concentration of 2-chlorophenol and 2,3-dichlorophenol in sediment cultures were

periodically monitored. For that, a derivatization process (i.e., acetylation) was applied

involving the addition of NaHCO3 and acetic acid prior to analysis. Thus, the 2-

chlorophenol and 2,3-dichlorophenol were derivatized to the esters 2-chlorophenyl

acetate and 2,3-dichlorophenyl acetate, respectively. A 0.5 ml sample from sediment

cultures was taken with sterile plastic syringes through the sediment culture septum.

The 0.5 ml sample was loaded in a 20 ml headspace vial, which already contained 5–

10 mg of NaHCO3. Immediately after, 5 µl of pure acetic acid was added, and the vial

Material & Methods

was closed with Teflon-coated stoppers and aluminium crimps. Samples were mixed by

shaking vials with the hand and were analysed with the same GC, and using the same

GC column and method as described for trichlorobenzenes. Vials were preconditioned

in an auto-sampler in the same way as for the trichlorobenzenes analyses. The GC

temperature program started at 50°C for 1 min followed by an increase of 30°C per min

until 150°C was reached; increase at a rate of 5°C per min until 180°C was reached;

increase of 30°C per min until 250°C were reached. The FID temperature was 250°C

and nitrogen was the makeup gas, set at a flow of 20 ml min-1.

2.6.3 Cell visualization by epifluorescence microscopy

Cells were visualized by epifluorescence microscopy after staining with SYBR Green I

(SYBR® Green I Nucleic Acid Gel Stain - 10,000 x concentrate in DMSO, Invitrogen).

SYBR Green I was diluted 1:100 using sterile TE-buffer (consisting of 10 mM Tris and

1 mM EDTA, at pH 7.2), aliquoted, and stored at -20ºC. The staining was performed for

10 min in the dark in a ratio of 20:1 (v/v), where 20 µl of a sample from a sediment

culture was mixed with 1 µl of diluted SYBR I Green solution (final concentration in

the sample of SYBR Green I was 1:2000). Then, a sample of 18 μl from the stained

sediment culture was loaded on an agarose-coated slide for cell immobilization, and

subsequently covered with a cover glass. The agarose-coated slides were prepared by

homogeneously covering a glass slide with ~2 ml of liquefied agarose solution and

dried overnight inside a clean bench (Adrian et al 2007). The agarose solution was

prepared with two grams of low-melting agarose (SeaPlaque®, Biozym Scientific)

solved in 120 ml water, stirred, and heated until liquefied. The agarose-coated slides

allowed cell immobilization in a defined focus level between the slide and the cover

glass (Adrian et al 2007). Fluorescence microscopy was performed by using a Nikon

Eclipse TE300 microscope associated to a Nikon DXM 1200F digital Camera.

2.7 MOLECULAR BIOLOGY METHODS

2.7.1 DNA isolation from sediments, cultures and colonies

DNA isolation from sediments and sediment cultures was done using the FastDNA Spin

Kit for Soil (MP Biochemicals) with a FastPrep instrument (FastPrep FP120; Savant

Instruments) following the recommendations of the manufacturer with the following

general modifications: the silica matrix settled for 30 min prior to removal of

supernatant, and samples were incubated for 15 min at 42°C prior to final DNA elution

to increase yields. DNA was aliquoted and stored at -20ºC.

Material & Methods

Further specific modifications, in particular groups of samples, were applied: i) for

sediments from the Baffin Bay, DNA was isolated from 0.86 g of sediment. For the

bead beating step, beads and 780 µl of sodium phosphate buffer were added. In the final

step, DNA was eluted in 100 µl DNase/pyrogen-free ultra-pure water supplied with the

kit. DNA was isolated in triplicate for each sediment sample. Triplicates were pooled

and stored at -20ºC; ii) for sediments from Chile and Århus, DNA were isolated from

0.5 g of sediment, and eluted in 30 µl DNase/pyrogen-free ultra-pure water; and iii) for

sediment cultures, 1 ml (eventually 1.5 ml for subcultures with lower biomass) was

sampled with sterile syringes after homogenization of the sediment culture by mixing

by hand. The extracted 1 ml sample was placed in a sterile Eppendorf tube and

centrifuged for 20 min at 13,200 rpm. The supernatant from the tube was discarded. The

pellet was re-suspended in sodium phosphate buffer supplied by the kit. DNA was

eluted in 30, 40, or 50 µl DNase/ pyrogen-free ultra-pure water supplied with the kit.

DNA isolation from colonies was performed using the Nucleospin Tissue Kit

(Macherey & Nagel). For that, a colony was picked from a deep-agarose dilution tube

and placed into an Eppendorf tube with some sterile medium. A total of 16 colonies

were selected for DNA isolation. Additionally, DNA was isolated from samples (1 ml)

taken from sub-cultured colonies, which were incubated over 3–6 months. A total of 16

sub-cultured colonies were selected for DNA isolation. For both cases, samples were

centrifuged for 15–20 min at 10,400 rpm, and the supernatant was removed. In the case

that no visible pellet could be observed, a volume of 0.2 ml was left in the Eppendorf

tube. Pellets were re-suspended with Buffer T1 (180 µl) and Proteinase K (25 µl),

previously solved in 3.35 ml of Buffer PB, and mixed; all supplied by the manufacturer.

The re-suspended pellets were then incubated at 56ºC for 2 h in a heat-block at 1,100

rpm, and the following steps were done as recommended by the manufacturer. DNA

was eluted in 30 µl of elution buffer BE, supplied by the manufacturer. For colonies

transferred in a medium with humic acids, a second wash step with Buffer B5 was

repeated to further remove humic acids, and the eluted DNA was further diluted, to

avoid interference of humic acids in downstream processes, i.e., PCR.

2.7.2 Determination of DNA concentration

The DNA concentration was determined by measuring the absorbance at 260 nm using

a Nanodrop spectrophotometer (NanoDrop ND 1000, NanoDrop Technologies). The

ratio of absorbance 260/280 was used to evaluate the purity of the DNA in the samples,

considering values of ~1.8 as pure DNA.

Material & Methods

The concentration of purified amplicons amplified for further high-throughput

sequencing (either 454-pyrosequencing or Illumina) were determined for each sample

using a Quant-iT™ PicoGreen® dsDNA Assay Kit (Invitrogen), and from these

concentrations, the samples were pooled in equimolar amounts for all samples derived

from each of the primer sets, i.e., bacterial and universal primer sets. These two pools

were then mixed, so 71% of the total amplified sequences were derived from bacterial

primers, and 29% of the total amplified sequences were derived from universal primers,

in a final tube containing all amplicons prior to the 454-pyrosequencing. For the

Illumina sequencing, bacterial amplicons were mixed with archaeal amplicons (archaeal

data and information is not shown in this study; for further information see (Algora et al

2015)). The final ratio of amplified sequences was 62.5% bacterial and 37.5% archaeal

in a final tube.

2.7.3 Quantification of the 16S rRNA gene by qPCR

For qPCR analysis, 1 µl of DNA template was used. For the case of sediment samples,

the DNA template was ten-fold diluted to avoid PCR inhibition by co-extracted organic

substances, such as humic acids, coming from the sediment sample.

All the qPCR assays were performed on triplicate samples using a StepOne detection

system (StepOne/ StepOnePlus version 2.0, Applied Biosystems) and analysed with the

StepOne v2.1 software. Resulting amplicons from the qPCR assay were checked by

agarose gel electrophoresis when necessary, e.g., when melt-curve irregularities were

detected.

Amplification by qPCR of various microbial groups

For amplification of the 16S rRNA gene from the various targeted microbial groups,

i.e., total Bacteria, class Dehalococcoidia, order Desulfuromonadales, genera

Desulfitobacterium spp. and Dehalobacter spp., and total Archaea, various primer

combinations were used as specified in Table 4.

For the quantification of Bacteria and Dehalococcoidia 16S rRNA gene copy numbers,

the qPCR reaction mix had a final reaction volume of 20 µl. The qPCR reaction mix

components were 10 µl of SensiMix SYBR (Bioline) (including DNA polymerase,

dNTPs, SYBR Green I dye, stabilizers and the ROX reference dye), 7 µl of PCR-grade

water, and each primer to a final concentration of 1 µM.

Material & Methods

Table 4. Combination of primers used to amplify and quantify 16S rRNA genes of the various microbial

groups by qPCR.

Targeted microbial group Set of primers* Chemistry

Total Bacteria (variable region v3) 341f and 534r SYBR Green I

Total Archaea Arch349F and Arch806R, with the TaqMan probe Arch516F TaqMan

Class Dehalococcoidia DEH-Fa and DEH-R¹ SYBR Green I

Class Dehalococcoidia DehalF5 and DehalR² SYBR Green I

Order Desulfuromonadales GEO494F and GEO825R SYBR Green I

Genus Dehalobacter Dre441F and Dre645R_Ch SYBR Green I

Genus Desulfitobacterium DSB406F and DSB619R SYBR Green I

*Primer sequences and references are specified in Table 7.

¹Used for samples of sediments and sediment cultures from Baffin Bay and Chile.

²Used for samples of sediment cultures from Chile, Ireland, and Århus sediments (Table 1). Primers were designed based on

Dehalococcoidia 16S rRNA gene sequences retrieved from sediments of Chile, site 7155, after a clone library, and using the ‘probe

design’ function of the ARB software package (http://www.arb-home.de/) (Ludwig et al 2004), together with the SILVA database

(Pruesse et al 2007) release ‘SILVA 100’. Primers were designed by a collaborator, Dr. Kenneth Wasmund, UFZ–Leipzig.

The PCR program for both Bacteria and Dehalococcoidia (DEH-Fa and DEH-R) sets of

primers included a touchdown program as described in Table 5. Once amplification was

completed, a melt-curve was run to check the specificity of products. The melt-curve

had the following parameters: initial denaturation for 15 min at 95°C, renaturation for

1 min at 60°C, which was followed by a gradient (collecting fluorescent signal data

every 0.3°C) until 95°C was reached, and finally followed by 15 min at 95°C, as

described (Algora et al 2013, Wasmund et al 2015).

Table 5. PCR touchdown program for qPCR amplification of Bacteria and Dehalococcoidia (primer set

DEH-Fa and DEH-R).

Step Temperature time

Initial denaturation 95ºC 15 min

Number of cycles

Step

1 1 1 1 1 33

T time T time T time T time T time T time

Denaturation 95ºC 30 s 95ºC 30 s 95ºC 30 s 95ºC 30 s 95ºC 30 s 95ºC 30 s

Annealing 65ºC 30 s 64.6ºC 30 s 64.2ºC 30 s 63.8ºC 30 s 63.4ºC 30 s 63ºC 30 s

Elongation* 72ºC 45 s 72ºC 45 s 72ºC 45 s 72ºC 45 s 72ºC 45 s 72ºC 45 s

* Fluorescence was acquired during each elongation step

Material & Methods

The PCR program for Dehalococcoidia using the primers DehalF5 and DehalR is

described in Table 6. Melt-curve parameters were the same as for the bacterial and

DEH-Fa and DEH-R set of primers.

Table 6. PCR touchdown program for qPCR amplification of Dehalococcoidia (primer set DehalF5 and

DehalR).

Step Temperature time

Initial denaturation 95ºC 15 min

Number of cycles

Step

1 1 1 1 1 1

T time T Time T time T time T time T time

Denaturation 95ºC 30s 95ºC 30s 95ºC 30s 95ºC 30 s 95ºC 30s 95ºC 30s

Annealing 66ºC 30s 64.8ºC 1 min 63.6ºC 30s 62.4ºC 30 s 62.2ºC 30s 61.2ºC 30s

Elongation* 72ºC 40s 72ºC 40s 72ºC 40s 72ºC 40 s 72ºC 40s 72ºC 40s

Followed by 35 cycles at

Step Temperature time

Denaturation 95ºC 30 s

Annealing 60ºC 30 s

Elongation* 72ºC 40 s

* Fluorescence was acquired during each elongation step

The qPCR assay for quantification of total Archaea and order Desulfuromonadales 16S

rRNA gene copy numbers in sediment samples from the Baffin Bay was performed with

the primers described in Table 7, and as previously reported (Algora et al 2013, Holmes

et al 2002, Schippers and Neretin 2006, Takai and Horikoshi 2000). The quantification

of Dehalobacter restrictus and Desulfitobacterium spp. in sediment cultures from Chile,

site 7155, was performed as previously described (Smits et al 2004) (Table 7). The

reverse primer, Dre645R, targeting the Dehalobacter 16S rRNA gene was modified (see

Table 7) accordingly to Dehalobacter sequences found in Chile sediments after a clone

library of the 16S rRNA gene was set up, amplified with the primers 27f and 1492r

(Table 7). The clone library and the Dre645R primer modification were performed by a

collaborator, Dr. Kenneth Wasmund, UFZ–Leipzig.

Material & Methods

Conversion of qPCR data to gene copy numbers ml-1culture or g-1 sediment

For the determination of the number of 16S rRNA gene copies for total Bacteria and

Dehalococcoidia in the qPCR assays, a DNA standard was prepared after cloning 16S

rRNA genes into a pGEM-T vector. The 16S rRNA gene used came from a

Dehalococcoidia member amplified using the primers 27f and 1492r (Table 7), from

sediment samples of Chile as described (Wasmund et al 2015).

For the determination of DNA concentration (ng μl-1) in the DNA standard, a NanoDrop

ND 1000 (NanoDrop Technologies) was used as previously described (section 2.7.2).

The DNA concentration was measured in triplicate, and converted to 16S rRNA gene

copies μl-1 with the formula in Equation 1.

16𝑆𝑆 𝑟𝑟𝑟𝑟𝑟𝑟𝑟𝑟 𝑔𝑔𝑔𝑔𝑔𝑔𝑔𝑔 𝑐𝑐𝑐𝑐𝑐𝑐𝑐𝑐𝑔𝑔𝑐𝑐 𝜇𝜇𝜇𝜇−1

= {[𝐷𝐷𝑟𝑟𝑟𝑟](𝑔𝑔𝑔𝑔 𝜇𝜇𝜇𝜇−1)} x {6.022 𝑥𝑥 1023(𝑚𝑚𝑐𝑐𝜇𝜇𝑔𝑔𝑐𝑐𝑚𝑚𝜇𝜇𝑔𝑔𝑐𝑐 𝑚𝑚𝑐𝑐𝜇𝜇𝑔𝑔−1)}

𝜇𝜇𝑔𝑔𝑔𝑔𝑔𝑔𝑙𝑙ℎ (𝑏𝑏𝑐𝑐)𝑥𝑥 109(𝑔𝑔𝑔𝑔 𝑔𝑔−1) 𝑥𝑥 650 (𝑔𝑔 𝑚𝑚𝑐𝑐𝜇𝜇𝑔𝑔 𝑐𝑐𝑜𝑜 𝑏𝑏𝑐𝑐−1)

Equation 1. Formula used for the conversion of DNA concentration (ng μl-1) of a specific DNA strand of

a given length (bp) to the number of DNA copies μl-1.The formula assumes an average weight of one bp

to be 650 Dalton, and thus, one bp mole to weight 650 g. The molecular weight of a DNA strand is

estimated by multiplying 650 times the bp length. 6.022 x 1023 is Avogadro’s number

(http://cels.uri.edu/gsc/cndna.html, and (Ritalahti et al 2006)).

Subsequently, the calculated number of 16S rRNA gene copies in the DNA standard

was brought to 1011 copies µl-1 by dilution with molecular-grade water, which was used

as a quantified stock solution, aliquoted, ten-fold diluted, and stored at -80ºC (Wasmund

et al 2015). A standard curve was either freshly made after thawing an aliquot of the

quantified stock solution, or by thawing ten-fold diluted standard aliquots (107–102)

from a -80ºC stock. In each qPCR assay, a standard curve ranging from 107 to 102 16S

rRNA gene copies µl-1 was always included in triplicate. The standard curve was used

for i) the calculation of qPCR amplification efficiency after association to a linear

regression line, and ii) conversion from the qPCR threshold cycle values (Ct; defined as

the qPCR cycle number where the intensity from the fluorescence reaches a set

threshold (Ritalahti et al 2006)) to gene copies µl-1 in the qPCR reaction. The

amplification efficiency was calculated from the calibration curve slope according to the

formula 10(-1/slope). Each qPCR run was checked for amplification efficiencies, and only

those qPCR runs which ranged between 90 and 110% were further processed. The

conversion of Ct values to 16S rRNA gene copy numbers µl-1 in each qPCR reaction for

each sample was performed using the standard curve, and calculated by the StepOne

v2.1 software (Applied Biosystems). Further conversion of 16S rRNA gene copy

Material & Methods

numbers µl-1 per reaction to 16S rRNA gene copies ml-1 or g-1 of sample was calculated

as previously described (Ritalahti et al 2006), and according to Equation 2.

𝑔𝑔𝑔𝑔𝑔𝑔𝑔𝑔 𝑐𝑐𝑐𝑐𝑐𝑐𝑐𝑐𝑔𝑔𝑐𝑐 𝑚𝑚𝜇𝜇−1𝑐𝑐𝑟𝑟 𝑔𝑔−1 𝑐𝑐𝑜𝑜 𝑐𝑐𝑠𝑠𝑚𝑚𝑐𝑐𝜇𝜇𝑔𝑔

= (𝑔𝑔𝑔𝑔𝑔𝑔𝑔𝑔 𝑐𝑐𝑐𝑐𝑐𝑐𝑐𝑐𝑔𝑔𝑐𝑐 𝑐𝑐𝑔𝑔𝑟𝑟 𝑟𝑟𝑔𝑔𝑠𝑠𝑐𝑐𝑙𝑙𝑐𝑐𝑐𝑐𝑔𝑔) 𝑥𝑥 �𝑔𝑔𝜇𝜇𝑚𝑚𝑙𝑙𝑔𝑔𝑒𝑒 𝑙𝑙𝑐𝑐𝑙𝑙𝑠𝑠𝜇𝜇 𝑣𝑣𝑐𝑐𝜇𝜇𝑚𝑚𝑚𝑚𝑔𝑔 𝑐𝑐𝑜𝑜 𝐷𝐷𝑟𝑟𝑟𝑟 𝑐𝑐𝑔𝑔 𝑙𝑙ℎ𝑔𝑔 𝐷𝐷𝑟𝑟𝑟𝑟 𝑐𝑐𝑐𝑐𝑐𝑐𝜇𝜇𝑠𝑠𝑙𝑙𝑐𝑐𝑐𝑐𝑔𝑔 (𝜇𝜇𝜇𝜇)�

�𝑣𝑣𝑐𝑐𝜇𝜇𝑚𝑚𝑚𝑚𝑔𝑔 𝑐𝑐𝑜𝑜 𝐷𝐷𝑟𝑟𝑟𝑟 𝑠𝑠𝑒𝑒𝑒𝑒𝑔𝑔𝑒𝑒 𝑐𝑐𝑔𝑔𝑟𝑟 𝑟𝑟𝑔𝑔𝑠𝑠𝑐𝑐𝑙𝑙𝑐𝑐𝑐𝑐𝑔𝑔 (𝜇𝜇𝜇𝜇)� 𝑥𝑥 �𝑣𝑣𝑐𝑐𝜇𝜇𝑚𝑚𝑚𝑚𝑔𝑔 𝑐𝑐𝑟𝑟 𝑤𝑤𝑔𝑔𝑐𝑐𝑔𝑔ℎ𝑙𝑙 𝑐𝑐𝑜𝑜 𝑐𝑐𝑠𝑠𝑚𝑚𝑐𝑐𝜇𝜇𝑔𝑔 (𝑚𝑚𝜇𝜇 𝑐𝑐𝑟𝑟 𝑔𝑔)�

Equation 2. Formula for the conversion of gene copies per reaction to gene copies per ml or g of sample.

The number of gene copies per reaction is given by the qPCR assay after using the standard curve. The

eluted volume of DNA within each DNA isolation from each sample, was either 30, 40, or 50 µl for

sediment cultures, or 100 µl for sediments. The volume of DNA added per reaction was 1 µl. The volume

of sediment culture sample from which the DNA was isolated was either 1 ml or 1.5 ml, and the weight of

sediment sample was 0.5 or 0.86 g.

Standard curves for the determination of Archaea and the order Desulfuromonadales

were done as described (Schippers and Neretin 2006).

2.7.4 Quantification of the functional genes mcrA and dsrA by qPCR

The quantitative amplification of the functional genes of the dsrA, which encodes the

dissimilatory sulphite reductase of sulphite/sulphate reducers and the mcrA encoding for

the methyl coenzyme M reductase subunit α gene of methanogens/anaerobic

methanotrophs was carried out as described (Algora et al 2013).

All primers used in this study for qPCR assays are presented in Table 7.

Table 7. Primers used in this study.

Primer

name

Sequence (5’-3’)

Target

gene

Target group

Reference

27F

AGAGTTTGATCMTGGCTCAG

16S rRNA

Bacteria

(Weisburg et al 1991)

1492R

GGTTACCTTGTTACGACTT

16S rRNA

Bacteria

(Lane 1991)

519R

TATTACCGCGGCKGCTG

16S rRNA

Bacteria

(Lane et al 1985)

341F

CCTACGGGAGGCAGCAG

16S rRNA

Bacteria

(Muyzer et al 1993)

534R

ATTACCGCGGCTGCTGGCA

16S rRNA

Bacteria

(Muyzer et al 1993)

343F

TACGGRAGGCAGCAG

16S rRNA

Bacteria

(Liu et al 2007)

534R

ATTACCGCGGCTGCTGGC

16S rRNA

Bacteria

(Liu et al 2007)

DEH-Fa

TACGGGAGGCAGCAGCDA

16S rRNA

Dehalococcoidia

(Wasmund et al 2015)

DEH-R

GRRAGGGTCGATACYCC

16S rRNA

Dehalococcoidia

(Wasmund et al 2015)

Dehal-F5

ATCTCYCRGCTYAACYGGGA

16S rRNA

Dehalococcoidia

Chile sediments

This study. Designed by

Wasmund

Dehal-R

ARRAGGGTCGATACYCC

16S rRNA

Dehalococcoidia

Chile sediments

This study. Designed by

Wasmund

GEO494F

AGGAAGCACCGGCTAACTCC

16S rRNA

Desulfuromonadales

(Holmes et al 2002)

GEO825R

TACCCGCRACACCTAGT

16S rRNA

Desulfuromonadales

(Anderson et al 1998)

Arch349F

GYGCASCAGKCGMGAAW

16S rRNA

Archaea

(Takai and Horikoshi

2000)

Arch806R

GGACTACVSGGGTATCTAAT

16S rRNA

Archaea

(Takai and Horikoshi

2000)

Arch516F

TGYCAGCCGCCGCGGTAAHA

CCVGC

16S rRNA

Archaea

(Takai and Horikoshi

2000)

U789F

TAGATACCCSSGTAGTCC

16S rRNA

Prokaryotes

(Baker et al 2003, Barns

et al 1994)

U1068R

CTGACGRCRGCCATGC

16S rRNA

Prokaryotes

(Lee et al 2011)

Material & Methods

Primer

name

Sequence (5’-3’)

Target

gene

Target group

Reference

ME1F

GCMATGCARATHGGWATGTC

mcrA

methanogens

(Hales et al 1996)

ME3R

TGTGTGAASCCKACDCCACC

mcrA

methanogens

(Wilms et al 2007)

DSR1F+

ACSCACTGGAAGCACGGCGG

dsrA

sulphate reducers

(Kondo et al 2004)

DSR-R

GTGGMRCCGTGCAKRTTGG

dsrA

sulphate reducers

(Kondo et al 2004)

Dre441F

GTTAGGGAAGAACGGCATCT

16S rRNA

Dehalobacter restrictus

(Smits et al 2004)

Dre645R_

CCTCTCCTGTCCTCAAGCCAH

16S rRNA

Dehalobacter restrictus

This study. (Smits et al

2004), modified by

Wasmund

DSB406F

GTACGACGAAGGCCTTCGGG

16S rRNA

Desulfitobacterium

(Smits et al 2004)

DSB619R

CCCAGGGTTGAGCCCTAGGT

16S rRNA

Desulfitobacterium

(Smits et al 2004)

RRF2

SHMGBMGWGATTTYATGAA

rdh-genes

Dehalococcoides mccartyi

(Krajmalnik-Brown et al

2004)

B1R

CHADHAGCCAYTCRTACCA

rdh-genes

Dehalococcoides mccartyi

(Krajmalnik-Brown et al

2004)

RDH F1C

TTYMVIGAYITIGAYGA

rdh-genes

Dehalococcoides mccartyi

(Chow et al 2010)

RDH R1C

CCIRMRTYIRYIGG

rdh-genes

Dehalococcoides mccartyi

(Chow et al 2010)

dehaloF3

ATCGWTSMRGGTAT

rdh-genes

Dehalobacter restrictus and

Desulfitobacterium spp.

(von Wintzingerode et al

2001)

dehaloR2

TYTGTACCATAGCC

rdh-genes

Dehalobacter restrictus and

Desulfitobacterium spp.

(von Wintzingerode et al

2001)

dehaloF5

GGTTGCATTGCYGTCAT

rdh-genes

Dehalobacter restrictus and

Desulfitobacterium spp.

(von Wintzingerode et al

2001)

dehaloR4

TGCTTYATGGAACCAGG

rdh-genes

Dehalobacter restrictus and

Desulfitobacterium spp.

(von Wintzingerode et al

2001)

M13F

GTAAAACGACGGCCAGT

pGEM –T vector

M13R

GCGGATAACAATTTCACACA

pGEM –T vector

IUPAC-Code: A – adenine; B – cytosine/ guanine/ thymine; C – cytosine; D – adenine/guanine/ thymine; G – guanine; H –

adenine/cytosine/ thymine; I – inosine; K – guanine/ thymine; M – adenine/cytosine; R – adenine/ guanine; S – cytosine/

guanine; T – thymine; V – adenine/ cytosine/ guanine; W – adenine/ thymine; Y – cytosine/ thymine

2.7.5 Amplification of reductive dehalogenase genes

Several sets of primers were used for amplification of the functional gene of the key

enzyme for organohalide respiration, i.e., the reductive dehalogenase (RDase).

Amplification of reductive dehalogenase homologous genes (rdh-genes) in sediment

cultures was performed with four sets of primers. Two sets were designed for targeting

the rdh-genes from Dehalococcoides mccartyi (Chow et al 2010, Krajmalnik-Brown et

al 2004) and the other two set was designed for targeting the rdh-genes from

Dehalobacter restrictus and Desulfitobacterium spp. (von Wintzingerode et al 2001).

The first set of primers, RRF2 and B1R (Table 7), were specific for Dehalococcoides

mccartyi and were used as described (Krajmalnik-Brown et al 2004). In addition, the

primers RDH F1C and RDH R1C (Table 7) were used with a S-Tbr DNA polymerase

(DyNAmo II, Finnzymes) that is specific for short primers (Isenbarger et al 2008). The

reaction mix contained 2.5 µl of buffer and 0.7 DNA polymerase (DyNaMo Taq), 2.5 µl

of dNTPs (0.2 mM; final concentration in PCR reaction mix), 1 µl of each primer

(0.8 µM; final concentration in PCR reaction mix), 0.75 µl of MgCl2 (1.5 mM; final

concentration in PCR reaction mix), 0.1 µl BSA (0.2 g l-1 final concentration PCR

reaction mix; New England Biolabs), 1 µl of DNA template, and molecular grade water

Material & Methods

up to 25 µl. The PCR program was the following: initial denaturation at 95ºC for 2 min,

44 cycles of denaturation at 95 ºC for 30 s, annealing at 47 ºC for 30 s, elongation at

72ºC for 1 min 20 s, and a final elongation step of 72ºC for 5 min.

For the amplification of rdh-genes described for Dehalobacter restrictus and

Desulfitobacterium spp., and dechlorinating mixed cultures, the pair of degenerate

primers dehaloF3 together with dehaloR2, and a second pair of primers dehaloF5

together with dehaloR4 were used (Table 7). The PCR program for both primer sets

involved the following parameters: initial denaturation at 95ºC for 5 min, 35 cycles of

denaturation at 95ºC for 30 s, annealing at 50ºC for 30 s, and elongation at 72ºC for

60 s, and a final elongation step at 72ºC for 7 min. Individual PCR reactions had a total

volume of 25 µl, and contained 12.5 µl of master mix (Qiagen), which already

contained DNA polymerase, buffer and dNTPs (https://www.qiagen.com/de/

products/catalog/assay-technologies/end-point-pcr-and-rt-pcr-reagents/taq-pcr-master-

mix-kit), 1.5 µl of each primer (0.6 µM; final concentration in PCR reaction mix),

1.5 µl of MgCl2 (1.5 mM; final concentration in PCR reaction mix), 0.1 µl BSA

(0.2 g l-1 final concentration PCR reaction mix; New England Biolabs), 6.9 µl of

molecular grade water, and 1 µl of DNA template.

2.7.6 Amplification of 16S rRNA gene from colonies

For PCR amplification of nearly entire-length bacterial 16S rRNA genes from the

various colonies, the primers 27F and 1492R (Table 7) were used. Single PCR reactions

were performed using a Taq PCR Master Mix Kit (Qiagen), and consisted of 12.5 µl of

master mix (Qiagen), which already contained DNA polymerase, buffer and dNTPs,

both primers (0.2 µM final concentration in PCR reaction mix), BSA (0.08 g l-1 final

concentration in PCR reaction mix; Promega), 1 µl of DNA template, and molecular

grade water up to 25 µl.

The PCR program consisted of an initial denaturation of 95°C for 4 min, followed by 35

cycles of denaturation at 95°C for 30 s, annealing at 55°C for 2 min, elongation at 72°C

for 3 min, and a final elongation step of 72°C for 10 min.

2.7.7 Amplification of 16S rRNA gene for 454-pyrosequencing

Bacterial 16S rRNA genes amplification was done using the primers: 27F and 519R

(Table 7), targeting the V1–V3 hypervariable regions of the 16S rRNA gene, and the

universal primers U789F and U1068R for both Bacteria and Archaea targeting the

hypervariable V6 region of the 16S rRNA gene (Baker et al 2003, Lee et al 2011, Wang

and Qian 2009).

Material & Methods

Primers were extended with barcoded multiplex identifiers (MID), which were

sequences of 10 bp specific to each sample (added to both forward and reverse primer

sequences, except for the universal reverse primer which had no MID sequence). The

MIDs allowed the later determination of the origin of each sequence from pooled

sequencing libraries. In addition, all primers were extended with the library “key”

TCAG sequence next to the barcoded MID sequence, or to the primer in case of the

universal reverse primer, and followed by the adapter A

(CGTATCGCCTCCCTCGCGCCA) in the forward primers, and adapter B

(CTATGCGCCTTGCCAGCCCGC) in the reverse primers. The design of these fusion

primers was done as requested by 454/Roche (http://454.com/products-

solutions/experimental-design-options/amplicon-sequencing. asp). Total amplicon

lengths were 599 bp for bacterial primers and 373 bp for universal primers.

PCR single reactions contained 10 μl of 2x Phusion Flash High-Fidelity Buffer

(Finnzymes), which already contained the Phusion® High-Fidelity DNA Polymerase

(Finnzymes) and dNTPs, each primer in a final concentration of 5 µM, and 1.0 μl of

DNA template, and deionized molecular-grade water up to 20 μl.

Bacterial PCR thermocycling conditions comprised an initial denaturation of 98°C for

10 s, followed by 30 cycles of 98°C for 5 s, 52°C for 10 s and 72°C for 10 s. PCR

thermocycling conditions for PCRs using universal primers were the same as for PCR

reactions using bacterial primers, except the number of cycles was 35. PCR reactions

were done in five replicates for each sample. Amplicons from the same sample were

pooled

2.7.8 Amplification of the 16S rRNA gene for Illumina sequencing

For Illumina sequencing, 16S rRNA genes were amplified with the bacterial primer set

343F and 534R (Table 7) targeting the V3 hypervariable region of the 16S rRNA gene.

To enable multiplexed sequencing in a single Illumina run, forward primers were

extended at the 5´end with a ‘barcode’ sequence of 7 bp. The ‘barcode’ sequence was

specific for each sample and differed to the rest of the barcodes by at least two bases. In

between the primer and the barcode, the 2 bp “TA” linker was inserted in order to

reduce possible bias effects of the barcodes (Degnan and Ochman 2012, Vasileiadis et

al 2012, Wu et al 2010). Amplicon lengths were 190 bp.

Single PCR reactions contained 4 µl Phusion® High-Fidelity Buffer (Finnzymes),

200 μM of each dNTP, 1 µM of each primer, 0.5 U of Phusion® High Fidelity

Material & Methods

Polymerase (Finnzymes), 1 µl of DNA template, and molecular biology grade water up

to 20 µl.

Bacterial PCR conditions were: initial denaturation of 98°C for 30 s, followed by 35

cycles of a two-step cycling program of 98°C for 10 s and 72°C for 30 s, and a final

elongation of 72°C for 1 min. Triplicate PCR reactions per sample were performed, and

amplicons from each triplicate PCR reaction from each sample were pooled.

2.7.9 Agarose gel electrophoresis and amplicon purification

After PCR, amplicons were loaded on agarose gels, and gel electrophoresis was run to

check if successful amplification of the correct band size occurred. For that, agarose

gels were prepared by solving 1–1.5 g of agarose in 100 ml of 1 x TAE buffer (40 mM

Tris, 20 mM acetic acid, 1 mM EDTA, pH 8.3). Agarose gels with loaded amplicons

and a DNA ladder (GeneRulerTM 100 bp Plus DNA Ladder, Fermentas) were run for

30 min at 110 V, as a standard basis, and for 70 min at 90 V, when a better band size

separation was needed (i.e., when bands were excised from agarose gels). The running

buffer in the electrophoresis chamber was 1 x TAE. Then, staining of the agarose gel

with ethidium bromide was performed and amplicons were visualized under UV-light

with a UV-detector (BioRad Gel DocTM XR+Imager).

When needed, and prior to loading into agarose gels, amplicon purification from the

PCR reaction was done using a Wizard® SV Gel and PCR Clean-Up Kit (Promega)

following the recommendations from the manufacturer.

2.7.10 Elution of amplicons from agarose gels

The elution of amplicons from agarose gels were done prior to 454-pyrosequencing and

Illumina sequencing. After gel electrophoresis, bands of the correct size were excised

using sterile scalpels, placed in sterile Eppendorf tubes, weighed and purified using a

Wizard® SV Gel and PCR Clean-Up Kit (Promega) following the manufacturer’s

recommendations.

2.7.11 Cloning of 16S rRNA gene amplicons from Dehalococcoidia qPCR

amplifications

Selected amplicons amplified in a qPCR assay with primers targeting the 16S rRNA

gene of the class Dehalococcoidia were cloned in order to determine their phylogenetic

affiliation within this class.

Material & Methods

Purified amplicons were cloned into Escherichia coli using the pGEM-T vector system

(Promega). The ligation reaction was performed according to the manufacturer’s

instructions, and incubated for 1h at ambient temperature. The pGEM-T vector

containing the corresponding ligated amplicons was introduced in Escherichia coli

strain JM109 high-efficiency competent cells (Promega). The transformation protocol

provided by the manufacturer was followed to transform the JM109 strain of E.coli.

Transformed cells were plated in LB agar plates, supplemented with ampicillin

(100 μg ml-1), X-gal (80 μg ml-1) and IPTG (0.1 mM). Colonies were allowed to grow

on the plates overnight in an incubator at 30ºC. White colonies (containing the cloned

fragment) were picked using sterile toothpicks and transferred to a new LB agar plate

supplemented with ampicillin, IPTG and X-gal. Plates were incubated overnight at 30ºC

to allow colonies to grow. Part of each isolated white colony was transferred with a

sterile toothpick to a sterile Eppendorf tube containing 100 µl of molecular-grade water.

One Eppendorf tube was used for each colony. Colony cells inside the Eppendorf tube

were intensively vortexed to homogeneously dissolve them in the water. Eppendorf

tubes containing colony cells were incubated on the bench at ambient temperature for

2h. After this time, the colony dissolved in water was used as template for a colony

PCR reaction to confirm the presence of cloning fragments with the correct size.

PCR single reactions contained 3.5 µl of buffer (Fermentas), 3.5 µl dNTPs, 1 µl M13F

primer (Table 7), 1 µl M13R primer (Table 7), 2.8 µl MgCl2, 0.175 µl BSA (0.2 g l-1

final concentration PCR reaction mix; New England Biolabs), 0.175 µl Taq DNA

Polymerase (Fermentas), 1 µl of template, and sterile molecular-grade water up to 35 µl.

The PCR program consisted of an initial denaturation of 95ºC for 5 min, 40 cycles of

95ºC for 30 s, 55ºC for 30 s, and 72ºC for 40 s and a final elongation of 72ºC for 5 min.

2.7.12 454-pyrosequencing, Illumina sequencing and subsequent data analysis

The emulsion PCR and sequencing was done by means of the GS FLX Titanium

chemistry, following protocols from the manufacturer and using a 454 GS FLX

pyrosequencer (Roche), as recommended by the developer, and performed by a

collaborator, Dr. Richard Reinhardt and his group at the Max Planck Genome Centre in

Cologne.

Initial quality filtering and extraction of raw pyrosequencing reads was performed using

the ‘amplicon’ settings of the GS Run Processor (Roche). Additional quality control

processing included screening sequences using the Greengenes (DeSantis et al 2006)

‘Trim’ tool (http://greengenes.lbl.gov/cgi-bin/nph-trim_fasta_by_qual.cgi) with the

Material & Methods

following settings: good quality score=25, window size=25 bp, and window

threshold=90%. Following this, sequences with >1 mismatch to the barcode, >2

mismatches to the forward primer, <200 bp in length and containing homopolymers of

>10 bp were removed using mothur (v1.35.0) (Schloss et al 2009). Further, bases after

300 bp were removed from the 3’ ends of all reads. Chimeric sequences were then

detected using the ‘chimera.uchime’ command within mothur (Edgar et al 2011), using

the silva.gold.alignment file (supplied by the mothur website) as a reference alignment.

Potential chimeric sequences were then removed. Taxonomic classification of sequence

reads was performed using ‘classify.seqs’ command within mothur with a bootstrap cut-

off of 50% and using the SILVA taxonomy files (v119) supplied by the mothur website

(http://www.mothur.org/wiki/Silva_reference_files). The classify.seqs command is the

mothur implementation of the RDP naïve Bayesian rRNA Classifier (Wang and Qian

2009). The analysis of the 454-pyrosequencing data was performed in collaboration

with Dr. Kenneth Wasmund, UFZ–Leipzig.

The Illumina sequencing was performed by using an Illumina Genome Analyzer IIx

with the TruSeq SBS Kit v5-GA sequencing reagents (Illumina Inc) by our

collaborators Dr. Edoardo Puglisi and Dr. Sotirios Vasileiadis from the Universita

Cattolica del Sacro Cuore, at Piacenza, Italy as described (Algora et al 2015). The

subsequent bioinformatics sequence data analysis was performed by a collaborator Dr.

Sotirios Vasileiadis, using mothur v1.28 (Schloss et al 2009) together with the statistical

analyses (Spearman’s rank correlation and Hierarchical clustering with the UPGMA

algorithm), which were performed in R (R Development Core Team 2011) as described

(Algora et al 2015).

2.7.13 Sanger sequencing and subsequent data analysis

Sanger sequencing of 16S rRNA gene amplicons was performed with the 27F primer

using an ABI Prism® 3139xl genetic analyzer instrument (Applied Biosystems) at the

Environmental Microbiology Department of the UFZ–Leipzig.

Prior to sequencing, a PCR reaction with the BigDye terminator v3.1 cycle sequencing

kit (Applied Biosystems) was performed. Single reactions contained BigDye (1 µl),

primer 27F (1 µl; Table 7), BigDye Terminator 5 x sequencing Buffer (1 µl), molecular-

grade sequencing water (4–6 µl) and amplicons (1–3 µl). The PCR program was 25

cycles of 96ºC for 30 s, 55ºC for 15 s, 60ºC for 4 min. Afterwards, amplicons were

precipitated with ethanol. For that, 10 µl of amplicons were mixed with 10 µl sodium

acetate, (3 M, pH 4.8), 150 µl ethanol (100%) and 80 µl molecular-grade water, and

Material & Methods

centrifuged for 10 min at 12,000 rpm. The supernatant was discarded, and 300 µl

ethanol (70%) was added, and subsequently centrifuged. The supernatant was discarded

and the pellet was vacuum-dried before injected to the sequencer.

The sequences obtained were analysed for their quality with the software Chromas Lite

(Technelysium Pty Ltd). Sequences were compared to public databases using the Blast

tool (Basic Local Alignment Search Tool; (Altschul et al 1990)) and the NCBI database

(http://www.ncbi.nlm.nih.gov/blast/).

Results

3 RESULTS

3.1 APPROACHES TO CULTIVATE DEHALOCOCCOIDIA FROM VARIOUS

MARINE SEDIMENTS

Quantification of Dehalococcoidia was performed by qPCR using the primers DehalF5

and DehalR, which are specific for the class Dehalococcoidia, and target the 16S rRNA

gene (Table 7). 16S rRNA gene copy numbers of Dehalococcoidia were used as a proxy

to monitor Dehalococcoidia growth with time, and thus investigate if Dehalococcoidia

could be cultivated in the laboratory at atmospheric pressures and to which time scale,

i.e., within a timespan of days, weeks, months, or years, they may grow. Additionally,

16S rRNA gene copy numbers of Dehalococcoidia were compared among the various

sediment cultures supplied with different potential electron acceptors to investigate

which potential electron acceptor may promote Dehalococcoidia growth. Total bacterial

numbers were additionally monitored with a 16S rRNA gene-targeted qPCR assay.

3.1.1 Dehalococcoidia abundance under various terminal electron acceptors in

sediment cultures

Dehalococcoidia were detected with the qPCR assay in all sediment cultures. The

sediment cultures with highest Dehalococcoidia numbers were those inoculated with

sediments of the Århus core at depths of 430–440 cmbsf, with values between 104–

105 16S rRNA gene copies ml-1 culture. Cultures set up with sediments from site 1317

of Ireland also showed values of 104 16S rRNA gene copies ml-1. The rest of cultures

showed values of ~103 16S rRNA gene copies ml-1, as it was the case for cultures

inoculated with sediments from the site 1318 of Ireland and Chile, sites 7155 and 7165.

Values lower than 103 16S rRNA gene copies ml-1 could not be precisely quantified as

they were out of range of the calibration curve.

In order to study if Dehalococcoidia numbers increased in the sediment cultures with

time, a culture was specifically monitored from start of the incubation and after 68 days.

This culture was inoculated (13%, v/v) with a sediment culture that had been previously

set up with sediments from Chile site 7155, and that dehalogenated 1,2,3-

trichlorobenzene (as further detailed in section 3.2). 16S rRNA gene copy numbers of

Dehalococcoidia increased one order of magnitude, from a number of 687 16S rRNA

gene copies ml-1 at the start point to a number of 2.1 x 104 after 68 days. Moreover,

nearly no change was detected in total bacterial 16S rRNA gene copy numbers with

time, from a 16S rRNA gene copy number of 3.8 x 107 at the start, to a number of

2.7 x 107 after 68 days of incubation (Figure 13). The relative proportion of

Results

Dehalococcoidia with respect to the total bacterial community, were of 0.002% at the

start of the incubation time, which increased to 0.08% after 68 days of incubation

Figure 13. 16S rRNA gene copy numbers of Dehalococcoidia (“DEH”) and total Bacteria (“Tot Bact”)

after 3 and 68 days of incubation, in a culture inoculated with a previously established sediment culture

set up with sediments from Chile, site 7155. Shown are standard deviations of triplicate measurements in

the qPCR assay.

Cloning and sequencing of the Dehalococcoidia qPCR amplicons revealed a

phylogenetical affiliation of the twelve cloned sequences to the class Dehalococcoidia

(Figure 14). These cloned Dehalococcoidia sequences did not fall into the clade Ord-

DEH, which includes the organohalide-respiring characterized Dehalococcoidia, but

were affiliated with the Dehalococcoidia-sister clades, particularly DSC-D (the name of

the clades of Dehalococcoidia are indicated as described by (Wasmund et al 2015)).

Results

Figure 14. Neighbour joining tree based on partial 16S rRNA sequences done with the SILVA 100 database and the ARB program. Shown in red are the cloned sequences

from the amplicons amplified with qPCR from a sediment culture inoculated with sediments of Chile site 7155. Shown in green are cultured strains.

Results

Dehalococcoidia abundance among the various tested electron acceptors showed

altogether no clear preferential growth under any of the tested electron acceptors, either

halogenated or non-halogenated, i.e., 16S rRNA gene copies ml-1 varied no more than

one order of magnitude at a given time point. However, differences in Dehalococcoidia

abundance were observed in the various tested sediments.

For cultures inoculated with Århus sediments from depths of 410-420 cmbsf,

Dehalococcoidia copy numbers were highest in cultures containing sulphate and 2,4,6-

tribromoresorcinol (upper panel, Figure 15). In addition, Dehalococcoidia copy

numbers were stable after nine months with a value of 104 16S rRNA gene copies ml-1

in cultures containing sulphate. Dehalococcoidia copy numbers could not be measured

in the time point of nine months of incubation in cultures amended with 2,4,6-

tribromoresorcinol.

In sediment cultures inoculated with sediments from Århus at a depth of 430-440 cmbsf,

Dehalococcoidia copy numbers increased with time exclusively in those cultures

supplied with sulphate and 2,4,6-tribromoresorcinol (lower panel, Figure 15).

Dehalococcoidia copy numbers of 1.4 x 104 16S rRNA gene copies ml-1 culture were

observed in sediment cultures amended with sulphate, which increased an order of

magnitude to 2.6 x 105 copies ml-1 after nine months of cultivation. Similarly,

Dehalococcoidia numbers increased one order of magnitude, from 1.3 x 104 to 1.4 x 105

16S rRNA gene copies ml-1 culture, in sediment cultures amended with 2,4,6-

tribromoresorcinol after an incubation time of nine months.

No substantial differences were detected in Dehalococcoidia numbers in the presence of

various reducing agents: either the inorganic reducing agent sodium sulphide (sediment

cultures supplied with 2-chlorophenol and TCB were reduced with sodium sulphide), or

the organic compound (titanium III citrate), which may be fermented.

Results

Figure 15. Dehalococcoidia quantification of sediment cultures set up with sediments from Århus Bay,

measured as 16S rRNA gene copy numbers by qPCR. In the upper panel, cultures inoculated with the top

sediment layer from Århus Bay (410-420 cmbsf) are shown. The lower panel shows cultures from the

deeper sediment layer from Århus Bay (430-440 cmbsf). Standard deviations refer to triplicate

measurements of the qPCR assay. Abbreviations: SO4 - sulphate; TCB - 1,2,3- and 1,2,4-trichlorobenzene

mixed in a 1:1 proportion; DCP - 2,3-dichlorophenol; CP - 2-chlorophenol; PCE - tetrachloroethene;

BMBA - 4-bromo-3,5-dimethoxybenzoic acid; TBR - 2,4,6-tribromoresorcinol; IMB - 1-iodo-2,6-

dimethoxybenzene. TCB and CP sediment cultures were reduced with Na2S, all others with titanium (III)

citrate.

Results

Differences of two orders of magnitude in Dehalococcoidia copy numbers were found

between the cultures set up with sediments from the two sites of the Porcupine Seabight

(Ireland; Table 1). Site 1318 had Dehalococcoidia numbers of 103 16S rRNA gene

copies ml-1 culture, compared to site 1317, which had Dehalococcoidia numbers of 105

16S rRNA gene copies ml-1 (Figure 16). In site 1318, only those sediment cultures

incubated with 2,4,6-tribromoresorcinol, 2,3-dichlorophenol, iron(III), and

tetrachloroethene had Dehalococcoidia numbers above the detection threshold of 103

16S rRNA gene copies ml-1 (Figure 16). Sediment cultures inoculated with site 1317

sediments showed highest Dehalococcoidia numbers of 9.2 x 104 16S rRNA gene

copies ml-1 within a culture that contained no supplied electron acceptor, which was

prepared as a control. In addition, for site 1317, increased Dehalococcoidia numbers of

4.8 x 104 and 8.2 x 104 16S rRNA gene copies ml-1 were observed in cultures amended

with sulphate and 2,4,6–tribromoresorcinol, respectively.

Figure 16. Dehalococcoidia quantification of sediment cultures set up with sediments from site 1318 at a

depth of 23.15 mbsf, and site 1317 at a depth of 227 mbsf. Both sites are located in the Porcupine

Seabight at the continental margin of Ireland (see Table 1 for details). Standard deviations of triplicate

measurements from the qPCR assay are shown. Abbreviations: NEA - no electron acceptor as a control,

NEA-I and NEA-II are two parallel sediment cultures without electron acceptor; Mn – manganese(IV); Fe

– iron(III); SO4 - sulphate; TCB - 1,2,3- and 1,2,4-trichlorobenzene mixed in a 1:1 proportion; DCP - 2,3-

dichlorophenol; CP - 2-chlorophenol; PCE - tetrachloroethene; BMBA - 4-bromo-3,5-dimethoxybenzoic

acid; TBR - 2,4,6-tribromoresorcinol.

Results

In line with the sediment cultures inoculated with sediments from Århus Bay, sediment

cultures from Ireland indicated that sulphate and 2,4,6–tribromoresorcinol may promote

Dehalococcoidia growth. However, the high numbers of Dehalococcoidia in a culture

supplied without any electron acceptor reveals that the inoculated sediment may be a

source of nutrients and/or electron acceptors which may support Dehalococcoidia

growth in sediment cultures.

Sediment cultures inoculated with sediment from the coast of central off Chile, site

7165 showed similar Dehalococcoidia numbers of 4 x 103 16S rRNA gene copies ml-1

culture (mean of duplicates) for all electron acceptors (upper panel, Figure 17).

Therefore, none of the electron acceptors supplied specifically enriched

Dehalococcoidia.

For the case of sediment cultures set up with sediment from site 7155, similar

Dehalococcoidia numbers of 103 16S rRNA gene copies ml-1 culture (mean of

duplicates) were observed after 4 months of incubation for all electron acceptors tested

(lower panel, Figure 17). However, after an incubation time of 10 months, highest

Dehalococcoidia abundance (and the only ones above the threshold) was observed in

sediment cultures supplied with sulphate and 2–chlorophenol as potential electron

acceptors and in those amended with no electron acceptor. Highest Dehalococcoidia

increments of an order of magnitude compared to the previous time point of 4 months

were observed in those sediment cultures supplied with 2–chlorophenol and without any

potential electron acceptor (lower panel, Figure 17). For a third time point

corresponding to a year of incubation, sediment cultures supplied with iron(III) and 2,3–

dichlorophenol had highest Dehalococcoidia numbers and increases in respect to the

previous measured time point. Altogether, sediment cultures from site 7155 did not

indicate a sustained growth of Dehalococcoidia under any of the potential electron

acceptors tested here.

Results

Figure 17. Dehalococcoidia quantification of sediment cultures set up with sediments from Chile, site

7165 at a depth of 635– 640 cmbsf (upper panel), and site 7155 at a depth of 437– 442 cmbsf (lower

panel). Shown are means of duplicate cultures ± SD. Abbreviations: NEA – no electron acceptor, as a

control; Mn – manganese(IV); Fe – iron(III); SO4 – sulphate; HA – humic acids; TCB – 1,2,3- and 1,2,4-

trichlorobenzene mixed in a 1:1 proportion; DCP – 2,3-dichlorophenol; CP – 2-chlorophenol; PCE –

tetrachloroethene; BMBA – 4-bromo-3,5-dimethoxybenzoic acid; TBR – 2,4,6-tribromoresorcinol.

Results

Due to the variability of the results in Dehalococcoidia numbers from the different time

points of sediments from Chile, an assay to prove if the methods that were used (i.e.,

sampling of sediment culture and subsequent DNA isolation and qPCR) were reliable

for quantification of bacteria was conducted. For this, three samples of 1, 2, and 4 ml

were taken from a sediment culture of Chile site 7165, followed by DNA isolation and

the performance of triplicate qPCR measurements with primers targeting for total

Bacteria (primer pair 341F and 534R, Table 7). Results showed a cycle of difference

between the 1 and 2 ml sampled sediment cultures in the qPCR assay and 2 cycles of

difference between the 2 and 4 ml sampled sediment cultures. Thus, this approach of

sample withdrawal from the sediment culture, DNA isolation, and qPCR assay was

reliable for quantifying 16S rRNA gene copy numbers.

In general, these experimental results show that Dehalococcoidia were growing in the

sediment cultures, demonstrating that they withstood the changes in temperature and

pressure during the sampling campaign and that the supplied media based on the

medium used for Dehalococcoides mccartyi strain CBDB1 was appropriate for the

cultivation of marine Dehalococcoidia.

3.1.2 Total Bacteria abundance under various terminal electron acceptors in

sediment cultures

The total bacterial community abundance within Chile site 7155 sediment cultures was

measured with primers targeting the 16S rRNA gene of all bacteria (primer pair 341F

and 534R, Table 7). The results showed similar total bacterial abundance for all electron

acceptors of 107 16S rRNA gene copies ml-1 culture after four months of incubation

(Figure 18). Therefore, Dehalococcoidia represent around 0.01% of the total bacterial

community in the Chile 7155 sediment cultures. A second time point, after ten months

of incubation, showed a decrease in total bacterial numbers for all electron acceptors

except for sulphate.

Results

Figure 18. Total bacterial 16S rRNA gene copy numbers in sediment cultures set up with sediments from

the continental margin of Chile, site 7155 (see Table 1 for details). Shown are means of duplicate cultures

± SD. Abbreviations: NEA – no electron acceptor, as a control; Mn – manganese(IV); Fe – iron(III); SO4

– sulphate; HA – humic acids; TCB – 1,2,3- and 1,2,4-trichlorobenzene mixed in a 1:1 proportion; DCP –

2,3-dichlorophenol; CP – 2-chlorophenol; PCE – tetrachloroethene; BMBA – 4-bromo-3,5-

dimethoxybenzoic acid; TBR – 2,4,6-tribromoresorcinol.

3.1.3 Colony formation in deep-agarose dilution tubes under various terminal

electron acceptors

None of the deep-agarose dilution tubes prepared with the various electron acceptors for

both Århus and Ireland sediments showed any colony formation, except two tubes,

which may also have been due to contamination. One of these deep-agarose tubes was a

third sequential dilution from a sediment culture inoculated with sediments from site

1317 from Ireland, amended with humic acids, which had white colonies. The second

deep-agarose tube was inoculated with an Århus sediment culture inoculum, amended

with 4-bromo-3,5-dimethoxybenzoic acid, and showed growth of small white colonies

all over the tube, but particularly under the surface, and although the redox dye

resazurin within the agarose tube did not change in colour (i.e., to pink; when at positive

redox potentials), maybe traces of oxygen may have induced these colonies growth.

In contrast, deep-agarose dilution tubes prepared from cultures inoculated with

sediments from Chile, site 7155, showed many colonies after an incubation time of two

weeks to one month. Colonies in the first dilution were small and very numerous,

however with increasing dilution, colonies began to be less numerous and bigger.

Results

Deep-agarose dilution tubes prepared with 2,3-dichlorophenol, humic acids, iron(III),

tetrachloroethene, and with no potential electron acceptor had the highest colony density

from all. Colony morphologies varied from small black and/or white in tubes amended

with 2,3-dichlorophenol or with no electron acceptor, to large ones, in the case of humic

acids and iron(III) (Figure 19). These large colonies were brown for humic acids and

dark red for iron, which were especially large. In agarose tubes with PCE, colonies were

black and with a black surrounding halo (Figure 19). This black colour from the

colonies may be iron sulphide, which is a black precipitate. Iron sulphide results from

the conversion of sulphate to sulphide by sulphate-reducing bacteria and the further

combination of sulphide with iron(II), which is present in the medium. Sulphate may

come from the inoculum (i.e., the sediment), as no sulphate is provided in the medium.

PCE conversion to lower chlorinated compounds was not observed. Thus, the black

colour from these colonies may indicate that the colonies may be formed by sulphate

reducers. Deep-agarose dilution tubes amended with humic acids developed gas bubbles

throughout the tube which may be due to methanogens forming methane and/or

fermenters forming carbon dioxide. Growth in agarose tubes amended without any

electron acceptor indicated that bacteria may be using the citrate from titanium (III)

citrate or residual compounds from the sediment inoculum, which even though diluted,

may still be present. Deep-agarose dilution tubes containing sulphate were observed to

develop a black precipitate in parts of the tube after longer periods of incubation

indicating the presence of sulphate reducers.

Figure 19. Colony colours, shapes and visual appearance varied from white to black, brown or red in the

semisolid deep-agarose dilution tubes. Colonies from deep-agarose dilution tubes amended with no

electron acceptor (A, B), with iron(III) (C), humic acids (D), PCE (E) and with 2,3-dichlorophenol (F, G)

are shown.

Some colonies were selected for sub-cultivation (Figure 20). For that, colonies were

picked with sterile Pasteur pipettes or with sterile syringes associated to thick sterile

needles. Picked colonies were immediately transferred into liquid. Deep-agarose

dilution tubes were prepared again from these sub-cultivated in-liquid-medium colonies

after two weeks of incubation. These newly prepared deep-agarose dilution tubes

showed in general consistent morphology with the original picked colony (A, B and C

in Figure 20). However, in some other cases, the newly prepared agarose tubes showed

Results

differing colonies to the original colony (see E in Figure 20), indicating that more than

one colony was picked or that contamination occurred in the newly prepared agarose

tubes. The newly prepared agarose tubes had high colony concentration with smaller

colony sizes, due to competition for space and nutrients. Those deep-agarose dilution

tubes amended with humic acids showed gas bubbles, as observed previously.

Figure 20. Colonies from sub-cultivated deep-agarose dilution tubes compared to the original colony.

Colonies formed in deep-agarose dilution tubes amended with humic acids (A), with PCE (B and D), and

with iron(III) (C).

3.1.4 Colony identification after 16S rRNA gene sequencing

Thirty of the observed colonies were picked, DNA was isolated from them, the 16S

rRNA gene was amplified and amplicons were sequenced. All sequenced colonies

belonged to the phylum Firmicutes, class Clostridia, although three colonies belonged

to the class Bacillales (Table 8). Many of them showed high similarity to the genera

Pelotomaculum (8/30 sequences), Gracilibacter (4/30) and Pelosinus (4/30),

Vulcanibacillus (3/30), Clostridium (3/30), Morella (2/30) and Desulfotomaculum

(2/30).

Results

Table 8. Colony similarities to closest cultured bacterial member based on the 16S rRNA gene partial

sequence. Colonies 1-16 belong to colonies which were picked exclusively to know their phylogenetic

affiliation. Colonies 17-32 come from sub-cultivated colonies, from which 1 ml of liquid culture was

sampled for DNA to find out their phylogeny. All of them were cultivated with titanium (III)-citrate as

reducing agent. The assessment of the 16S rRNA gene sequence similarity was done using Blast (Basic

Local Alignment Search Tool; (Altschul et al 1990)) and the NCBI database. Abbreviations: TCB – 1,2,3-

and 1,2,4-trichlorobenzene mixed in a 1:1 proportion; BMBA – 4-bromo-3,5-dimethoxybenzoic acid.

No.

Electron

acceptor

Closest cultivated bacterial sequence

Literature

2,4,6-

tribromo-

resorcinol

Gracilibacter thermotolerans strain JW/YJL-S1

Firmicutes; Clostridia; Clostridiales; Graciibacteraceae;

Gracilibacter.

95% coverage; 88% identity

(Lee et al

2006)

Gracilibacter thermotolerans strain JW/YJL-S1

Firmicutes; Clostridia; Clostridiales; Graciibacteraceae;

Gracilibacter.

93% coverage; 96% identity

(Lee et al

2006)

TCB

Candidatus Heliobacterium aridinosum

Firmicutes; Clostridia; Clostridiales; Heliobacteriaceae;

Heliobacterium

95% coverage; 88% identity

Girija et al,

unpublished

BMBA

Thermincola carboxydiphila strain 2204

Firmicutes; Clostridia; Clostridiales; Peptococcaceae;

Thermincola.

100% coverage; 89% identity

(Sokolova et

al 2005)

humic acids

Sporotalea propionica strain TmPM3

Firmicutes; Clostridia; Clostridiales; Veillonellaceae; Sporotalea

100% coverage; 85% identity

(Boga et al

2007)

humic acids

Pelotomaculum isophthalicicum

Firmicutes; Clostridia; Clostridiales; Peptococcaceae;

Pelotomaculum.

99% coverage; 85% identity

(Qiu et al

2006)

humic acids

Desulfotomaculum sp. 16S rRNA gene, DSM 7440

Firmicutes; Clostridia; Clostridiales;

Peptococcaceae;Desulfotomaculum.

92% coverage; 85% identity

(Stackebrandt

et al 1997)

Fe(III)

Pelotomaculum isophthalicicum

Firmicutes; Clostridia; Clostridiales; Peptococcaceae;

Pelotomaculum

99% coverage; 87% identity

(Qiu et al

2006)

PCE

Pelotomaculum isophthalicicum

Firmicutes; Clostridia; Clostridiales;

Peptococcaceae;Pelotomaculum

90% coverage; 90% identity

(Qiu et al

2006)

Methanol

Pelotomaculum isophthalicicum

Firmicutes; Clostridia; Clostridiales; Peptococcaceae;

Pelotomaculum.

100% coverage; 86% identity

(Qiu et al

2006)

2-Chloro-

phenol

Clostridium sp. JC3

Firmicutes; Clostridia; Clostridiales; Clostridiaceae;Clostridium.

94% coverage; 83% identity

Syutsubo, et

al,

unpublished

Fe(III)

Lutispora thermophila

Firmicutes; Clostridia; Clostridiales; Clostridiaceae; Lutispora.

99% coverage; 87% identity

(Shiratori et

al 2006)

Fe(III)

Pelotomaculum isophthalicicum

Firmicutes; Clostridia; Clostridiales;

Peptococcaceae;Pelotomaculum.

100% coverage; 87% identity

(Qiu et al

2006)

Results

No.

Electron

acceptor

Closest cultivated bacterial sequence

Literature

Fe(III)

Pelotomaculum isophthalicicum

Firmicutes; Clostridia; Clostridiales;

Peptococcaceae;Pelotomaculum.

100% coverage; 86% identity

(Qiu et al

2006)

Fe(III)

Pelosinus sp. UFO1

Firmicutes; Clostridia; Clostridiales; Veillonellaceae;Pelosinus.

100% coverage; 99% identity

Ray et al,

unpublished

PCE

Pelosinus sp. UFO1

Firmicutes; Clostridia; Clostridiales; Veillonellaceae;Pelosinus.

100% coverage; 99% identity

Ray et al,

unpublished

BMBA

Moorella glycerini strain JW/AS-Y6

Firmicutes; Clostridia;

Thermoanaerobacterales;Thermoanaerobacteraceae; Moorella

group; Moorella.

94% coverage; 84% identity

(Slobodkin et

al 1997)

BMBA

Pelotomaculum isophthalicicum

Firmicutes; Clostridia; Clostridiales;

Peptococcaceae;Pelotomaculum.

99% coverage; 84% identity

(Qiu et al

2006)

BMBA

Pelotomaculum isophthalicicum

Firmicutes; Clostridia; Clostridiales;

Peptococcaceae;Pelotomaculum.

100% coverage; 84% identity

(Qiu et al

2006)

TCB

Vulcanibacillus modesticaldus

Firmicutes; Bacillales; Bacillaceae; Vulcanibacillus.

99% coverage; 95% identity

Swiderski,

unpublished

TCB

Vulcanibacillus modesticaldus

Firmicutes; Bacillales; Bacillaceae; Vulcanibacillus.

100% coverage; 93% identity

Swiderski,

unpublished

TCB

Vulcanibacillus modesticaldus

Firmicutes; Bacillales; Bacillaceae; Vulcanibacillus.

99% coverage; 93% identity

Swiderski,

unpublished

2,3-

dichloro-

phenol

Clostridium sp. AAN11

Firmicutes; Clostridia; Clostridiales; Clostridiaceae; Clostridium.

91% coverage; 84% identity

Ueno &

Yamazawa,

unpublished

2,3-

dichloro-

phenol

Clostridium acetireducens strain DSM 10703

Firmicutes; Clostridia; Clostridiales; Clostridiaceae; Clostridium.

95% coverage; 91% identity

(Orlygsson et

al 1996)

none

Desulfotomaculum carboxydivorans strain CO-1-SRB

Firmicutes; Clostridia; Clostridiales;

Peptococcaceae;Desulfotomaculum.

100% coverage; 84% identity

(Parshina et

al 2005)

none

Moorella glycerini strain JW/AS-Y6

Firmicutes; Clostridia;Thermoanaerobacterales;

Thermoanaerobacteraceae; Moorella group; Moorella.

95% coverage; 82% identity

(Slobodkin et

al 1997)

Fe(III)

Pelosinus sp. UFO1

Firmicutes; Clostridia; Clostridiales; Veillonellaceae; Pelosinus.

100% coverage; 99% identity

Ray et al,

unpublished

Fe(III)

Pelosinus sp. UFO1

Firmicutes; Clostridia; Clostridiales; Veillonellaceae; Pelosinus.

100% coverage; 99% identity

Ray et al,

unpublished

Results

3.2 ORGANOHALIDE TRANSFORMATION IN MARINE SEDIMENT

CULTURES

Sediment cultures were amended with various halogenated compounds (1,2,3-TCB and

1,2,4-TCB, 2,3-dichlorophenol, 2-monochlorophenol, 2,4,6-tribromoresorcinol, 4-

bromo-3,5-dimethoxybenzoic acid, hexachlorobenzene, hexabromobenzene, or PCE) as

potential electron acceptors for all tested sediments (Århus, Ireland, Chile). From the

halogenated compounds, the concentrations of 1,2,3-TCB, 1,2,4-TCB, 2,3-

dichlorophenol, 2-monochlorophenol, PCE, and hexachlorobenzene transformation

products in the cultures were monitored with GC-FID. Exclusively 1,2,3-TCB was

transformed to 1,3-DCB in two sediment cultures, one inoculated with sediments from

Århus (after 346 days of incubation), and the other with sediments from site 7155 of

Chile as detailed in section 3.2.1. No further transformation of 1,3-DCB to

monochlorobenzene was observed.

3.2.1 Transformation of 1,2,3-TCB to 1,3-DCB in marine sediment cultures

Complete transformation of 1,2,3-TCB (80 μM, which was solved in 1,2,4-TCB in a 1:1

proportion) to 1,3-DCB was observed in one sediment culture inoculated with sediments

from Chile, site 7155 (at a depth of 437–442 cmbsf, Table 1), after six months of

incubation. This sediment culture did however not transform 1,2,4-TCB.

After complete 1,2,3-TCB transformation, the sediment culture was transferred in three

parallels with a 13% (v/v) inoculum to fresh medium amended with 80 μM of 1,2,3–

TCB (solved in 1,2,4-TCB in a 1:1 proportion). One of the three daughter cultures

showed complete conversion of 1,2,3-TCB to 1,3-DCB after two months of incubation.

In the other two replicate daughter cultures, no transformation of 1,2,3-TCB to DCBs

was observed.

Further transferring consisted of six subcultures (using inocula of 4% (v/v) in fresh

medium, and amended with 80 μM 1,2,3-TCB solved in acetone for three subcultures,

and 40 μM for the other three), and aimed for an enrichment in a larger number of

subcultures and thus higher total biomass, and for an enrichment of specifically 1,2,3-

TCB transforming microorganisms when increasing the 1,2,3-TCB concentrations.

However, none of the six subcultures transformed 1,2,3-TCB.

After this, and to reproduce the organohalide transformation, a new batch of sediment

cultures was set up. Sediment cultures were prepared in four replicates with 9 or 6%

inoculum (v/v) of sediment from Chile site 7155 and amended with 45 µM of 1,2,3-

TCB, solved in acetone. Sediment cultures were prepared in four parallels and reduced

Results

either with titanium (III) citrate, iron sulphide, or sodium sulphide together with L-

cysteine (Table 2). All sediment cultures reduced with iron sulphide showed a change of

colour of the redox dye resazurin after the inoculation of the sediment, indicating a

positive redox potential in the cultures; and these cultures were discarded. Cultures

reduced with titanium (III) citrate or sodium sulphide together with L-cysteine remained

anoxic (at negative redox potentials as indicated by the transparent colour of the redox

indicator resazurin), and they were monitored and further maintained.

The formation of 1,3-DCB was observed in two of the four parallel sediment cultures

(labelled as G0, see section 3.2.2, Figure 23) reduced with titanium (III) citrate after 58

days (Figure 21). On day 65, all the cultures were amended with 40 μM of 1,2,3-TCB,

before G0–a had depleted all 1,2,3-TCB. The parallels G0–b and G0–d transformed

1,2,3-TCB to 1,3-DCB more slowly. The formation of 1,3-DCB was observed after an

incubation time of 270 and 325 days for G0–b and G0–d, respectively (results not

shown). All cultures were amended with more 1,2,3-TCB at day 65, and subsequently

whenever 1,2,3-TCB was depleted. In all cultures and also in negative controls without

bacteria, a slow decrease of 1,2,3-TCB was observed and was attributed to the sampling

procedure and the piercing of the Teflon liner of the rubber stoppers.

Results

Figure 21. Time-course of 1,2,3-TCB transformation (upper panel) and 1,3-DCB formation (lower panel)

in four parallel G0 cultures reduced with titanium (III) citrate. G0 cultures were inoculated with 9% (v/v)

sediment from Chile site 7155 (from a depth of 437–442 cmbsf). No production of dichlorobenzene was

observed in medium blanks or in autoclaved sediment cultures reduced with titanium (III) citrate.

Results

Three of the four parallel sediment cultures reduced with sodium sulphide plus L-

cysteine transformed 1,2,3-TCB to 1,3-DCB (Figure 22). The sediment culture G0–e

was not observed to form 1,3-DCB within 277 days. 1,2,3-TCB (40 μM) was supplied

to the cultures after 65 and 221 days. The formation of 1,3-DCB was observed within

42 days for two sediment cultures (G0–g and G0–h, lower panel of Figure 22).

Figure 22. Time-course of 1,2,3-TCB transformation (upper panel) and 1,3-DCB formation (lower panel)

in four parallel G0 cultures reduced with sodium sulphide and L-cysteine. G0–e, –f, –g cultures were

inoculated with 6% (v/v), and G0–h culture with 9% (v/v) of sediment from Chile site 7155 (at a depth of

437–442 cmbsf). No production of dichlorobenzene was observed in medium blanks or in autoclaved

sediment cultures reduced with sodium sulphide and L-cysteine.

Results

Additionally, triplicate sediment cultures reduced with either titanium (III) citrate or

sodium sulphide with L-cysteine were supplied with vancomycin or with ampicillin plus

the methanogen-inhibitor 2-bromoethanesulfonate (BES). None of the cultures supplied

with antibiotics and/or BES did transform 1,2,3-TCB to dichlorobenzenes. Thus, the

specific enrichment of dehalogenating bacteria by the addition of vancomycin or

ampicillin/BES was not possible, suggesting that Gram-positives may have a direct or

indirect role in 1,2,3-TCB transformation.

3.2.2 Enrichment of trichlorobenzene dechlorinating bacteria from sediment

cultures

The enrichment procedure consisted of transferring 9% of inoculum into fresh media

once all the 1,2,3-TCB was completely transformed or about to be transformed in the

culture. The hypothesis of this experiment was that the microorganisms performing

1,2,3-TCB transformation obtain a growth advantage over other microorganisms by

repeated subcultivation in the presence of organohalides. Four subsequent transfers

(subcultures G1–G4; Figure 23) maintained organohalide transformation in cultures

reduced with titanium (III) citrate (Table 9). In cultures reduced with sodium sulphide

and L-cysteine, two subsequent transfers (subcultures G1–G2) maintained organohalide

transformation (Table 9).

Figure 23. Enrichment scheme of dehalogenating cultures. The nomenclature G0–G4 will be consistently

used in this thesis to describe the different cultures.

Results

Table 9. Summary of cultures set up within this study reduced either with titanium (III) citrate or sodium

sulphide together with L-cysteine. The total number of cultures per generation (G0–G4; “Total”) and the

number of cultures that transformed 1,2,3-TCB to 1,3-DCB (“Active”) are shown.

Reduced with: G0 G1 G2 G3 G4

Total1 Active2 Total1 Active2 Total1 Active2 Total1 Active2 Total1 Active2

Titanium (III)

citrate

4 4 34 29 46 38 16 7 8 2

Sodium

sulphide and

L-cysteine

4 3 20 10 15 6 3a 0 6a 0

1Total number of cultures; 2Number of active cultures, where formation of 1,3-DCB was observed associated to 1,2,3-TCB

transformation. aInoculum was a G2 o G3 culture which was originally reduced with titanium (III) citrate.

A first set of G1 subcultures containing 2% (v/v) inoculum from G0–a, –c, –g, and –h

was prepared in triplicate. This first set of subcultures was prepared when G0 cultures

had been incubated for 73 days and the formation of 1,3-DCB had been observed

(Figure 21 and Figure 22). The three triplicate G1–c subcultures transformed 1,2,3-TCB

to 1,3-DCB. Two from the three G1–c subcultures had to be discarded due to the change

in colour to pink of the resazurin, which indicated that the subcultures were no longer

reduced. The third G1–c subculture was maintained and subsequently transferred. G2

(three subcultures), G3 (16 subcultures) and G4 (six subcultures) were obtained and

dehalogenated 1,2,3-TCB to 1,3-DCB. All G3 and G4 subcultures of this study were

subcultures of this G1–c subculture. G1–a, and –g did not transform 1,2,3-TCB to 1,3-

DCB. One of the triplicates from G1–h showed transformation of 1,2,3-TCB to 1,3-

DCB and was transferred into fresh medium, however, no active subculture was

obtained from it.

A second set of G1 subcultures were produced with a higher proportion of G0 inoculum

of 9% (v/v), when the G0 cultures were 245 days old for G1–a and G1–c, and 277 days

for G0–f, –g, and –h. The subcultures G1–a and –c, as well as G1–f, –g and –h were

prepared in parallel, respectively. G1–b and –d subcultures were prepared later on, after

dechlorination in the G0–b and –d cultures was observed. All G1 subcultures

transformed 1,2,3-TCB to 1,3-DCB either if reduced with titanium (III) citrate (Figure

24) or with sodium sulphide together with L-cysteine (Figure 25). All G1 subcultures

transformed 1,2,3-TCB to 1,3-DCB in a shorter time spam that the G0 culture from

which they were inoculated.

Results

Figure 24. Time-course of 1,2,3-TCB transformation (upper panel) and 1,3-DCB formation (lower panel)

in twelve G1 subcultures reduced with titanium (III) citrate. The mean of 1,2,3-TCB and 1,3-DCB

concentration in triplicate subcultures are plotted ± SD. The “–a, –b, –c, –d” in the labels refer to the

specific G0 culture from which each G1 subculture was inoculated from. For example, G1–a are triplicate

subcultures inoculated with 9% (v/v) of culture G0–a. Subcultures G1–a and –c are parallels, inoculated

and measured at same time. The subcultures G1–b and –c were set up later at different times.

Results

Figure 25. Time-course of 1,2,3-TCB transformation (upper panel) and 1,3-DCB formation (lower panel)

in six parallel G1 subcultures reduced with sodium sulphide and L-cysteine. The mean of 1,2,3-TCB and

1,3-DCB concentration in duplicate subcultures are plotted ± SD. The “–f, –g, –h” in the labels refer to

the specific G0 culture from which each G1 subculture was inoculated from. For example, G1–f refers to

duplicate subcultures inoculated with 9% (v/v) of culture G0–f. No G1–e subcultures were set up as G0–e

did not transform 1,2,3-TCB to 1,3-DCB. All subcultures were set up and measured at same time point-s

until 1,2,3-TCB was depleted and subsequently transferred to fresh medium.

Results

The subcultures G1–a, –c, –d, and –g were selected for further subculturing. 1,2,3-TCB

transformation to 1,3-DCB was observed in the great majority of G2 subcultures (Table

9). Activity (defined here as formation of 1,3-DCB from 1,2,3-TCB) was observed in a

greater proportion of G2 subcultures reduced with titanium (III) citrate compared to

those reduced with sodium sulphide and L-cysteine (Table 9).

G2 subcultures transformed 1,2,3-TCB in a longer time spam than G1 and sometimes

also than G0 cultures, either when reduced with titanium (III) citrate or sodium sulphide

together with L-cysteine (Figure 26).

Figure 26. Time-course of 1,2,3-TCB transformation (upper panel) and 1,3-DCB formation (lower panel)

in 24 G2 subcultures. G2–a and –c are each the mean of concentration values coming from nine parallel

subcultures reduced with titanium (III) citrate ± SD. G2–g are the mean of concentration values of six

parallel subcultures reduced with sodium sulphide and L-cysteine ± SD.

Results

G3 subcultures were reduced with titanium (III) citrate or sodium sulphide plus L-

cysteine. The G3 subcultures reduced with sodium sulphide plus L-cysteine were

inoculated with 9% (v/v) of a G2 subculture which was previously reduced with

titanium (III) citrate to investigate if a change of reducing agent may affect

dechlorination. No active G3 subculture reduced with sodium sulphide and L-cysteine

was observed in any of the triplicates prepared. Sixteen G3 subcultures were prepared

and reduced with titanium (III) citrate and seven of them were active. Out of those

seven subcultures, five and two, respectively, were parallel subcultures. The five G3

parallel subcultures formed 1,3-DCB after 21 (three subcultures) and 42 (two

subcultures) days of incubation. The two G3 parallel subcultures formed 1,3-DCB

within 85 and 134 days of incubation. One of each parallels (a total of two G3

subcultures) were further transferred to fresh medium. A total of eight G4 subcultures

were prepared and two of them were active after 149 and 274 days of incubation. A

further transfer, G5 triplicate subcultures reduced with titanium (III) citrate, was

additionally prepared. During the monitoring time of 167 days, no formation of 1,3-

DCB was observed. Due to time limitations, the G5 subcultures could not be further

monitored.

G2 and G3 subcultures (inoculated with 12% v/v from G1–b and –f, and G2–c and –g)

either reduced with titanium (III) citrate or with sodium sulphide and L-cysteine were

tested with hexachlorobenzene (HCB) instead of 1,2,3-TCB to investigate if the

subcultures may further transform higher chlorinated compounds. However after 337

days of incubation, no lower halogenated products (1,2,3-TCB or 1,3-DCB) were

observed in any of the six subcultures tested.

In addition, it was investigated if hydrogen (1 bar overpressure) and acetate (5 mM)

affected organohalide transformation. For that, 32 subcultures (either G2 or G3) were

prepared amended with 60 µM of 1,2,3-TCB solved in acetone. From those, eight were

amended with hydrogen and acetate, eight with acetate alone, eight with hydrogen

alone, and eight without hydrogen and acetate. After 253 days of incubation, 1,3-DCB

was observed in five of the subcultures. One of the eight parallel subcultures amended

only with hydrogen, one of the subcultures with neither hydrogen nor acetate, and three

of the subcultures amended only with acetate showed 1,3-DCB formation (Table 10).

Results

Table 10. Transformation of 1,2,3-TCB to 1,3-DCB by G2 and G3 subcultures after 253 days of

incubation in the presence or absence of hydrogen and/or acetate. “+” refers to culture amended with

either acetate (Ac) or hydrogen (H2), and “–“ to non-amended.

Culture

generation

Number of cultures per

treatment

Number of cultures which transformed 1,2,3-TCB

Ac+/H

–

Ac–/H

3.2.3 Quantification of total Bacteria and known organohalide-respiring bacteria

by qPCR

Three parallel G0 cultures were selected for the quantification of total Bacteria and

other known organohalide-respiring bacteria, i.e., Dehalococcoidia, Dehalobacter, and

Desulfitobacterium. These G0 cultures were G0–a, –b, and –c, which were reduced with

titanium (III) citrate. Bacterial quantification was performed at various time points of

incubation (i.e., 3, 36, 74, and 238 days) while dehalogenation of 1,2,3-TCB occurred,

except for G0–b (Figure 21). The formation of 1,3-DCB was observed after 270 days of

incubation for the culture G0–b.

After three days of incubation, the average number (± SD) of total bacterial 16S rRNA

gene copies was 1.2 x 107 ± 2.9 x 106 ml-1 for the triplicate G0 cultures. Bacterial 16S

rRNA copies decreased with incubation time to 2.6 x 106 ± 2.2 x 106, 6.8 x 106 ±

2.9 x 106, and 1.9 x 106 ± 4.6 105 ml-1, for the time points of 36, 74, and 238 days,

respectively (Figure 27).

Figure 27. Total bacterial 16S rRNA gene copy numbers in G0 cultures reduced with titanium (III)

citrate. Three parallel G0 cultures are shown. G0–a and G0–c completely dehalogenated 1,2,3-TCB

within 238 days. Shown are means of qPCR triplicate measurements ± SD.

Results

In the G0 cultures, Dehalococcoidia 16S rRNA genes were amplified with both the

primer pairs DEH-Fa and DEH-R (matching to most Dehalococcoidia), and Dehal-F5

and Dehal-R (matching to Chile-specific Dehalococcoidia) (Table 7), although at low

copy numbers for both primer pairs. The average number of Dehalococcoidia 16S

rRNA gene copies of 2.8 x 103 ± 1.3 x 103 ml-1 (amplified with DEH-Fa and DEH-R)

was observed in triplicate G0 cultures after three incubation days. Dehalococcoidia 16S

rRNA gene copies decreased with time to average values of 5.3 x 102 ± 9.1 x 10, 6.5 x

102 ± 1.8 x 102, and 3.5 x 102 ± 1.6 x 102, for the time points of 36, 74, and 238 days of

incubation, respectively (Figure 28).

Figure 28. Dehalococcoidia 16S rRNA gene copy numbers in G0 cultures reduced with titanium (III)

citrate amplified the primer pair DEH-Fa and DEH-R targeting most Dehalococcoidia. Three parallel G0

cultures are shown. G0–a and G0–c completely dehalogenated 1,2,3-TCB within 238 days. Shown are

means of qPCR triplicate measurements ± SD.

The proportion of Dehalococcoidia 16S rRNA gene copies relative to total bacterial 16S

rRNA gene copies in G0 parallel cultures was ~0.02%, which did not increase with time

meanwhile dehalogenation was taking place (Table 11).

Table 11. Proportion of Dehalococcoidia 16S rRNA gene copies relative to total bacterial 16S rRNA

gene copies in triplicate G0 cultures at different time points is shown.

Time of incubation (days)

Culture 3 36 74 238

G0-a 0.02% 0.05% 0.01% 0.02%

G0-b 0.02% 0.01% 0.01% 0.01%

G0-c 0.03% 0.02% 0.01% 0.02%

Results

G1 and G2 subcultures were also monitored for Dehalococcoidia copy numbers. In the

qPCR assay, amplification was obtained for G1 and G2 with both the primer pairs

(DEH-Fa and DEH-R; Dehal-F5 and Dehal-R). However, the melt-curves from the

qPCR assay revealed no clear peaks indicative of specific amplification, and further

visualization of these amplicons by agarose gel electrophoresis revealed no clear bands.

Together, these results indicated that the primers amplified non-specifically for G1 and

G2.

In addition, quantification of 16S rRNA gene copies from Dehalobacter and

Desulfitobacterium was performed using the primers Dre441F and Dre645R_Ch, and

DSB406F and DSB619R (Table 7), respectively, in the triplicate G0 cultures. For the

quantification of 16S rRNA gene copy numbers, no standard for both Dehalobacter and

Desulfitobacterium was run in the qPCR assay due to lack of cloned 16S rRNA genes of

Dehalobacter and Desulfitobacterium. In Table 12, the values for the threshold cycle

(Ct) for the triplicate G0 cultures is shown as a proxy to indicate Dehalobacter and

Desulfitobacterium increase or decrease with time. As both Dehalobacter and

Desulfitobacterium qPCR assays were performed in the same qPCR run, Ct values can

be compared among the samples. The qPCR assay revealed that Dehalobacter 16S

rRNA gene copies decreased with time (Ct values increased with time, Table 12) in G0–

a and –c cultures, which dehalogenated 1,2,3-TCB during the time span of 3–238 days

of incubation (Figure 21). The G0–b culture showed an increase in Dehalobacter 16S

rRNA gene copies after 36 and 74 days compared to the Ct value of 3 days (Table 12).

However, G0–b did not dehalogenated 1,2,3-TCB during the first 74 days (Figure 21).

Thus, Dehalobacter increase cannot be associated to dehalogenation in G0–b.

Desulfitobacterium 16S rRNA gene copies in G–a increased with time with respect of

the first time point of 3 days. However, for G–b and –c, Desulfitobacterium copies

remained stable with time (Table 12).

Results

Table 12. qPCR threshold cycles (Ct) for each sample amplified with primers targeting the 16S rRNA

gene of Dehalobacter and Desulfitobacterium. Shown are mean of duplicate values for Ct ±SD run in the

same qPCR assay. Triplicate G0 cultures at different time points of incubation were analysed together

with a sediment sample from Chile, site 7155 (at a depth of 437–442 cmbsf), and a negative control (1 μl

water as template). Samples with Ct values ≤29 indicate abundant 16S rRNA gene copies, meanwhile Ct

values of 38–40 indicate few or no16S rRNA gene copies

Sample

Dehalobacter Desulfitobacterium

Ct mean ± SD Ct mean ± SD

Sediment1 24.7 ± 0.1 27.5 ± 0.1

Negative Control 37.0 ± 0.1 35.5 ± 0.1

G0-a 3 days 30.0 ± 0.2 26.2 ± 0

36 days 31.7 ± 0 23.0 ± 0.1

74 days 32.8 ± 0 24.0 ± 0.1

238 days 35.0 ± 0.2 25.3 ± 0

G0-b 3 days 32.4 ± 0.3 18.5 ± 0

36 days 28.1 ± 0.2 19.9 ± 0

74 days 29.4 ± 0.2 20.8 ± 0.1

238 days 31.4 ± 0.7 21.1 ± 0.3

G0-c 3 days 30.7 ± 0.1 19.4 ± 0.2

36 days 31.7 ± 0.2 21.2 ± 0.1

74 days 32.8 ± 0.3 21.7 ± 0.1

238 days 33.0 ± 0.2 21.7 ± 0

1 1 μl DNA template was used from DNA isolated from Chile site 7155.

No amplification of known reductive dehalogenase genes was possible with the various

primers either for Dehalococcoides mccartyi reductive dehalogenase genes rdhA

(primers RRF2 and B1R; RDH F1C and RDH R1C, Table 7) or for Dehalobacter-

Desulfitobacterium rdhA genes (primers dehaloF3 and dehaloR2; dehaloF5 and

dehaloR4, Table 7).

3.2.4 Bacterial community changes in enrichment cultures studied by 454-

pyrosequencing

Several cultures from different enrichment stages (G0, G2, and G4) were selected

together with a sediment sample from Chile site 7155 (depth of 437–442 cmbsf) to

study the development of the microbial community by using 454-pyrosequencing.

Various samples were taken at different time points as specified in Table 13. For the

case of titanium (III) citrate-reduced cultures: G0–c was prepared with 9% of sediment

inoculum and completely transformed 1,2,3-TCB to 1,3-DCB within 58 days of

Results

incubation (Figure 21); G2 and G4 subcultures transformed 1,2,3-TCB within 53 and

149 days, respectively (Table 13). For the case of sodium sulphide and L-cysteine-

reduced cultures: G0–g was prepared with 6% of sediment inoculum and formed 1,3-

DCB within 42 days, and depleted 1,2,3-TCB after 58 days (Figure 22). The G2

subculture formed 1,3-DCB within 20 days. G2 had 5 μM of 1,2,3-TCB after 61 days

when sampled.

Table 13. Samples used for the study of the microbial community by 454-pyrosequencing.

Reducing agent in culture none titanium (III) citrate Na2S and L-cysteine

Sample origin sediment G0–c G2 G4 G0–g G2

Time 1,2,3-TCB depletion

(days of incubation)

n.a. 58 53 149 58 >61k

Time points of sampling

for 454-pyrosequencing

(days of incubation when

sampled)

n.a. 3 a 78 a 1 b 3 a 4 a

36 a 31 b 36 a 61 a

74 a 149 b 74 a

238 a 238 a

n.a. – not applicable; a A sample of 1 ml was withdrawn from the culture b A sample of 1.5 ml was withdrawn

from the culture. kComplete depletion of 1,2,3-TCB in the culture could not be monitored as it was transferred

after 65 days.

The microbial community was studied by amplifying bacterial and archaeal 16S rRNA

genes with two sets of primers (Table 14). Five replicates from parallel PCRs were

pooled for each sample. The total number of bacterial and archaeal sequences obtained

after the 454-pyrosequencing are detailed in Table 14.

Table 14. Sequences obtained from the 454-pyrosequencing of sediment and sediment cultures.

Primer pair1 Target group Size of amplified

PCR product

(bp)

Average size of

sequence after

trimming2 (bp)

Total number

of sequences

Total number of

sequences per

Domain

27F and 519R Bacteria 599 300 44019 44019

U789F and U1068R Bacteria and Archaea 373 286 28202 Archaea: 26

Bacteria: 28176

1Sequences are detailed in Table 7; 2 The trimming was done for quality control purposes of the sequences with the program

mothur as described in the Material and Methods section.

The results from samples amplified with the primers 27F and 519R are described in this

section. Those amplified with the universal primers, U789F and U1068R, are described

in the next section (3.2.5).The number of bacterial sequences obtained in total and for

the phyla Firmicutes, Proteobacteria, and Chloroflexi for each sample is shown in

Table 15.

Results

Table 15. Number of sequences obtained for total Bacteria, phyla Firmicutes, Proteobacteria, and

Chloroflexi after 454-pyrosequencing as classified by mothur in samples of sediment and sediment

cultures.

Sample Number of sequences,

total Bacteria

Number of sequences,

phylum Firmicutes

Number of sequences,

phylum Proteobacteria

Number of sequences,

phylum Chloroflexi

sediment 2895 7 2884 0

TiCia G0 – 3 days 2430 1767 595 7

G0 – 36 days 636 596 31 0

G0 – 74 days 1006 902 28 0

G0 – 238 days 1485 1334 71 1

G2 – 78 days 3999 3999 0 0

G4 – 1 day 8610 8601 7 0

G4 – 31 days 7059 7059 0 0

G4 – 149 days 2657 2657 0 0

Na2Sb G0 – 3 days 2759 684 2029 6

G0 – 36 days 1199 1084 91 0

G0 – 74 days 708 659 32 2

G0 – 238 days 293 279 8 0

G2 – 4 days 360 144 158 0

G2 – 61 days 2924 2233 280 0

a Titanium (III) citrate-reduced cultures; b Na2S and L-cysteine-reduced cultures.

The bacterial community structure experienced a clear shift from a dominance of the

phylum Proteobacteria, class Gammaproteobacteria, in the sediment, to a dominance

of the phylum Firmicutes in the cultures, either reduced with titanium (III) citrate or

with sodium sulphide and L-cysteine.

A shift to 75% relative abundance to the Firmicutes occurred within three days of

incubation in the G0 culture reduced with titanium (III) citrate (Figure 29). The

dominance of Firmicutes was maintained in further subcultures (G2 and G4), where the

100% of the bacterial community was affiliated with the Firmicutes. Other phyla such

as Deinococcus-Thermus, Actinobacteria, Aquificae, and Bacteroidetes were also

present in the G0 culture, although at considerably lower relative abundance (≤ 5%)

than Proteobacteria and Firmicutes. The phylum Deinococcus-Thermus was the third

most abundant phyla, with a maximum relative abundance of 5% at the time point of 74

days of incubation (Figure 29).

Results

Figure 29. Bacterial community analysis of titanium (III) citrate-reduced cultures. Bacterial community

structure is shown as relative proportions of phyla in sediments from Chile, site 7155 (“Sed”), and in

sediment culture G0 and subcultures G2 and G4 from the Chile sediment. The labels indicate the

subculture (see Figure 23) and the incubation time in days.

Higher diversity of Firmicutes was observed in G0 culture than in G2 or G4

subcultures, with members affiliated to the genera Gracilibacter, Heliorestis,

Desulfotomaculum, Thermincola, Caldalkalibacillus, and Geosporobacter being present

(Figure 30). In G0, a strong shift was observed for the first 36 days of incubation. After

36 days of incubation, a stable bacterial community was established with variations only

in the relative abundance of the different genera (Figure 30). In subcultures G2 and G4,

Anaerobacter sequences were dominant, forming 90% of the bacterial community

(Figure 30). The remaining 10% of the sequences were affiliated with the Clostridium,

Desulfosporosinos, Tepidanaerobacter and Gracilibacter.

Results

Figure 30. Relative abundance of Firmicutes in cultures reduced with titanium (III) citrate and sediments

from Chile, site 7155. Shown proportions are at the genus level for the phylum Firmicutes. Phyla other

than the Firmicutes are indicated by the “other phyla” proportion in the stacked-bar section in grey. The

proportion for “other Firmicutes” refers to genus within the Firmicutes at presence lower than 1%. For

details on the labels and the cultures see the legend of Figure 29.

More in depth analysis of sequences from within the phylum Firmicutes showed several

sequences closely associated phylogenetically to known organohalide-respiring

microorganisms such as Dehalobacter and Desulfitobacterium to be present in the

sediment cultures. The relative abundance of Dehalobacter sequences in G0 increased

with time from 0.1% and 0% on days 3 and 36, respectively, to 3% on day 74, when

complete 1,2,3-TCB transformation had occurred and the culture had further been

amended with 40 μM 1,2,3-TCB on day 65. However, Dehalobacter sequences

decreased to 2% on day 238 (Table 16). The number of Dehalobacter sequences was

very low or even below the detection limit in further G2 and G4 subcultures (Table 16

and Figure 30). Desulfitobacterium sequences were detected in G0, however at very low

relative abundance which did not increase with time. In subcultures G2 and G4, no

Desulfitobacterium sequences were detected (Table 16).

Results

Table 16. Relative abundance of various genera within the family Peptococcaceae in sediment and

cultures reduced with titanium (III) citrate from Chile site 7155. The relative abundance of each genus

with respect to whole bacterial community is shown. Sequences were obtained after 454-pyrosequencing

and classified by mothur in each sample using the primer pair 27F and 519R. Cultures G0 and G2 are

shown at different time points as stated in Table 13.

Sample



Sediment G0 G2 G4

Incubation days



- 3 36 74 238 78 1 31 149

Dehalobacter 0% 0.1% 0% 3% 2% 0.1% 0% 0% 0%

Desulfitobacterium 0% 0.9% 0% 0.2% 0.1% 0% 0% 0% 0%

Desulfosporosinus 0% 0.1% 0% 0% 0.1% 1% 0% 0% 3%

Desulfotomaculum 0% 2% 16% 10% 15% 0% 0% 0% 0%

Desulfurispora 0% 0.2% 0% 0.3% 0.1% 0% 0% 0% 0%

Pelotomaculum 0% 0.2% 0.5% 0.1% 1% 0% 0% 0% 0%

Thermincola 0% 0.4% 4% 3% 10% 0% 0% 0% 0%

For cultures reduced with sodium sulphide and L-cysteine, Firmicutes also took over as

the dominant bacterial phylum (Figure 31). This shift to Firmicutes dominance

happened at a slower rate than for those cultures reduced with titanium (III) citrate, with

25% of the bacterial community belonging to Firmicutes after three days of incubation

(Figure 31). After 36 days of incubation, 90% of the bacterial total community belonged

to members of the phylum Firmicutes (Figure 31).

In the G2 subcultures, the bacterial community evolved differently to the community in

cultures reduced with titanium (III) citrate, with other bacterial phyla apart from

Firmicutes also being detected. G2 subcultures showed proportions of the phylum

Proteobacteria of 44% and 10% at time points of 4 and 61 days of incubation,

respectively. In these G2 subcultures, the phylum Bacteroidetes increased from 2% to

10% for the period of time between 4 to 61 days.

Results

Figure 31. Bacterial community analysis of sodium sulphide plus L-cysteine-reduced cultures. Bacterial

community structure is shown as relative proportions of phyla in sediments from Chile, site 7155 (“Sed”),

and in culture G0 and subculture G2 from the Chile sediment. The labels indicate the subculture (see

Figure 23) and the incubation time in days.

The diversity within the phylum Firmicutes in the sulphide-reduced sediment cultures

was different to the titanium (III) citrate-reduced sediment cultures. Most importantly,

very few Anaerobacter members were observed in either G0 or G2 cultures. High

proportions of Desulfosporosinus were present in G0 (36–238 days of incubation) and

G2 cultures, together with Gracillibacter, Gracillibacillus, and Cryptaenobacter as

most dominant genera (Figure 32).

Results

Figure 32. Relative abundance of Firmicutes in cultures reduced with sodium sulphide and L-cysteine and

sediments from Chile, site 7155. Shown proportions are at the genus level for the phylum Firmicutes.

Phyla other than the Firmicutes are indicated by the “other phyla” proportion in the stacked-bar section in

grey. The proportion for “other Firmicutes” refers to genus within the Firmicutes at presence lower than

1%. For details on the labels and the cultures see the legend of Figure 31.

More in depth analysis of sequences from the family Peptococcaceae (phylum

Firmicutes), indicated the presence of Dehalobacter and Desulfitobacterium sequences

at very low relative abundance (Table 17). A stable enrichment in Desulfosporosinus

was achieved in the G0 culture with time from 0% to 34% relative abundance after 3

and 74–238 incubation days, respectively, and further on, in subculture G2 (Table 17).

Results

Table 17. Relative abundance of various genera within the family Peptococcaceae in sediment and

cultures reduced with sodium sulphide and L-cysteine from Chile site 7155. The relative abundance of

each genus with respect to whole bacterial community is shown. Sequences were obtained after 454-

pyrosequencing and classified by mothur in each sample using the primer pair 27F and 519R. Cultures G0

and G2 are shown at different time points as stated in Table 13.

Sample



Sediment G0 G2

Incubation days



- 3 36 74 238 4 61

Dehalobacter 0% 0% 0.3% 0% 0.3% 0% 0.3%

Desulfitobacterium 0% 0% 0.2% 0.4% 0.3% 0% 0%

Desulfosporosinus 0% 0% 29% 34% 34% 12% 27%

Desulfotomaculum 0% 0% 1% 0.7% 0% 0% 2%

Desulfurispora 0% 0% 0.2% 0.1% 1% 0% 0.1%

Pelotomaculum 0% 0% 0.2% 0% 0% 0% 0%

Thermincola 0% 0% 0% 0% 0.3% 0% 0%

3.2.5 Microbial community study with universal primers in the enrichment

cultures using 454-pyrosequencing

Samples from the same G0, G2, and G4 cultures as for the bacterial-specific community

study were used together with a sediment sample from Chile, site 7155 (depth of 437–

442 cmbsf) (Table 13) for the amplification of 16S rRNA genes with the universal

primers U789F and U106R, which targets both Archaea and Bacteria.

Less time points compared to the bacterial-specific community study (amplified with

primers 27F and 519R) were selected, and related to an incubation time when 1,2,3-

TCB was about to be depleted. These time points were 36 and 238 incubation days for

G0, 149 days for G4 in cultures reduced with titanium (III) citrate, and 238 days for G0

and 173 days for G2 in cultures reduced with sodium sulphide and L-cysteine.

The sequences amplified with the universal primers belonged mostly to Bacteria (Table

14). Archaeal sequences were only found in G0 cultures reduced with titanium (III)

citrate making up 0.31% of the total microbial sequences. No archaeal sequences were

amplified in sediment cultures reduced with sodium sulphide and L-cysteine, as well as

in the sediment sample. Archaea were therefore most likely not associated to the

transformation of 1,2,3-TCB in sediment cultures.

Results

The bacterial sequences amplified with the universal primers also confirmed the shift

from phylum Proteobacteria in sediment to phylum Firmicutes in the cultures as

previously observed (Figure 33). However, there were minor differences, especially that

there was a larger contribution of Nitrospirae sequences in G0 after 238 incubation days

than previously observed, and the Proteobacteria sequence contribution was also lower

for cultures reduced with sodium sulphide and L-cysteine than previously observed.

Figure 33. Bacterial relative phyla abundance of samples amplified with universal primers in sediment

and cultures from Chile, site 7155. G0 and G4 are cultures reduced with titanium (III) citrate and G0 and

G2 are cultures reduced with sodium sulphide plus L-cysteine. The labels indicate the subculture (see

Figure 23) and the incubation time in days (see Table 13). Those cultures reduced with sodium sulphide

and L-cysteine are indicated with “(Na2S)”.

Results

Within the Firmicutes, large relative shares belonged to Gracilibacter (in G0),

Tepidanaenobacter (in G4), and Sporomusa (in G0 and G2 cultures reduced with

sodium sulphide and L-cysteine) in contrast to the bacterial-specific community

analyses (Figure 34).

Figure 34. Relative abundance of Firmicutes in samples amplified with universal primers. Shown

proportions are at the genus level for the phylum Firmicutes from cultures reduced with sodium sulphide

plus L-cysteine set up with sediments from Chile site 7155. Phyla other than the Firmicutes are indicated

by the “other phyla” proportion in the stacked-bar section in grey. The labels indicate the subculture (see

Figure 23) and the incubation time in days (see Table 13). Those cultures reduced with sodium sulphide

and L-cysteine are indicated with “(Na2S)”.

Among phylotypes that were present in all samples are Desulfosporosinus and

Desulfitobacterium (Figure 34 and Table 18). The relative abundance of

Desulfosporosinus was however, less than in the bacterial-specific community analyses.

In addition, no Dehalobacter sequences were observed, in contrast to the bacterial-

specific community analyses (Figure 34 and Table 18).

Results

Table 18. Relative abundance of sequences amplified with universal primers and affiliated to specific

genera within the family Peptococcaceae in sediment and cultures from Chile site 7155. The relative

abundance of each genus with respect to whole bacterial community is shown.

Reducing agent none Titanium (III) citrate Sodium sulphide and L-

cysteine

Sample



Sediment G0 G4 G0 G2

Incubation days



- 36 238 149 238 173

Dehalobacter 0% 0% 0% 0% 0% 0%

Desulfitobacterium 0% 2% 1% 1% 0.5% 0.4%

Desulfosporosinus 0% 8% 3% 7% 0.2% 1%

Desulfotomaculum 0% 0.1% 0.1% 0% 0% 0%

Desulfurispora 0% 9% 3% 0% 0% 0%

Pelotomaculum 0% 1% 0.1% 0% 0.1% 0%

Sporomusa 0.3% 0.2% 0% 0% 84% 88%

3.3 BAFFIN BAY GEOCHEMISTRY AND MICROBIAL COMMUNITIES

This chapter describes the in situ study performed in sediments of the Baffin Bay, in the

Arctic. The goal of this study was to gain insights into the biogeochemistry and

microbial communities which may drive the major element cycles (i.e., carbon, sulphur,

iron, manganese) in sediments of the Baffin Bay. For that purpose, sediments were

cored at 34 sites, as previously described (section 2.2). Ten sites were selected based on

the site location and the recovery of the core after sampling (highest recovery length of

4.69 m), and further sampled for the site- and depth-dependent geochemistry and

microbiology study (described in section 2.2.2). These ten sites were from various

locations of the Greenlandic part of the Baffin Bay, which included the continental

shelf, slope and basin areas. Sites 363, 365, 371 belonged to the Northern continental

shelf (Figure 4 and Figure 5). Site 383 was located in between the shelf and the

beginning of the continental slope (Figure 5). Site 387 belonged to the continental slope

(Figure 5). Sites 389, 391, and 453 were located at the central deep basin, and sites 486

and 488 were at a Southern continental slope (Figure 4 and Figure 5). In this chapter,

the geochemistry (including the geology, and the solid- and interstitial-phase

geochemistry of sediments; section 3.3.1), and microbiology (microbial ecology

deciphered with molecular biology techniques; section 3.3.2) of the cores from the

selected ten sites of the Baffin Bay are described.

Results

3.3.1 Geochemistry of Baffin Bay cores

An overview of the oceanographic data of the ten investigated sites and the

corresponding sediment recoveries of each core is presented in Table 19.

Table 19. Oceanographic data of the selected sites in the Baffin Bay investigated in this study. The

corresponding core recovery after the sediment sampling at each site is presented as “core depth”. Apart

from the geochemistry analysis, each core was further analysed for either one or two microbiology

studies, named study A (including three geographically distinct areas: shelf, basin, and Southern slope),

and study T (including sites located along a transect from shelf to basin).

Site

Area

Study

Latitude

Longitude

Water Depth (m)

Core depth (m)

363

Shelf

A & T

76° 52.92' N

71° 34.01' W

938

4.69

365

Shelf

76° 39.04' N

71° 18.79' W

658

3.67

371

Shelf

A & T

75º 58.24' N

70° 34.86' W

598

4.05

383

Shelf-slope

75º 17.69' N

69º 53.75' W

674

2.32

387

Slope

74° 50.42' N

69º 27.14' W

1,300

3.32

389

Basin

A & T

74° 37.05' N

69º 13.75' W

1,716

4.24

391

Basin

A & T

74° 23.36' N

69º 01.22' W

1,864

4.27

453

Basin

73° 19.37' N

64° 58.11' W

2,300

4.69

486

S Slopea

72° 24.51' N

60° 48.85' W

645

4.69

488

S Slopea

72° 08.80' N

60° 58.86' W

1,493

4.69

aSouthern Slope

The geology of the cores varied among the different sites (see core photos in the

Appendix 2, section 6.2) indicating changes in the local conditions. In general, many

dropstones were found in the cores as a result of glacial transport.

Cores from sites 363 and 371 had similar sediment composition of olive-grey silty muds

along the entire core length, except in deeper parts where a change in colour to greyish-

brown occurred. In deeper sections of the core from site 371 (346–395 cmbsf),

sediments became coarser (sandy and silty mud with dropstone granules) and changed

to a dark-brown colour.

Shelf site 365 had olive-grey silty muds in the core section from the surface down to

~200 cmbsf. At increased depths, site 365 sediment became gradationally coarser to

silty muds with sand granules (200–274 cmbsf) and to muddy sands (274–322 cmbsf).

A gradational change of colour from the olive-grey to brownish-grey was observed at

150 cmbsf for site 365, which turned to greyish-red (185–192 cmbsf), and to a distinct

red colour lamination (247–260 cmbsf), and finally, to brown in deeper sediments (300

to circa 322 cmbsf).

Results

The core from site 383, a transition site from shelf to slope areas, was composed of

olive–grey silty muds in surface sediments, which turned to brown silty muds (core

sections of 46–57 cmbsf, and 112–154 cmbsf) and to brownish-black sandy and silty

muds (57–100 cmbsf, and 154 to 232 cmbsf).

Shallow sediment layers from basin sites 389, 391, 453 and slope site 387 had diatom-

bearing silty muds, which extended from the surface to 64–63 cmbsf for sites 389 and

391, to 100 cmbsf for site 453, and to 21 cmbsf for slope site 387. Sites 387, 389 and

391 had increased composition of sand in their sediments, with alternating layers of silty

muds and sandy muds throughout the three cores. Changes in sediment colour were

observed along the core lengths in sites 387, 389 and 391. Site 387 sediment colour

changed from yellowish-brown (21–52 cmbsf) to a layer where a transition from

reddish- to brownish-colours were observed (52–71 cmbsf), and to brown (71–332

cmbsf). Site 389 changes in sediment colour were mainly in the shallower 200 cmbsf,

where a gradational change from yellowish-brown to olive-grey was observed at 64–

100 cmbsf, followed by a layer of olive-grey sediments (100–145 cmbsf), which

changed to brown (145–166 cmbsf) and to greyish brown (166–424 cmbsf). Site 391

sediments changed from yellowish-brown to olive-grey (63–97 cmbsf) to dark greenish-

grey (97–100 cmbsf), to a layer of dark yellowish-brown to light olive-grey (100–120

cmbsf), to pale brown (120–123 cmbsf), moderate brown (123–142cmbsf), brownish-

grey (145–200 cmbsf), and to brownish-black (200–420 cmbsf). Site 453 sediments

were composed of silty muds (100–300 cmbsf) with intercalations of clayey silt and

sand at section 200–273 cmbsf. From 300 to 430 cmbsf, sediments contained increased

concentrations of carbonate, with a layer of carbonate-rich brownish-grey silty marl at

300–317 cmbsf. Silty marls were observed from 300 to 430 cmbsf at site 453. At deeper

sediment layers (430–470 cmbsf) of site 453, dark-olive silty muds were observed.

Southern slope sites 486 and 488 were composed of olive-grey silty muds throughout

the entire cores, which eventually changed colour to more brownish in some sections of

both cores.

The main minerals composing the mineral fraction of the sediments for three analysed

sites (363, 389 and 486) were quartz and feldspar. In deeper layers of site 363, the main

mineral fraction changed to be composed of only quartz (454 cmbsf) and quartz with

dolomite (460 cmbsf). The mineral composition of site 389 changed from quartz and

feldspar (51–192 cmbsf; except layers at 129 and 157 cmbsf that were composed of

only quartz) to quartz and feldspar together with dolomite and calcite at 216 cmbsf,

followed by layers of quartz and calcite (240–304, and 358 cmbsf; except a intercalated

Results

layer of quartz and feldspar at 320 cmbsf) and only quartz in deepest layers (387–

409 cmbsf).

The chemical composition of the sediment was of various oxides (50% silica, 13%

alumina, 4% magnesia, 3–7% calcium oxide, 2–4% sodium oxide, 3% potassium

oxide). Among them, iron(III) oxide values of 5% and 7% were observed for sites 363

and 486, respectively. Site 389 had an iron(III) oxide value of 6–7%, except at

sediments layers of 187–192 cmbsf where the value decreased to 3–4% (see Appendix

3, section 6.3). A manganese(II) oxide value of ~ 0.1% was observed in all sediments

analysed expect at a depth of 51 cmbsf from site 389, where a value of 0.7% was

observed (see Appendix 3, section 6.3).

Values for the total organic carbon (TOC) in samples from shelf sites 363, 371 and the

uppermost 125 cmbsf of 365 were higher than from other sites (Figure 35). The highest

TOC value was 3% at surface sediments of site 371. Sites from the continental slope

and central deep basin had similar TOC values around 0.5–1%. Lowest TOC values

were found in cores 453, 486, and deepest sections of the shelf site 365. TOC decreased

gradually with depth generally for all cores except for site 391, which at 175 cmbsf

increased to values higher than 1.5%, which were maintained until the deepest part of

the core.

In addition, total carbon (TC) was measured and the values indicated a contribution of

carbonates at some sediment layers of sites 383, 389, 391, and 453. The highest TC

value corresponded to site 453 at 277 cmbsf (Figure 36), which had a contribution of

3% for carbonates. Sediment layers from depths of 122–415 cmbsf at the site 389 had

carbonate contributions of ~1.5%. Sediment layers at 122–214 and 221–410 cmbsf for

sites 383 and 389, respectively, had carbonate contributions of ~ 1%.

The stable isotopic composition of organic carbon, measured as δ13Corg calibrated vs.

the VPDB standard, was measured from the sites 363, 389 and 486. Shelf site 363 had

average δ13Corg values of -25‰, which became lighter (-26 – -27‰) in deepest core

sections (Figure 37). Sites 389 and 486 had lighter overall δ13Corg values than site 363.

Site 389 had average δ13 Corg values of -28‰, with lighter values of -31‰ at core

sections of 130 cmbsf, and heavier values of -25‰ at deepest sections of the core (~ 400

cmbsf). Site 486 had average δ13C values of -28 ‰, which became lighter with depth.

Results

Figure 35. Total organic Carbon (TOC) values for sediments from the Baffin Bay. Shown are depth

profiles of all the sites analysed within this study. The symbol and colour code for each site are

consistently used in all graphs of this thesis.

Figure 36. Total Carbon (TC) values of sediment sites from the Baffin Bay. Shown are depth profiles of

all sites analysed within this study.

Results

Figure 37. Organic carbon isotopic composition of sediment from the Baffin Bay. Shown are depth

profiles of three sites that were selected, one site per area (shelf, basin, slope).

Pore-water constituents of sediments from the Baffin Bay included the ions sodium,

chloride, potassium, magnesium, sulphate, iron(II), and manganese(II).

Sodium and chloride depth-dependent concentrations were stable with mean values of

461 mM ± 10 mM and 534 mM ± 13 mM, respectively.

Potassium concentrations of 10 mM were observed in near-surface sediments, which

decreased to 8−9 mM with increasing depth.

Magnesium depth-dependant concentrations were stable at 45−50 mM in sediments

from all sites, except for sites 363 and 453, where concentrations decreased with

increasing depth (results not shown; for details see (Algora et al 2013)).

Sulphate concentrations were 25−27 mM in near-surface sediments and decreased

gradually with increasing depths except for sites 365, 383, 389, and 391, which

remained stable (Figure 38). Pore-water at sites 363 and 453 had highest decreases in

sulphate concentrations with increasing depths, reaching 18 mM and 4 mM,

respectively, at 450 cmbsf (Figure 38).

Results

Figure 38. Sulphate concentration in pore-water of sediments from the Baffin Bay. Shown are depth

profiles of all the sites investigated within this study.

Iron(II) was present in the pore-water from all sites as shown by the depth profiles

(Figure 39). However, the concentration of Fe(II) differed substantially among the

various sites and depths. At shelf sites 363, 365, and 371, Fe(II) concentration increased

with increasing depth up to 32 μM for site 363, and up to 57 μM for sites 365 and 371,

at the deepest parts of the cores. However, Fe(II) concentration increased at higher rates

at site 365 (29 μM were measured at 175 cmbsf), compared to sites 371 and 363 (Figure

39). Fe(II) concentration decreased with increasing depth at site 383, except at 175

cmbsf. However, site 387 showed increasing concentration of Fe(II) with increasing

depth reaching 58 μM. At the basin sites 389 and 391, Fe(II) concentration increased

with increasing depth. The maximum Fe(II) concentration was 27 μM for site 389,

observed at the depths of 75 and 225 cmbsf. For site 391, a substantially high value

(maximum of all measured values) of 138 μM at 175 cmbsf was detected (Figure 39).

Fe(II) concentration at site 453 increased with increasing depth to 15 μM. At Southern

slope sites 486 and 488, near-surface Fe(II) concentration of 28 µM and 47 µM were

detected, respectively. At both sites, the concentration of Fe(II) decreased with

increasing depth to subsequently increase at deeper core sections (Figure 39).

Results

Figure 39. Iron(II) concentration in pore-water of sediments from the Baffin Bay. Shown are depth

profiles of all the sites investigated within this study.

Manganese(II) was detected in the pore-water of all sites, however, with high

variabilities in its concentration among the various sites and depths, which resulted in

distinct trends in depth profiles (Figure 40). The concentration of Mn(II) in depth

profiles from shelf sites 363 and 371 slightly increased with increasing depth (Figure

40). Similarly, the Mn(II) concentration in shelf site 365 increased with depth, however

at higher rates, reaching a maximum concentration of 23 μM at 225 cmbsf. The Mn(II)

concentration in the depth profiles of sites 383 and 387 were stable at a value of 15–

20 μM, except at surficial sediments where 39 μM and 3 μM was observed,

respectively.

The sediment profiles of sites 389 and 391 indicated a pronounced increase in Mn(II)

concentration at 75–175 cmbsf, especially for site 389, where Mn(II) concentrations

reached a maximum of 115 μM. At both sites, the Mn(II) concentration decreased to

~20 µM at 225/275 cmbsf and remained stable with increasing depth (Figure 40). The

concentration of Mn(II) at site 453 increased with increasing depth to 16 μM at 375

cmbsf, to subsequently slightly decrease at deepest core sections. The Mn(II)

concentration at site 486 was stable at ~20 μM with depth, and at site 488 decreased

with increasing depth from 50 μM (at 25 cmbsf) to 24 μM (at 450 cmbsf).

Results

Figure 40. Mn(II) concentration in pore-water of sediments from the Baffin Bay. Shown are depth

profiles of all the sites investigated within this study.

Methane was detected at all sites, however with no specific trend in the depth profiles.

Methane concentration values were in general low with a maximum concentration of

17 µM at site 486 (Figure 41). Deepest core sections (225/275–425 cmbsf) of Southern

slope sites 486 and 484 showed highest methane concentrations of ~15 µM, followed by

shelf sites 363 and 365, with methane concentrations of ~11 µM (Figure 41).

Figure 41. Methane concentration in sediment depth profiles from cores of the Baffin Bay.

Results

3.3.2 Microbial ecology of Baffin Bay cores

The microbial ecology study investigated i) the microbial composition present in

sediments of the Baffin Bay, ii) if differences in the microbial composition existed

among the various sediment sites, and iii) if these differences in the microbial

composition may be associated to changes in environmental parameters, i.e., sediment

geochemistry, depth, and geographical location within the Baffin Bay.

The microbial composition was studied in terms of the abundance and spatial

distribution of specific microbial groups, the microbial community structure and

diversity. For that, two approaches were designated, the first one named “study A”

focused on the abundance and spatial distribution of Bacteria, Archaea and specific

microbial groups, e.g., Dehalococcoidia, in three geographically distinct areas: Northern

Greenlandic shelf, central deep basin, and a Southern slope (Figure 4). The second

approach was named “study T” and focused on the microbial community structure and

diversity in sediments along a North-to-South, shelf-to-basin transect. This transect was

267 km long, from the Northern Greenlandic shelf to the central deep basin of the

Baffin Bay (Figure 5).

Study A: Microbial composition in three distinct areas of the Baffin Bay

The abundance and site- and depth-dependant distribution of microbial populations

were investigated in a total of seven sediment sites from the areas: Northern

Greenlandic shelf (sites 363 and 371, Figure 4), central deep basin (sites 389, 391, and

453, Figure 4), and Southern slope (sites 486 and 488, Figure 4). This study involved

the quantification of total Bacteria and Archaea, as well as specific microbial

phylogenetical groups, i.e., the class Dehalococcoidia and the order

Desulfuromonadales, and specific microbial physiological groups, i.e., sulphate-

reducers, and methanogens. Quantification of Bacteria, Archaea, Dehalococcoidia and

Desulfuromonadales was performed by qPCR using primers targeting the 16S rRNA

gene (Table 7). Quantification of the physiological groups of sulphate-reducers and

methanogens was done by qPCR with primers targeting the functional genes dsrA and

mcrA (Table 7).

 Quantification of total Bacteria and Archaea in sediment cores

In general, highest 16S rRNA gene copy numbers for both Bacteria and Archaea were

observed in uppermost sediments for all sites, except for site 486, where the highest

number of bacterial copies (7.9 x 107 g-1) was detected at 120 cmbsf. Similar patterns in

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the depth profiles were observed for both archaeal and bacterial 16S rRNA gene copy

numbers.

In the Northern Greenlandic shelf sites 363 and 371, 16S rRNA gene copy numbers for

Bacteria were stable with increasing depth, averaging 1.4 x 107 ± 4 x 106 for site 363 at

25–430 cmbsf, and 2.3 x 107 ± 1.8 x 107 for site 371 at 22–271 cmbsf. Similar depth

profile patterns were observed for archaeal 16S rRNA gene copies at both shelf sites.

Archaeal gene copy numbers were in average 2.1 x 107 ± 8.4 x 106 for site 363 at 25–

430 cmbsf, and 1.5 x 107 ± 4.11 x 106 for site 371 at 22–271 cmbsf. In core sections

deeper than 300 cmbsf for site 371 and deeper than 425 cmbsf for site 363, both

bacterial and archaeal copy numbers decreased one or two orders of magnitude (Figure

42).

In the central deep basin sites, bacterial copy numbers of 4.1 x 107 and 6.6 x 107 g-1

were detected at ~25 cmbsf in sites 391 and 389, respectively (Figure 43). Bacterial

copy numbers rapidly decreased down to three orders of magnitude with increasing

depth, and became in average 2.2 x 104 g-1 at 221–410 and 172–415 cmbsf for sites 389

and 391, respectively (Figure 43). Similarly to Bacteria, archaeal copy numbers

declined rapidly and steadily with increasing depth at sites 389 and 391 (Figure 43). The

highest archaeal copy number of all sites was 1.8 x 108 g-1 and was detected at 20 cmbsf

at site 389. Archaeal copy numbers at site 389 decreased to 8.4 x 106 g-1 at 75 cmbsf and

to 8 x 105 g-1 at 122 cmbsf. A similar pattern was found at site 391, with highest

archaeal copy numbers of 5.4 x 106 g-1 at 28 cmbsf that declined to 2.9 x 104 at 122

cmbsf. In general, no amplification was obtained in the qPCR assay for Archaea in both

sites 389 and 391 at sediments deeper than 175 cmbsf, indicating that the presence of

Archaea at increasing depths was very rare.

At site 453, the bacterial and archaeal depth profiles differed substantially from the

other central deep basin sites. Bacterial copy numbers were in average 2.3 x 106 ± 1.6 x

106 g-1 at site 453. An order of magnitude decrease in bacterial copy numbers (1.9 x 105

g-1) was observed at 277 cmbsf (Figure 43), where a high percentage of carbonates in

the sediment had been detected. Archaeal copy numbers at site 453 were one order of

magnitude lower than the bacterial copy numbers, and could be detected throughout the

core, except at 22 cmbsf (Figure 43).

In the site 486 from the Southern slope, bacterial copy numbers of 1 x 107 g-1 were

observed at 20 cmbsf and increased to a maximum value of 7.9 x 107 g-1 at 120 cmbsf

(Figure 44). At 173–321 cmbsf, bacterial copy numbers remained stable with increasing

depth, averaging 1.2 x 107 ± 1.2 x 106 g-1. Bacterial copy numbers decreased at layers

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deeper than 321 cmbsf down to a minimum number of 1.6 x 106 g-1 by 448 cmbsf

(Figure 44).

Site 488 from the Southern slope showed a maximum bacterial copy number of 5.6 x

106 g-1 at 70 cmbsf, and a minimum of 2.8 x 105 g-1 at 171 cmbsf. Surface sediments

(20–70 cmbsf) had considerable higher bacterial copy numbers, nearly one order of

magnitude, than the deeper core sections (Figure 44).

Archaeal copy numbers were generally one order of magnitude lower than bacterial at

Southern slope sites 486 and 488, in contrast to the shelf and basin areas, where similar

bacterial and archaeal copy numbers were observed. The only exception was surface

sediments of site 488, and deep sediments (421 cmbsf) of site 486, where similar

archaeal and bacterial copy numbers were found (Figure 44).

Results

Figure 42. Depth profiles of total Bacteria (upper panel) and Archaea (lower panel) quantified as 16S

rRNA gene copy numbers per gram of sediment (wet weight) for the sites 363 and 371 within the

Northern Greenlandic shelf area of the Baffin Bay.

Results

Figure 43. Depth profiles of total Bacteria (upper panel) and Archaea (lower panel) quantified as 16S

rRNA gene copy numbers per gram of sediment (wet weight) for the sites 389, 391, and 453 within the

central deep basin area of the Baffin Bay.

Results

Figure 44. Depth profiles of total Bacteria (upper panel) and Archaea (lower panel) quantified as 16S

rRNA gene copy numbers per gram of sediment (wet weight) for the sites 486 and 488 within the

Southern slope area of the Baffin Bay.

Results

 Quantification of class Dehalococcoidia and order Desulfuromonadales in

sediment cores

Members from the class Dehalococcoidia were quantified with the primers DEH-Fa and

DEH-R targeting the 16S rRNA gene (Table 7). From the three areas under

investigation, highest Dehalococcoidia 16S rRNA gene copy numbers, of 3.7 x 105 ±

2.8 x 105 g-1 in average, were detected in sediment sites 363 and 371, from the Northern

Greenlandic shelf (Figure 45). Lowest Dehalococcoidia copy numbers, of 5 x 104 g-1 ±

3.7 x 104 in average, were found at Southern slope site 486 (Figure 47).

In the central deep basin, highest Dehalococcoidia copy numbers were observed in

shallow sediments, with 4 x 105 g-1 at depths of 22 and 78 cmbsf at sites 453 and 391,

respectively (Figure 46). At site 453, similar Dehalococcoidia copy numbers with

increasing depth (1.7 x 105 ± 1.3 x 105 in average) were found. However, sites 389 and

391 had pronounced decreases in the number of Dehalococcoidia copies with increasing

depth (Figure 46). At site 389, the minimum value (1.1 x 103 g-1) of Dehalococcoidia

copy numbers from all sites and depths was detected at 221 cmbsf.

Even though shelf sites 363 and 371 showed highest Dehalococcoidia copy numbers,

with a maximum of 8.4 x 105 g-1 at site 363, and depth of 78 cmbsf, the relative

proportion of Dehalococcoidia with respect of the total bacterial 16S rRNA gene copy

numbers (calculated as 16S rRNA gene copy numbers of Dehalococcoidia divided by

total bacterial 16S rRNA gene copies), was fewer than 10%, with the only exception of

site 371, at 374 cmbsf, where Dehalococcoidia accounted for 29% of the total bacterial

16S rRNA gene copy numbers (Figure 48). In contrast, sites 389 and 391, in the central

deep basin, had low Dehalococcoidia copy numbers at 150–400 cmbsf, nevertheless,

Dehalococcoidia accounted for more than 50% for site 391 and between 11–51% for

site 389 (Figure 48).

Southern slope site 488 had a distinct depth profile of Dehalococcoidia copy numbers,

which differed from the rest of the sites, as numbers increased more than one order of

magnitude in deepest core sections (423–448 cmbsf) compared to shallower sediments

(20–171 cmbsf). Highest Dehalococcoidia copy numbers of 6.4 x 105 g-1 were detected

at 448 cmbsf, where Dehalococcoidia accounted for 21% of the total bacterial copy

numbers (Figure 47 and Figure 48).

The order Desulfuromonadales includes known members of Bacteria able to reduce

insoluble Fe(III) and Mn(IV) oxides such as Geobacter spp. and Desulfuromonas spp.

The primers GEO494F and GEO825R (Table 7) targeted the order Desulfuromonadales

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and were used here as an approximation to provide information on potential metal

reducing bacteria in the sediments of the Baffin Bay.

Desulfuromonadales 16S rRNA gene copy numbers were in general one order of

magnitude less abundant than bacterial and archaeal copy numbers (Figure 45, Figure

46, and Figure 47). The depth profile for Desulfuromonadales exhibited similar patterns

to the total bacterial and archaeal copy numbers, and did not relate to the concentration

of iron(II) and manganese(II) in the pore-water. For instance, at deep core sections of

454 and 323–374 cmbsf at shelf sites 363 and 371, respectively, the concentration of

iron(II) notably increased (Figure 39); however, the copy numbers of

Desulfuromonadales decreased two orders of magnitude (Figure 45). Similarly,

substantial high concentrations of iron(II) at 175 cmbsf of site 391, and of

manganese(II) at 75–175 cmbsf of site 389 were observed (Figure 39 and Figure 40);

however, at those depths and sites, no Desulfuromonadales could be detected (Figure

46). Desulfuromonadales accounted for more than 20% of the total 16S rRNA bacterial

sequences in near-surface shelf sediments, where a low concentration of iron(II) or

manganese(II) was measured (Figure 39, Figure 40, and Figure 48).

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Figure 45. Depth profiles of the class Dehalococcoidia (upper panel) and order Desulfuromonadales

(lower panel) quantified as 16S rRNA gene copy numbers per gram of sediment (wet weight) for the sites

363 and 371 within the Northern Greenlandic shelf area of the Baffin Bay.

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Figure 46. Depth profiles of the class Dehalococcoidia (upper panel) and order Desulfuromonadales

(lower panel) quantified as 16S rRNA gene copy numbers per gram of sediment (wet weight) for the sites

389, 391, and 453 within the central deep basin area of the Baffin Bay.

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Figure 47. Depth profiles of the class Dehalococcoidia (upper panel) and order Desulfuromonadales

(lower panel) quantified as 16S rRNA gene copy numbers per gram of sediment (wet weight) for the sites

486 and 488 within the Southern slope area of the Baffin Bay.

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Figure 48. Relative proportion of the 16S rRNA gene copy numbers of class Dehalococcoidia (upper

panel) and order Desulfuromonadales (lower panel) to the total 16S rRNA gene copy numbers of

Bacteria for all sites of the Baffin Bay investigated in this study.

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 Quantification of functional genes (dsrA, mcrA) in sediment cores

The functional genes dsrA and mcrA encode the enzymes dissimilatory sulphite (bi)-

reductase subunit A (dsrA) and the α-subunit of the methyl coenzyme M reductase

(mcrA). The qPCR analysis of dsrA and mcrA genes was used here as a proxy to

quantify the physiological microbial groups of sulphate reducers and methanogens/

anaerobic methanotrophs, respectively.

dsrA genes were detected in sediment sites from the Northern Greenlandic shelf and the

Southern slope at all depths except at 374 cmbsf of site 371 (Figure 49 and Figure 51).

However, at central deep basin sites 389 and 391, dsrA genes were only detected in

shallow sediments (20–122 and 28–78 cmbsf at sites 389 and 391, respectively; Figure

50). dsrA genes at site 453 were detected at 123–374 cmbsf (Figure 50).

The number of dsrA copies was highest in the Northern Greenlandic shelf than in any

other area, with maximum dsrA copies of 1.4 x 109 g-1 at site 371 and a depth of 22

cmbsf, which decreased with increasing depths down to a minimum of 1.2 x 107 g-1 at

323 cmbsf (Figure 49). In central deep basin sites 389 and 391, dsrA gene copies were

highest in shallowest sediments (Figure 50). dsrA gene copies at site 453 were the

lowest of all sites.

In the Southern slope site 486, dsrA copy numbers remained generally stable with

depth, at a value of 8.6 x 106 ± 3.3 x 106 g-1 in average (Figure 51). However, at site 488,

dsrA copy numbers varied with increasing depth, from an average value of 3.4 x 106 ±

1.6 x 106 g-1 in the uppermost 75 cmbsf, to a minimum value of 7.8 x 104 g-1 at 121, and

to an average value of 5.3 x 105 ± 2.2 x 105 at deep core sections of 171–448 cmbsf

(Figure 51).

In general, few mcrA gene copy numbers were observed in any of the sites from all

areas. The maximum number of mcrA gene copies was 2.9 x 106 g-1, observed at a depth

of 20 cmbsf at site 389 (Figure 50.). Lowest mcrA gene copies were detected at sites

453 and 488 (Figure 50 and Figure 51). Generally, depth profiles for mcrA gene copies

were similar to those of dsrA. However, mcrA gene copies were one or two order of

magnitude less abundant than dsrA (Figure 49, Figure 50, and Figure 51).

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Figure 49. Depth profiles of functional genes mcrA and dsrA of methanogens/anaerobic methanotrophs

and sulphate reducing prokaryotes, respectively, per gram of sediment (wet weight) for two selected sites

within the Northern Greenlandic shelf area within the Baffin Bay.

Figure 50. Depth profiles of functional genes mcrA and dsrA of methanogens/anaerobic methanotrophs

and sulphate reducing prokaryotes, respectively, per gram of sediment (wet weight) for two selected sites

within the central deep basin area within the Baffin Bay.

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Figure 51. Depth profiles of functional genes mcrA and dsrA of methanogens/anaerobic methanotrophs

and sulphate reducing prokaryotes, respectively, per gram of sediment (wet weight) for two selected sites

within the Southern slope area within the Baffin Bay.

Study T: Microbial composition in sediments along a shelf to basin transect

The microbial community structure and diversity was investigated using high-

throughput Illumina sequencing in a total of seven sites (363, 365, 371, 383, 387, 389,

and 391; Figure 5) along a North-to-South, shelf-to-basin transect. Several samples

from various depths were selected for each site (five from site 363, seven from site 365,

four from site 371, three from site 383, seven from site 387, five from site 389, and nine

from site 391), forming a total of 40 samples (see right-hand legend in Figure 52, where

site_depth in cmbsf is shown). The bacterial community structure was determined as

relative abundance of bacterial taxonomical groups at the phylum and/or class and/or

order level present. The diversity was determined as numbers of operational taxonomic

units (OTUs). In addition, the correlation of specific bacterial groups to geochemical

parameters, i.e., TOC content, and concentration of sulphate, iron(II), and

manganese(II), was also investigated.

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 Bacterial community structure analysed by Illumina sequencing

The Illumina sequence analysis of the 40 samples yielded a total of 113,120 sequences

(2,828 sequences per sample) after quality filtering and normalising the number of

sequences in any sample to the lowest number of sequences found in a sample. The

sequences had a length of ≥110 bp and were taxonomically classified using the SILVA

and Greengenes databases via the naïve Bayesian classification method (Wang et al

2007).

The phylum Proteobacteria was dominant within the bacterial community at all sites

and depths. More in-depth classification of the proteobacterial sequences showed that

38–64% of the total bacterial community belonged to the class Betaproteobacteria, and

6–13% to the class Alphaproteobacteria (Figure 52). Other classified groups within the

phylum Proteobacteria, although in minor relative abundance, were the classes

Deltaproteobacteria and Gammaproteobacteria. The betaproteobacterial sequences

affiliated mostly (average of 99.7%) to the order Burkholderiales (Figure 52).

The second most abundant phylum was Chloroflexi accounting for 8–22% of the total

bacterial community (Figure 52). More in-depth classification indicated that 75% of

these Chloroflexi belonged to the class Dehalococcoidia. Relative abundance for

Dehalococcoidia was 6%–17% of the total bacterial community. The third most

abundant phylum was the Actinobacteria (Figure 52).

Hierarchical clustering of the analysed bacterial community structure in all the samples

resulted in two main clusters of samples showing highest similarities (dendrogram at the

right part of Figure 52, shaded in grey). One cluster (upper cluster in Figure 52)

included near-surface sediments of 25 cmbsf from all sites, and deeper layers from shelf

sites 363, 371, and 365, as well as two samples from basin sites 391 and 389 at 125 and

410 cmbsf, respectively. In the lower cluster (in Figure 52), there were mostly deeper

layer samples from basin and slope sites and the shelf sites 365, 371, and 383. One of

the main differences between the two clusters was the higher relative abundance of the

phylum Chloroflexi in the samples from the upper cluster. Indeed, highest relative

abundance of the class Dehalococcoidia was present in samples of the shelf sites 363

and 371 at 425 and 225 cmbsf, respectively. Samples in the lower cluster showed higher

relative abundance of the class Betaproteobacteria and specifically from the order

Burkholderiales than the first cluster (Figure 52).

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Figure 52. Relative abundance of classified bacterial sequences in samples (bar plot, with colour-code legend on the left) and the associated hierarchical clustering (UPGMA,

right and shaded in grey) calculated using the Bray-Curtis dissimilarities. The scale below the bar plot indicates relative abundance in samples and in total. Sediment sample

names indicate site and depth in cmbsf where the sample is coming from (e.g., sample “391_125” comes from site 391 at depth 125 cmbsf). The site labels are shaded in

colours that correspond to the colour code in Figure 5. Modified after (Algora et al 2015).

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 Bacterial diversity analysed as number of OTUs

The bacterial diversity in samples from the various sites and depths was measured as

number of OTUs, defined here as groups of sequences with an identity of ≥ 97%.

Highest OTU numbers were detected at 25 cmbsf, with values ranging around 550

OTUs for all sites (Figure 53). In general, OTU numbers decreased with increasing

depth in all sites. The depth profiles of OTUs indicated highest diversity in shelf sites

363, 365, and 371, and in shallow sediments down to 125 cmbsf of basin site 389.

Figure 53. Bacterial diversity, measured as OTU numbers, in depth profiles for each site of the Baffin

Bay. Colour code according to each site as given in Figure 5.

 Association of bacterial groups to geochemical parameters

The association of OTUs (≥ 97% sequence identity) to the geochemical parameters was

evaluated with Spearman’s rank correlation tests. An OTU correlated significantly to a

geochemical parameter when the p-value was <0.05. OTUs having significant

Spearman correlation with an environmental parameter were taxonomically classified

using mothur and the Greengenes database via the naïve Bayesian classification

method. The relative amount of OTUs affiliated to a taxonomical group and

significantly correlating with a specific geochemical parameter in relation to the total

number of OTUs from that taxonomical group was calculated to evaluate the proportion

of a taxonomical group correlating to a specific geochemical parameter (Figure 54). The

highest 25% identified correlations were defined as strong (either positive or negative).

OTUs that were strongly positive correlating with Mn(II) concentrations affiliated to the

families Alcaligenaceae, Burkholderiaceae and Comamonadaceae within the class

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Betaproteobacteria (Figure 54). Indeed, ≥ 90% of the OTUs belonging to these three

betaproteobacterial families indicated a strong positive correlation with the

concentration of Mn(II), and a negative correlation with TOC content. The opposite was

observed for the GIF-9 cluster within the class Dehalococcoidia and members of

candidate phylum “JS1”, both with ≥ 90% of the OTUs having strong positive

correlation with TOC and negative correlation with Mn(II) (Figure 54). Positive

correlations with TOC were also observed for nearly half of the OTUs of the order

Dehalococcoidales within the class Dehalococcoidia. Moreover, Dehalococcoidia

(order Dehalococcoidales and GIF-9) showed a negative correlation with sulphate

concentration. Similarly, ≥ 90% of the OTUs falling into the candidate phylum “OP8”

correlated negatively with sulphate and positively with TOC. Taxa that strongly positive

correlated with the concentration of Fe(II) belonged to the class Alphaproteobacteria.

Around 40% of OTUs falling into Alphaproteobacteria strongly correlated to Fe(II),

and another 40% correlated to sulphate (Figure 54).

Figure 54. The relative amount of OTUs within a selected taxonomical group which had a significant

(P<0.05) correlation with a geochemical parameter is shown. The correlation was individually calculated

for each OTU by Spearman rank correlation tests. The top 25% positive and/or negative correlations are

defined as “strong positive” (++) or “strong negative” (--). The remaining significant correlations are

indicated as positive (+) or negative (-). The order Dehalococcoidales and GIF-9 belong to the class

Dehalococcoidia. The families Alcaligenaceae, Burkholderiaceae and Commamonadaceae belong to the

class Betaproteobacteria. Modified after (Algora et al 2015).

Discussion

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4 DISCUSSION

Marine sediments comprise a vast area of our planet, and are a major habitat for huge

amounts of highly diverse and novel microorganisms (Parkes and Sass 2009). Members

belonging to the class Dehalococcoidia are among the most abundant and widespread

microorganisms in marine sediments (Durbin and Teske 2011, Parkes et al 2014).

However, very little is known apart from their presence in marine sediments deciphered

after 16S rRNA gene clone library studies. The ecological role of marine

Dehalococcoidia is completely unknown. This study is a contribution for gaining some

insights into the ecological role of marine Dehalococcoidia, and some understanding of

the reasons why marine Dehalococcoidia are so abundant and widespread in marine

sediments. For that, an in situ study in sediments of the Baffin Bay was carried out,

investigating the distribution of marine Dehalococcoidia in various depths and sites,

and the environmental parameters, i.e., depth, geographical location, geochemistry,

which may influence such a distribution and abundance. The geochemical parameter of

organic carbon was observed to strongly and positively correlate to the abundance of a

specific group within the Dehalococcoidia, named GIF-9. Other parameters, such as

sulphate-, and manganese(II)- concentration negatively correlated with GIF-9

abundance. Interestingly, the clade GIF-9 was present in shelf sediments (site 371), but

not in basin sediments (site 453) of the Baffin Bay (Wasmund et al 2015). High

differences among the presence and abundance of the members encompassing the class

Dehalococcoidia were observed depending on the depth and site of the Baffin Bay,

suggesting variability in the diversity of Dehalococcoidia based on sediment local

conditions. In addition, marine Dehalococcoidia were observed to persist with depth

and account for high percentages (up to 70%) of the total bacterial community at depths

where low abundance of total Bacteria was detected.

Additionally, an ex situ study that relied on the cultivation of marine sediments from

various locations and using an anoxic defined minimal medium supplied with various

potential electron acceptors was performed to investigate possible respiration modes for

Dehalococcoidia. Sulphate, iron(III), manganese(IV), and various halogenated

compounds were selected as potential electron acceptors that were individually supplied

to sediment cultures. Halogenated compounds were chosen because the

phylogenetically closest bacteria to marine Dehalococcoidia are strict organohalide-

respiring bacteria, such as Dehalococcoides mccartyi (Löffler et al 2013, May et al

2008, Moe et al 2009). Although dehalogenation of 1,2,3-TCB was observed in this

study, it could not be associated to Dehalococcoidia growth, suggesting that marine

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Dehalococcoidia did not respire the halogenated compounds used here. However, no

other electron acceptors tested here promoted a substantial growth of Dehalococcoidia,

suggesting alternative respiratory modes or fermentation as a mode of living for marine

Dehalococcoidia.

Altogether, the present study suggests marine Dehalococcoidia to be highly diverse

displaying different respiratory modes. Some of the marine Dehalococcoidia are most

likely heterotrophic bacteria, e.g., clade GIF-9 (Hug et al 2013a, Wasmund et al 2014).

Dehalococcoidia are widespread in marine sediments most likely due to their resilience,

i.e., their ability to survive burial, high pressures, long periods of starvation and/or low

concentration of nutrients, which is an advantage in an environment depleted in

nutrients such as deep marine sediments, and/or may use recalcitrant organic substrates

that most other bacteria cannot metabolize.

4.1 IN SITU ABUNDANCE OF DEHALOCOCCOIDIA

The determination of the abundance of the class Dehalococcoidia (phylum Chloroflexi)

in different marine sediment samples aimed to identify natural conditions, e.g., sediment

geochemistry, depth, and geographical location that support Dehalococcoidia natural

occurrence in an attempt to better understand the ecological role of Dehalococcoidia in

marine sediments. For this, an in situ approach determining the abundance of

Dehalococcoidia 16S rRNA genes in sediments of the Baffin Bay was performed. The

abundance was measured here as “relative abundance” of Dehalococcoidia 16S rRNA

genes within the total bacterial 16S rRNA genes, and as “absolute abundance”, which

refers to the number of 16S rRNA gene copies amplified and quantified with a qPCR

assay targeting specifically Dehalococcoidia. The relative abundance was determined

from the amplicon sequencing of 16S rRNA genes performed with Illumina sequencing.

In Baffin Bay samples, members of the class Dehalococcoidia were present at all

sediment sites and depths, suggesting a widespread distribution, which agrees with the

ubiquitous presence of Dehalococcoidia found in marine sediments worldwide in

previous studies (Fry et al 2008, Parkes et al 2014).

Relative abundance of bacterial classes in Baffin Bay sediments indicated

Dehalococcoidia as the second most abundant class, which accounted for 6 to 17% of

the total bacterial community, after bacteria of the class Betaproteobacteria (between 38

to 64%). On the phylum level, the relative abundance for Chloroflexi was of 8 to 22%,

which was slightly less than the average relative abundance of 25.5% for Chloroflexi, as

Discussion

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determined in a review of bacterial communities in various subsurface marine sediments

(Parkes et al 2014).

Absolute abundance of Dehalococcoidia was around 105 16S rRNA gene copy numbers

per gram of sediment for most Baffin Bay sediments. Shelf areas showed, however, one

order of magnitude higher copy numbers than other areas (Figure 45, Figure 46, and

Figure 47, and (Wasmund et al 2015)). Total bacterial numbers were maintained around

107 16S rRNA gene copies g-1 along the studied sediment profile in shelf sites of the

Baffin Bay. However, basin sites of the Baffin Bay showed significant decreasing

numbers of total Bacteria from nearly 108 to 104 16S rRNA gene copies g-1 with

increasing depths (this study Figure 43, and (Algora et al 2013, Wasmund et al 2015)).

At similar depth ranges (from seafloor down to 6 mbsf), sediments from other locations

showed similar copy numbers of Dehalococcoidia and an order of magnitude difference

in copy numbers depending on basin/shelf sample origin. For instance, the number of

Dehalococcoidia was of 105 16S rRNA gene copies g-1 in basin sediments from the

forearc basin in Sumatra, meanwhile shelf sediments from Århus Bay showed one order

of magnitude higher copy numbers of 106–107 g-1 (Wasmund et al 2015). The

percentage of Dehalococcoidia 16S rRNA gene copy numbers in relation to the total

bacterial 16S rRNA gene copy numbers showed Dehalococcoidia to account for ≤ 10%

of the bacteria in shelf sediments of the Baffin Bay (Figure 48 and (Wasmund et al

2015)). Interestingly, Dehalococcoidia appears to be a resilient bacterial group that

persists with burial and make up to 74% (at a depth of 222 cmbsf at site 391), and more

than 10% when low bacterial copy numbers are found at deeper sediment layers, i.e.,

depths of 374, and 290–320 cmbsf from the shelf sites 371, and 365, respectively; and

depths of 150–400 cmbsf at basin sites 389, 391 and 224–277 cmbsf at site 453 (Figure

48 and (Wasmund et al 2015)). Thus, Dehalococcoidia may have metabolisms requiring

low energy for cell maintenance and/or use recalcitrant substrates, which are not

degraded in shallow sediments.

Both absolute and relative abundance for Dehalococcoidia were highest at the shelf

compared to any other area. Geochemically, the shelf has higher contents of organic

carbon than the slope or basin, and highest indications of sulphate reduction, i.e.,

highest abundance of dsrA genes (functional markers for sulphate-reducing

microorganisms), and decreasing sulphate concentrations from ~25 to ~20 mM at

depths below 4 mbsf. Iron and manganese reduction also occur in the shelf sediments,

although at depths of ≥200 cmbsf. In sediments of the Baffin Bay, values for the content

of organic carbon strongly correlated with the relative abundance of Dehalococcoidia,

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115

and especially, the clade GIF-9 (formerly known as NT-B4, (Reed et al 2002)).

Accordingly, a literature review has described an average relative abundance of 41.3%

of bacterial 16S rRNA gene sequences belong to the phylum Chloroflexi in various

shelf sediment sites with high organic carbon contents (Parkes et al 2014). Some

examples of sediments containing high organic carbon contents are Mediterranean

sapropels and the Peru margin. In sediments from Mediterranean sapropels, almost the

whole bacterial community belonged to Dehalococcoidia (Coolen et al 2002). In the

Peru Margin, an average of 41% of the Chloroflexi correlated to 16S rRNA gene

libraries from subsurface organic rich shelf sediments (Parkes et al 2014). A recent

study specifically found the occurrence of the clade GIF-9 correlated with values of

organic carbon content through different depths of a single core from mud flats of

Helgoland, North Sea (Oni et al 2015). Previous studies have also suggested that the

GIF-9 clade is associated to organic-rich, as well as methane-bearing marine sediments

(Harrison et al 2009, Takeuchi et al 2009, Teske et al 2011). Together, previous studies

and this study from Baffin Bay sediments indicated that members of the GIF-9 are

associated with organic rich sedimentary environments. The GIF-9 clade of the

Dehalococcoidia may therefore have an ecological role in the carbon cycle for the

degradation of organic matter; however their exact role, e.g., whether they are involved

in a primary or secondary stage of the degradation of organic matter, or their

metabolisms, remain currently unknown.

Further indications about the ecological role and metabolism of the GIF-9 clade can be

inferred from the specific profiling of Dehalococcoidia 16S rRNA genes in sediments

of Baffin Bay sites 371 and 453 (Wasmund et al 2015). In general, Dehalococcoidia

were more diverse at shelf site 371 than at basin site 453. Site 371 contains higher

organic carbon content than site 453. Interestingly, the clade GIF-9 accounted for a high

percentage of the Dehalococcoidia present in sediments of site 371, and especially at a

depth of 173 cmbsf. However, no Dehalococcoidia belonging to the GIF-9 clade was

present at site 453. Clade GIF-9 therefore appears to be a Dehalococcoidia clade

inhabiting shelf sediments rich in organic matter, as they were also found in high

percentages in Århus Bay sediments and at the Peru margin site 1227, which at depths

of 60 mbsf accounted for the 95% of the total Dehalococcoidia present in the sample

(Inagaki et al 2006, Wasmund et al 2015). Interestingly, GIF-9 was absent at site 453 of

the Baffin Bay basin and also at sites from the Arctic Mid-Ocean Ridge cores GC6 and

GC12 (Wasmund et al 2015), where low organic matter content were found (Jørgensen

et al 2012). Therefore, microbial ecology studies in the Baffin Bay and other areas give

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116

indications that GIF-9 may be heterotrophic or fermenting bacteria thriving in organic-

matter rich sediments from coastal margins containing complex organic matter.

Recently, genomic content from members of GIF-9 clade were obtained. These include

a single cell named DEH-J10 retrieved from Århus Bay sediments (Wasmund et al

2014), a draft genome curated from metagenomic data named RBG-2 from sediments of

an aquifer next to the Colorado River (Hug et al 2013a), and a partial (~35%) single cell

named Dsc1 retrieved from site 1230 of the Peru Margin (Kaster et al 2014). Insights

into the metabolism of DEH-J10 indicated genes encoding for enzymes involved in

β-oxidation of organics, aromatic compounds, and other organics completely to CO2 via

the Wood-Ljungdahl pathway (this pathway is reversible and can function for

autotrophic CO2 fixation), in addition to fermentation (Wasmund et al 2014). Thus, a

heterotrophic, fermentative metabolism is most likely for DEH-J10, which may explain

the strong association of GIF-9 clade to organic matter in sediments of the Baffin Bay.

Furthermore, RBG-2 annotations indicated homoacetogenic metabolisms, producing

acetate as final product of the fermentation of glucose or plant polymers, i.e., pyrogallol.

Therefore, RBG-2 may have a saprophytic role in aquifer sediments. Additionally, the

RBG-2 genome encodes for the degradation of fatty acids via β-oxidation and also the

Wood-Ljungdahl pathway (Hug et al 2013a). Thus, both DEH-J10 and RBG-2 genomic

insights agree with the hypotheses put forward based on ecological observations,

providing further indications that the GIF-9 clade is most likely directly involved in the

remineralisation of organic matter and performs fermentation. These features may be

the reason why GIF-9 is commonly found in organic-rich areas.

Furthermore, more than 90% of the OTUs affiliated to GIF-9 strongly negatively

correlated to sulphate concentration (Figure 54 and (Algora et al 2015)). In sediment

depth profiles from a site in Århus Bay and site 1227 of the Peru margin, clade GIF-9

was detected in sediment depths with and without sulphate (Inagaki et al 2006,

Wasmund et al 2015). In Århus Bay, GIF-9 accounted for the highest relative

proportion of total Dehalococcoidia in sediments with low sulphate concentration. In

the Peru margin, highest relative proportion of GIF-9 from the total Dehalococcoidia

was found underneath the sulphate-methane transition zone. The presence of GIF-9 in

sediments with low or no sulphate in Århus Bay and Peru margin, together with the

strong negative correlation shown in Baffin Bay sediments, indicate that GIF-9

members most likely are not sulphate-reducing bacteria, or that they are linked to high-

affinity sulfate reduction, e.g., through syntrophic associations with microbes

specialised for the reduction of low concentrations of sulphate (Tarpgaard et al 2011).

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Further, no genes encoding for sulphate respiration and for a respiratory metabolism in

general, were found in both DEH-J10 and RBG-2.

Further research to get more insights into the metabolisms and ecological roles of

bacteria of the class Dehalococcoidia, and especially the clade GIF-9, may include

cultivation with different organic substances to document possible growth, as well as

cultivation-independent studies such as stable isotope probing and activity

measurements in sediments. In this line, it will be very helpful to investigate the organic

matter composition associated to GIF-9 members to suggest potential organic substrates

for GIF-9 bacteria cultivation and also as correlation studies in the field.

4.2 ELECTRON ACCEPTORS IN THE BAFFIN BAY

The Baffin Bay is located in the Arctic, between Canada and Greenland. It is a remote

area, covered by ice during the great majority of the year except for the summer months,

thus, affecting primary production, sedimentation rates, and organic content in its

sediments. Few studies have been carried out so far in sediments of the Baffin Bay.

Here, I studied the geochemistry and the indigenous microorganisms inhabiting the

Baffin Bay sediments at various sites and depths for gaining understanding of the

element cycling in its sediments (Algora et al 2013, Tang et al 2004). Various locations

were investigated including shelf, slope, and basin sites. In general, shelf sediments had

highest content of organic matter and sulphate decreases in concentration, indicating

that sulphate reduction was an important mineralization pathway for organic matter.

Additionally, highest bacterial and archaeal 16S rRNA gene copy numbers, and highest

bacterial diversity over nearly entire core lengths was observed in shelf sediments.

Highest dsrA gene copy numbers in shelf sediments revealed that most likely sulphate

was the main terminal electron acceptor for microorganisms. On the contrary, sediments

from the basin and slope had comparably much lower content of organic matter, which

was mainly mineralized by iron(III)- and manganese(IV)-reduction. Depth affected

more dramatically bacterial and archaeal abundance and diversity in basin sites, with

highest 16S rRNA gene copy numbers and OTUs in uppermost sediments, which

decreased substantially with increasing depths. 16S rRNA gene copy numbers from

members of the order Desulfuromonadales, which encompasses known metal reducers

such as the genera Geobacter and Desulfuromonas, revealed no association with iron or

manganese concentrations. Thus, other metal reducers may be present in Baffin Bay

sediments which may mediate the reduction of iron(III) and manganese(IV) oxides.

Altogether, the analysis of the geochemistry and the microbial communities revealed

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strong differences based on the local sedimentary conditions. These differences were

especially evident between shelf and basin sites.

4.2.1 Sulphate reduction and methanogenesis in shelf sediments

Due to the high concentration of sulphate in seawater, sulphate is the quantitatively

major electron acceptor for the microbial mineralization of organic matter in many

marine sedimentary environments (D'Hondt et al 2002, Jørgensen 1982). In the samples

measured here from sediments of the Baffin Bay, sulphate was available down to a

depth of 5 mbsf. At shelf and slope sites, as well as at basin site 453, sulphate

concentrations decreased with increasing depths, indicating that sulphate reduction in

sediments of the Baffin Bay occurs. Sulphate reduction was also measured in slurries

prepared with and without various substrates including hydrocarbons, monomeric and

polymeric carbohydrates (i.e., chitin, cellulose, peptone) from Baffin Bay sediments

(Algora et al 2013). Potential rates of sulphate reduction in the slurries indicated the

shelf as the most active area in sulphate reduction as a result of highest organic carbon

content compared to other sampled areas, which agrees with previous reports from

Arctic sediments. In those reports, in situ sulphate reduction was limited by organic

carbon content independently of the temperature (Arnosti and Jørgensen 2006,

Vandieken et al 2006). Methane concentrations in Baffin Bay sediments were very low

indicating that the sulphate-methane transition zone was below the studied sediment

layers. Previous pore-water measurements of the ODP site 645 showed depleted

concentrations of sulphate at 35 mbsf (Srivastava et al 1989). A plausible explanation

for such a deep sulphate-methane transition zone compared to other locations in the

world may be the low organic content. Low TOC values were especially found at basin

sites where long water columns and distance to land result in decreased sedimentation

of organic matter. Distance to land is critical for organic matter content in Baffin Bay

sediments due to low primary productivity in the overlying water column, which is the

result of seasonal ice cover. Terrestrial origin of the organic matter is indicated by the

isotope value of the organic carbon (Algora et al 2013, Sackett 1964, Srivastava et al

1989). Thus, shelf areas have increased organic carbon coming from Greenland.

Sulphate reducing bacteria use sulphate as terminal electron acceptor in their respiratory

chains. The enzyme dissimilatory sulphite reductase (Dsr), encoded by the genes dsrA

and B, catalyses the final reaction in sulphite reduction, which is the reduction of

sulphite to sulphide, and is present in all known sulphite/sulphate reducers using the

canonical sulphite/sulphate reduction pathway (Wagner et al 1998). Therefore, dsrA

genes are used as a proxy for presence and quantification of sulphate reducing

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119

microorganisms in marine sediments (Kondo et al 2004). dsrA genes were abundant in

Baffin Bay sediments, indicating the potential for sulphate reduction at all depths and

sites. The abundance of dsrA genes in Baffin Bay sediments was higher than at other

locations such as the Peru margin ODP site 1227 or the Wadden Sea in Germany

(Blazejak and Schippers 2011, Schippers and Neretin 2006, Wilms et al 2007). Within

Baffin Bay sites, dsrA gene abundance was highest in shelf sediments and Southern

slope site 486, indicating increased numbers of sulphate reducing microorganisms at

sites with higher organic carbon content.

The gene encoding for the alpha subunit from the methyl coenzyme M reductase (mcrA)

is present in methanogenic and anaerobic methanotrophic archaea (Friedrich 2005,

Hallam et al 2003, Nunoura et al 2008). mcrA gene abundance was used here for the

detection and quantification of methanogens and methanotrophs. Both low methane

concentrations and low abundance of mcrA indicated a minor role of methanogenesis

and/or methanotrophy in sediments of the Baffin Bay. Only in the shelf area, increased

mcrA gene copy numbers evidenced the presence of methanogens, although at low

abundance. Interestingly, slurries prepared with shelf sediments and the aliphatic

hydrocarbons hexadecane and hexadecanoic acid showed methane formation (Algora et

al 2013). However, methane was not formed in shelf sediment slurries amended with

aromatic hydrocarbons (Algora et al 2013). As a conclusion, methanogenesis and

methane oxidation do not play a pronounce role in the Baffin Bay in the sediment

depths analysed. Most likely, the low organic carbon content in Baffin Bay sediments is

mineralized using electron acceptors other than CO2, i.e., sulphate, iron, or manganese,

which yield higher energy. Therefore, organic matter quantity and availability are

driving the element cycles in marine sediments of the Baffin Bay as in many other

marine sediments worldwide inhabited by heterotrophic microorganisms.

4.2.2 Importance of metal biogeochemistry in the central Baffin Bay

In the Baffin Bay, accumulation of Mn(II) and Fe(II) in pore-water profiles indicated

the reduction of manganese and iron oxides (i.e., MnO2 and Fe2O3, respectively) from

depths of 0.25 mbsf down to 4.7 mbsf. Thermodynamically, Mn(IV) provides highest

energy yields followed by Fe(III) and then by sulphate, in the anoxic mineralization of

organic matter mediated by microorganisms (Froelich et al 1979). Mn(IV) and Fe(III)

are reduced to Mn(II) and Fe(II) in this process. The reduced forms were observed in

high concentrations in pore-water profiles from the central basin sites indicating that

metal element cycling is important in basin sediments of the Baffin Bay. Basin

sediments were characterized by low organic carbon contents, which may be mainly

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mineralized via Mn(IV) and Fe(III) reduction as organic carbon content decreases at

those depths where Fe(II) and Mn(II) concentration in pore-water increased. Generally,

low concentrations of manganese oxides are found in marine sediments (≤ 20 µmol cm-

3) and at shallow depths of ≤ 2 cmbsf (Thamdrup 2000). Thus, manganese is rapidly

depleted in surface sediments, and therefore collectively plays a minimal role for the

mineralization of the bulk of organic matter, especially in marine sediments rich in

organic matter (Nealson and Saffarini 1994, Sørensen and Jørgensen 1987, Vandieken

et al 2006). However, some sediment sites have high manganese oxide concentrations

of 25 to 185 µmol cm-3, such as sediments from the Panama Basin, the Black Sea, the

Barents Sea and some parts of the Skagerrak (Canfield et al 1993, Nickel et al 2008,

Thamdrup 2000, Vandieken et al 2006), where manganese oxides are found reaching

depths down to 410 cmbsf. Manganese reduction contributes from 25% to nearly 100%

to the anaerobic carbon oxidation at those sites. In particular, in Arctic sediments from

the Barents Sea, Mn(IV) and Fe(III) are reported to contribute between 69% to more

than 90% to the mineralization of organic matter within near-surface sediments of

10 cm (Vandieken et al 2006). Mn(IV) and Fe(III) reduction processes are therefore of

importance as the main respiratory pathways for the carbon mineralization in Arctic

sediments (Vandieken et al 2006). A reason for metal cycling predominance in polar

areas may be that polar ice sheets are enriched in iron, manganese and organic matter

(Lannuzel et al 2014). During the ice melt, iron, manganese and organic matter fuel

phytoplankton and microbial growth in the seawaters (Lannuzel et al 2013, Lannuzel et

al 2014, Martin and Fitzwater 1988). Some of the iron and manganese is deposited on

the seafloor, usually associated with organic particles and/or sediment grains (Lannuzel

et al 2014, Raiswell et al 2006). Apart from sea-ice, glacial melt-water and glacial

sediments transported at the base of icebergs additionally contribute to the release of

bioavailable iron into the ocean waters (Bhatia et al 2013, Raiswell et al 2006). The

Baffin Bay is ice-covered most of the year except the summer months (Tang et al 2004).

Therefore, Baffin Bay sediments may be supplied with manganese, iron, and organic

matter from the melting of the sea-ice and the surrounding glaciers.

Well-known bacterial groups associated to iron and manganese reduction are members

of the order Desulfuromonadales, class Deltaproteobacteria (Childers et al 2002,

Lovley et al 2004). In sediments of the Baffin Bay, members of the

Desulfuromonadales were detected. Desulfuromonadales 16S rRNA gene abundance

followed a similar pattern to total bacterial abundance with depth, independently of

Fe(II) and Mn(II) concentrations in the pore-water. Furthermore, no positive correlation

was found between the relative abundance of the class Deltaproteobacteria and Fe(II)

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121

or Mn(II) pore-water concentrations (Figure 54). On the contrary, 40% of the OTUs

affiliated to Deltaproteobacteria showed a negative correlation to Mn(II) pore-water

concentration. This suggests that Deltaproteobacteria are not predominantly involved in

the reduction of Fe(III) and Mn(IV) in Baffin Bay sediment pore-water. In the microbial

community analysis of Baffin Bay sediments, the class Betaproteobacteria was

dominant, especially in basin sediments. Pore-water concentration of Mn(II) negatively

correlated with microbial diversity (Algora et al 2015), indicating the enrichment for a

few specialized microbial groups, i.e., members of the class Betaproteobacteria, which

may be the bacteria performing the largest share of the Mn(IV) reduction, and therefore

oxidation of organic matter in the Baffin Bay.

4.3 BURKHOLDERIALES DOMINATE IN BAFFIN BAY SEDIMENTS

The class Betaproteobacteria, with a relative abundance of 38 to 64%, as determined

from 16S rRNA gene amplicon sequencing, was dominant in all analysed sediments

from the Baffin Bay. The average presence of Betaproteobacteria in marine sediments

is of 5% according to various studies from subseafloor sediments bordering the Pacific

Ocean, reviewed by (Fry et al 2008), and of 2% according to (Parkes et al 2014), which

reviewed bacterial communities from various subseafloor sediment studies worldwide

combining 205 prokaryotic 16S rRNA gene libraries. Thus, sediments from the Baffin

Bay are characterized by a considerably high abundance of Betaproteobacteria in their

bacterial community. Other marine sediments with high abundance of

Betaproteobacteria that substantially exceed the 2–5% average were found in the

Cascadia Margin site 889/890 (relative abundance of betaproteobacterial clones of 20%

at 9 mbsf, 55% at 198 mbsf, 47% at 222 mbsf, and 5% at 234 mbsf (Marchesi et al

2001)). Betaproteobacterial clones from the Cascadia Margin site 889/890 were

phylogenetically most similar to Ralstonia pickettii, which belongs to the order

Burkholderiales. R. pickettii is a heterotrophic bacterium inhabiting oligotrophic

habitats (Ryan et al 2007) that include metal-enriched environments such as in acid

mine drainage (Kimura et al 2011), and in deep-sea basalts and sediments from the Mid-

Atlantic Ridge (Rathsack et al 2009).

Within the class Betaproteobacteria, 99.7% on average of the clones found in sediments

of the Baffin Bay belonged to the order Burkholderiales. The order Burkholderiales

encompasses a metabolically diverse group of bacteria that includes aerobic, facultative

anaerobic, diazotrophic, chemoorganotrophic, and chemolitotrophic microorganisms

(Garrity et al 2005). Interestingly, Burkholderiales genomes encode many oxygenases

able to degrade a wide range of aromatic compounds (Pérez-Pantoja et al 2012).

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In the Baffin Bay, ≥ 90% of OTUs affiliated to Burkholderiales positively correlated to

Mn(II) concentrations and negatively to organic matter content. In basin and slope sites,

as well as deep sediment layers from shelf site 365, higher relative abundance of

Burkholderiales in accordance to lower organic matter content and higher Mn(II)

concentration were detected. These results, together with the negative correlation of

pore-water Mn(II) concentration with the microbial diversity (measured as number of

OTUs) already mentioned in the previous chapter, suggest that Burkholderiales may be

involved in manganese cycling. To my knowledge, no studies in the marine subsurface

link Burkholderiales abundance to the concentration of manganese. However, in the

terrestrial subsurface, an association of the order Burkholderiales with metal cycling

was observed after the isolation of Rhodoferax ferrireducens from aquifer subsurface

sediments of Oyster Bay, Virginia, USA (Finneran et al 2003). R. ferrireducens is a

psychrotolerant bacterium able to use Fe(III)- and Mn(IV)- oxides, nitrate, fumarate and

oxygen as electron acceptors, with acetate or lactate as electron donors (Finneran et al

2003). Indeed, many iron reducers are also able to reduce manganese oxides (Lovley et

al 2004, Nealson and Saffarini 1994). A couple more studies in the terrestrial subsurface

found Burkholderiales associated to Fe(III) reduction in either contaminated

groundwater or sediment, both after stimulation with acetate (Handley et al 2014,

Livermore et al 2013). Further terrestrial environments include ferromanganese nodules

either in natural freshwater sediments (Stein et al 2001) or in anthropogenic sugarcane

and rice paddy fields (Hu et al 2015). Identified Burkholderiales in the ferromanganese

nodules belonged to the genus Leptothrix, which are known to oxidize Mn(II) and Fe(II)

coupled to oxygen or nitrate reduction (Carlson et al 2013, De Vrind-De Jong et al

1990, Stein et al 2001). However, in the Baffin Bay subsurface, the Burkholderiales are

more probable associated to the Mn(IV) and Fe(III) reduction than Mn(II) and Fe(II)

oxidation, as deduced from the accumulation of the soluble metal forms Mn(II) and

Fe(II) in the pore-waters. In addition, Fe(II) accumulation at the bottom of cores 365

and 371 are associated with higher relative abundance of Burkholderiales. Interestingly,

the bacterial community in deep sediment layers from sites 365 and 371, where

increased Fe(II) concentrations were observed, cluster together with samples from the

basin sites where the metal cycling of both manganese and iron prevails. Thus, I

hypothesize that Burkholderiales are possibly reducing both Mn(IV) and Fe(III) in the

Baffin Bay. Future studies on cultivation of Baffin Bay sediments in microcosms

amended with Fe(III) and Mn(IV) may lead to clear relationships of betaproteobacterial

growth associated to reduction of metal oxides.

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Burkholderiales are prevalent in glaciers or glacially-associated environments.

Examples for this are an ice-core from Greenland frozen for over one-thousand-years

(Sheridan et al 2003), permanent lake ice from Antarctica (Gordon et al 2000), moraine

chronosequences in the primary succession of a recently deglaciated and unvegetated

soil from a Peruvian receding glacier (Nemergut et al 2007), Arctic moraines (Mapelli

et al 2011), and subglacial sediments (Carr et al 2013, Foght et al 2004, Lanoil et al

2009, Skidmore et al 2005, Stibal et al 2012, Yde et al 2010). The Baffin Bay is ice-

covered for the whole year except the summer and receives sediments and water from

glacial melt (Bhatia et al 2006, Stibal et al 2012, Wadham et al 2008). I therefore

hypothesize that the sedimentary Burkholderiales detected in the Baffin Bay could

come from the sediments underneath glaciers in Greenland and Canada. The

Burkholderiales may be transported by icebergs and released to the Baffin Bay waters

attached to sediment particles or iron aggregates after the ice-melt. Thus,

Burkholderiales may colonize sediments from the Baffin Bay, similarly to the way that

iron and manganese particles or organic matter are delivered to the oceans and later on

to the sediments (Lannuzel et al 2014, Raiswell et al 2006). Burkholderiales can be

aerobic and facultative anaerobic bacteria using nitrate or iron for respiration (Willems

et al 1991). For example, facultative anaerobic bacteria from the genus Rhodoferax

(family Comamonadacea; order Burkholderiales) may reduce iron (Stibal et al 2012,

Willems et al 1991). Rhodoferax–related bacteria have been commonly detected in basal

ice from glaciers (Lanoil et al 2009, Skidmore et al 2005, Yde et al 2010). Rhodoferax

can deal with oxic conditions, which is an advantage compared to other iron reducers

such as members of the Geobacteraceae family (order Desulfuromonadales; phylum

Deltaproteobacteria) in periods with oxic-anoxic conditions such as glacial sediments

or sea-ice where melting occurs annually (Yde et al 2010), and may be the reason of

Burkholderiales predominance in sediments of the Baffin Bay over

Desulfuromonadales. Therefore, Burkholderiales may change from aerobic to nitrate to

Mn(IV) and Fe(III) respiration in sediments for adaptation in glacial sediments in

Greenland, as already suggested (Stibal et al 2012), and may be the case for marine

sediments of the Baffin Bay, which may be the key to their dominance in Baffin Bay

sediments.

4.4 DEHALOCOCCOIDIA CULTIVATION

The close phylogenetical affiliation of Dehalococcoides mccartyi, Dehalogenimonas

lykanthroporepellens, Dehalogenimonas alkenigignens, and Dehalobium chlorocoercia

strain DF-1, which are all strict organohalide-respiring bacteria (Löffler et al 2013, May

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et al 2008, Moe et al 2009), to marine Dehalococcoidia members may suggest an

organohalide-respiring metabolism of marine Dehalococcoidia (Adrian 2009).

However, the presence of organohalide compounds in the sediment cultures did not

enhance growth of marine Dehalococcoidia, suggesting that either they use other

organohalides not tested here or they do not use organohalides. Interestingly, I observed

the conversion of 1,2,3-TCB to 1,3-DCB in one sediment culture, which was further

transferred. In the subculture, the transformation of 1,2,3-TCB to 1,3-DCB was

observed together with marine Dehalococcoidia growth of one order of magnitude

within two months of incubation time. However, organohalide respiration could not be

linked to marine Dehalococcoidia growth as further transfers failed to transform 1,2,3-

TCB, maybe due to the transfer of low biomass within the inoculum, or exposure to

inhibitory concentrations of 1,2,3-TCB. Cloned sequences of the Dehalococcoidia

present in that subculture revealed an affiliation to Dehalococcoidia sister-clade DSC-

D, falling far from the Ord-DEH cluster of the known dehalogenating Dehalococcoidia

(name of clades as indicated by (Wasmund et al 2015)). Further sediment cultivation

amended with 1,2,3-TCB revealed members of the phylum Firmicutes as the bacteria

transforming 1,2,3-TCB to 1,3-DCB. Regarding the wide diversity and high abundance

of the Dehalococcoidia in marine sediments, alternative metabolisms other than

organohalide respiration are plausible. In fact, recent genomic data from four marine

Dehalococcoidia did not contain genes related to organohalide respiration, i.e., RDases

(Hug et al 2013a, Kaster et al 2014, Wasmund et al 2014).

In this study, no electron acceptor among those tested was observed to preferably and

substantially support growth of Dehalococcoidia in the various sediments. All tested

electron acceptors resulted in similar Dehalococcoidia 16S rRNA gene copy numbers,

indicating a lack of a respiratory metabolism with these electron acceptors, or that the

preferred electron donors/vitamins/minerals were not supplied, or that growth rates were

not detectable within the available incubation time. Thus, marine Dehalococcoidia may

be fermenters, as already suggested by the in situ approach and genomic studies (Hug et

al 2013a, Wasmund et al 2014). An alternative explanation may be that the

Dehalococcoidia have diverse metabolisms, including both respiration and

fermentation. Metabolic diversity within the class Dehalococcoidia may explain the

patchy Dehalococcoidia abundance observed for some sediments amended with

electron acceptors and monitored at different incubation times. For example, cultures

prepared with sediments originating from Århus and a site in Ireland gave some

indications of sulphate as potential electron acceptor favouring the growth of marine

Dehalococcoidia. In addition, a single cell retrieved from Århus sediments named

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DEH-C11, which belongs to the class Dehalococcoidia (specifically to the

Dehalococcoidia sister-clade DSC-GIF3-B), contains a dsr gene operon as well as

genes for arsenate reduction, suggesting that Dehalococcoidia may, in fact, respire

certain compounds (Dr. Kenneth Wasmund, personal communication). On the other

hand, Dehalococcoidia abundance in sediment cultures from Chile varied randomly,

and clearly showed that none of the tested electron acceptors favoured growth.

Although variability in the abundance of Dehalococcoidia could be due to

heterogeneities or/and errors in the sampling and analysing method (sediment sub-

sample extraction together with DNA isolation and qPCR assay), a test trial indicated

this method as reliable for total bacterial identification. Altogether, Dehalococcoidia

variable abundance indicates diverse metabolisms, as already suggested by in situ

studies (Wasmund et al 2015) and as previously mentioned. In this case, an approach

whereby each separate clade is targeted by quantitative methods such as qPCR may be

most appropriate than per whole class, since the quantification of some clades and

therefore their growth may be masked by changes in quantities of others. For instance,

an approach to investigate if the clade GIF-9 encompasses fermenting microorganisms,

or if the clade DSC-GIF3 includes potential sulphite/sulphate reducers, as recently

suggested by DEH-C11 single-cell study (Dr. Kenneth Wasmund, personal

communication), might be more suitable than investigating the entire class

Dehalococcoidia. Moreover, addition of a sediment inoculum less than 9–10% used

here may be beneficial, as the inoculated sediment supplies with nutrients and electron

acceptors, e.g. sulphate, which may additionally support microbial growth

independently of the compounds added by the medium.

Although no clear respiratory mode could be deciphered, Dehalococcoidia were

detected in the sediment cultures and 16S rRNA gene copy numbers were maintained

with time indicating that it is possible to cultivate them using a minimal mineral

medium under atmospheric pressures and at temperatures of 30ºC. Therefore, the

marine Dehalococcoidia are not obligate piezophilic or psychrophilic bacteria. For

instance, Dehalococcoidia numbers increased one order of magnitude after an

incubation time of two months in a subculture amended with 1,2,3-TCB. Thus, growth

rates for marine Dehalococcoidia members are not of hundreds of years as may be

thought for subseafloor microorganisms. Previous authors suggested that subsurface

bacteria may be damaged when exposed to high substrate concentrations due to

uncoupling of reactions (Parkes et al 2014). However, it seems not to be the case either

for the used medium here, or for the marine Dehalococcoidia bacteria.

Discussion

126

Nevertheless, a transferable enrichment of marine Dehalococcoidia was not achieved

and a plausible reason may be that Dehalococcoidia were out-competed by other faster

growing organisms such as those from the phylum Firmicutes, which were observed to

be present in some of the sediment cultures by clone libraries and 454-pyrosequencing,

in particular, in enrichments from Chile site 7155 (discussed below). In addition, all

investigated colonies that were formed in deep-agarose dilution tubes from Chile site

7155 (the rest of sites and sediments showed very few to no colonies) belonged to

members of the Firmicutes. Possible ways to eliminate fast growing Gram-positives are

using antibiotics specific for Gram-positive cell types, i.e., ampicillin, vancomycin,

penicillin, in order to avoid out-competition. Additionally, using a medium with very

low nutrient concentrations to avoid spore germination or even cultivating at low

temperatures may be also favourable for cultivation of indigenous microorganisms in

marine sediments.

Isolation of marine Dehalococcoidia in deep-agarose dilution tubes was not possible. A

possible reason is that the Dehalococcoidia may form too small or non-visible colonies

as indeed other cultivated Dehalococcoidia such as Dehalococcoides mccartyi strain

CBDB1 colonies are hardly visible. In addition, some microorganisms may not be

isolated as pure strain, but may live in symbiosis or syntrophy with other strains or may

need other microorganisms for, e.g., quorum sensing, so isolation may also be difficult

or even impossible if the metabolite exchange between microbial species is not known.

Altogether, marine Dehalococcoidia are able to be maintained in culture under varying

conditions, e.g., Mn(IV)-, Fe(III)-, sulphate-, and bicarbonate-reducing conditions.

Nevertheless, these organisms are still hard to isolate. To my knowledge, no previous

study has enriched or maintained in sediment cultures members of the marine

Dehalococcoidia, so it is not possible to compare with such other reports for obtaining

further overall conclusions. From my perspective, it may be reasonable to hypothesize

that enrichments of Dehalococcoidia, which may lead to cell densities high enough to

give improved chances of isolation, were not possible due to lack of suitable substrates,

e.g., electron donor/acceptor combinations. Better knowledge on the chemical nature of

the complex organic matter present in marine sediments may substantially aid for this

endeavour of suppling suitable substrates for cultivation of marine Dehalococcoidia.

4.5 DEHALOGENATION OF ORGANOHALIDES IN SEDIMENT

CULTURES

In sediment cultures from Chile, 1,2,3-TCB was transformed to 1,3-TCB. Previous

studies in the marine subsurface also observed the transformation of organohalides, i.e.,

Discussion

127

2,4,6-tribromophenol, 2,4,6-triiodophenol, and TCE in deep marine sediments from the

Nankai Trough, Japan (Futagami et al 2009, Futagami et al 2013). Thus, organohalide

respiration may occur in deep marine sediments. Organohalides are naturally produced

in the marine environment (Gribble 2003) and buried to marine sediments. Thus,

organohalides may be potential electron acceptors in the marine subsurface.

Interestingly, the marine subsurface is most likely one of the very few environments on

Earth that may be truly uncontaminated, i.e., deep sediments deposited prior to the

industrial revolution are completely pristine. Therefore, the marine subsurface is a well-

suited environment to study the natural cycle of halogens. Reductive dehalogenase

homologous gene (rdhA) sequences have been amplified from various subsurface

marine sediments (Futagami et al 2009, Futagami et al 2013), however not always

associated to dehalogenation activities (Futagami et al 2013). The organohalide 1,2,3-

TCB was never observed to be transformed by autochthonous microbiota in deep

marine sediments. Interestingly, chlorinated compounds are usually more resistant to

microbial attack than brominated compounds, although all can be transformed by

specialist bacteria such as Dehalococcoides mccartyi strain CBDB1 (Cooper et al 2015,

Wagner et al 2012, Yang et al 2015). To study the dehalogenating capacity of the

sediment cultures transforming 1,2,3-TCB, my colleague Myriel Cooper (UFZ–Leipzig)

performed activity tests using a methyl viologen-based resting cell activity assay

(Cooper 2015). Various brominated compounds were selected for the activity test

because brominated compounds are more common than chlorinated compounds in the

marine environment (Ballschmiter 2003). Activity was found for 2,4,6-tribromophenol,

2,4- and 2,6-tribromophenol, 1,2,4-tribromobenzene, and 4-bromo-3,5-

dimethoxybenzoic acid (Cooper 2015). Hexachlorobenzene, hexabromobenzene, PCE

and TCE were not transformed. Most likely, the microorganisms responsible for the

organohalogen transformations may have a smaller set of reductive dehalogenases

(RDases), than for instance, Dehalococcoides mccartyi strains, resulting in a narrower

catalytic spectrum. Moreover, I was not successful in amplifying rdhA genes with the

primers available in the literature that target rdhA genes from Dehalococcoides

mccartyi, Dehalobacter restrictus and Desulfitobacterium spp. (Krajmalnik-Brown et al

2004, von Wintzingerode et al 2001). However, biochemical analysis using

2-iodopropane as a specific inhibitor for B12 enzymes indicated a potential RDase was

performing the 1,2,3-TCB transformation reaction with a cobalamin as a reactive centre

of the enzyme (Cooper 2015). Therefore, the rdhA sequence of the sediment cultures

may differ from the rdhA of Dehalococcoides mccartyi, Dehalobacter restrictus and

Desulfitobacterium spp., suggesting the need to improve rdhA primers to further

Discussion

128

evaluate the potential of organohalide transformation by molecular biology studies in

marine deep sediments. On the contrary, many deep marine sediments that amplified

rdhA genes with the available primers showed no activity when exposed to

organohalides in cultures (Futagami et al 2013). Thus, there is a need for better primers

for functional rdhA genes in the marine subsurface. Other PCR-independent approaches

such as metagenomics could also be an avenue to explore marine subsurface rdhA genes

in the future.

The microorganisms transforming 1,2,3-TCB to 1,3-DCB in sediment cultures were

Gram-positives, as indicated by the lack of organohalide transformation when amended

with either ampicillin or vancomycin. The microbial community in the cultures was

dominated by members of the phylum Firmicutes. Interestingly, members of the

Firmicutes have been suggested as bacteria transforming naturally occurring

organohalides in uncontaminated environments (Krzmarzick et al 2014). Additionally,

several pure strains belonging to Firmicutes, such as Dehalobacter spp. and

Desulfitobacterium spp., are described as dehalogenating organohalogens (Gerritse et al

1996, Holliger et al 1998, Miller et al 1997, Nelson et al 2014, Utkin et al 1994).

The type of the reducing agent used, i.e., titanium (III) citrate or sodium sulphide plus

L-cysteine, had an effect on the composition of the microbial communities as

deciphered by 454-pyrosequencing of 16S rRNA genes. Sediment cultures with either

reducing agents transformed 1,2,3-TCB to 1,3-TCB, and for both cases, the

microorganisms transforming 1,2,3-TCB belonged to the phylum Firmicutes, as

indicated by inhibitor studies. Interestingly, in sediment cultures reduced with titanium

(III) citrate, the genus Anaerobacter took over nearly the entire bacterial community

with a relative abundance of 90% in the most enriched subcultures, which were

transferred four times. Anaerobacter does not appear in the sediment culture line

reduced with sodium sulphide plus L-cysteine suggesting that it may ferment citrate,

and thus titanium (III) citrate enriched for Anaerobacter. Therefore, the use of a specific

reducing agent, as any medium component, strongly affects the microbial community,

especially if it is an organic reducing agent.

For sediment cultures (generation 0) reduced with titanium (III) citrate, members

affiliated to Dehalobacter and Desulfitobacterium were observed, indicating a potential

role for organohalide transformation. However, Desulfitobacterium relative abundance

was very low, and did not increase with dehalogenation and enrichment in any sediment

culture or subculture reduced with any of the reducing agents used here. However,

Desulfitobacterium 16S rRNA gene copy numbers increased with dehalogenation and

Discussion

129

with respect to the start point of incubation for one of the two sediment cultures

investigated, which dehalogenated 1,2,3-TCB. Nevertheless, Desulfitobacterium are

metabolically versatile bacteria that can use other electron acceptors than organohalides

(Villemur et al 2006), and thus an increase in numbers in one sediment culture may not

indicate a role in the organohalide transformation. Most likely, the genus

Desulfitobacterium does not play a substantial role in the transformation of 1,2,3-TCB

in the sediment cultures and enrichments.

On the other hand, Dehalobacter relative abundance increased with time (forming 3%

of the total bacterial community) while dehalogenation occurred in sediment cultures

reduced with titanium (III) citrate. Therefore, the genus Dehalobacter may play a role

for the dehalogenation of 1,2,3-TCB as they are known obligate organohalide-respiring

microorganisms, which cannot derive energy via fermentation or respiration of other

substrates (Holliger et al 1998, Yoshida et al 2009). Additionally, Dehalobacter are

known for respiring both aliphatic, e.g., PCE (Holliger et al 1998), and aromatic, e.g.,

DCBs (Nelson et al 2014). However, no increase in 16S rRNA gene copy numbers was

observed in any of the investigated sediment cultures reduced with titanium (III) citrate.

This may be due to the use of primers specific for Dehalobacter restrictus, which may

not match the Dehalobacter sequences in the sediment cultures. Interestingly, no

increase in Dehalobacter relative abundance was observed in sediment cultures reduced

with sodium sulphide plus L-cysteine. Thus, most likely, Dehalobacter had no role in

dehalogenation in those cultures.

Desulfotomaculum has also been documented to transform organohalides (Duan 2014).

In particular, Desulfotomaculum guttoideum strain VN1 is reported to dechlorinate

1,2,4- and 1,2,3-TCB, and 1,2-DCB, and debrominate hexabromobenzene, 1,2,4-

tribromobenzene and monobromobenzene to benzene (Duan 2014). Interestingly, strain

VN1 used citrate as an electron donor for dechlorination (Duan 2014). Members

affiliated to Desulfotomaculum are observed in sediment cultures reduced with titanium

(III) citrate, with an increase in their relative abundance from 2% at the start point to

15% during dehalogenation. Thus, Desulfotomaculum may play a role in the 1,2,3-TCB

dehalogenation in sediment cultures reduced with titanium (III) citrate, and as observed

for strain VN1, which was isolated from freshwater sediments, may use citrate as an

electron donor in dechlorination. The use of citrate as an electron donor is likely as the

relative abundance of Desulfotomaculum in sediment cultures reduced with sodium

sulphide plus L-cysteine is low (max of 1%). However, no Desulfotomaculum could be

detected in further enrichments reduced with titanium (III) citrate (subcultures G2 and

Discussion

130

G4), and in few proportions (up to 2%) in subcultures reduced with sodium sulphide

plus L-cysteine.

Although Dehalobacter and Desulfotomaculum may have a role in the dehalogenation

of 1,2,3-TCB in sediment cultures reduced with titanium (III) citrate, both of them were

not present in subcultures reduced with titanium (III) citrate and in cultures reduced

with sodium sulphide plus L-cysteine. Thus, there must have been another Firmicutes

member that performed the transformation of 1,2,3-TCB in subcultures and sediment

cultures reduced with sodium sulphide plus L-cysteine. However, no other known

organohalide-respiring microorganism was detected in the cultures. It is possible that

members affiliated to the Desulfosporosinus may be the bacteria dehalogenating 1,2,3-

TCB because Desulfosporosinus were present in the microbial community increasing in

their relative abundance throughout dehalogenation in the sediment cultures reduced

with sodium sulphide plus L-cysteine. Furthermore, Desulfosporosinus phylotypes were

present in further transfers (G2) and were increasing in their relative abundance after

complete dehalogenation of 1,2,3-TCB after two months of incubation (Figure 32). In

addition, Desulfosporosinus phylotypes increased in relative abundance in the most

enriched subculture reduced with titanium (III) citrate after complete dehalogenation

(G4, 149 incubation days; Figure 30.). Desulfosporosinus spp. are known to be

sulphate-reducing bacteria (Ramamoorthy et al 2006, Vatsurina et al 2008), however the

medium used in the present study contains no sulphate, and thus they may respire

alternative electron acceptors other than sulphate in the sediment and transferred

cultures. Interestingly, a novel group within the Firmicutes named the ‘Gopher group’

has been enriched in cultures using halogenated aromatic compounds, i.e., chlorinated

xanthones (Krzmarzick et al 2014). Members of the ‘Gopher group’ are related to the

genera Dehalobacter, Desulfitobacterium and Desulfosporosinus (Krzmarzick et al

2014). The members of the Firmicutes transforming 1,2,3-TCB in sediment cultures in

this study may be also related to the ‘Gopher group’, adding up evidence to the phylum

Firmicutes to be dehalogenating-bacteria in pristine environments such as deep marine

sediments.

4.6 FIRMICUTES ARE COMMON CULTURED BACTERIA FROM MARINE

SEDIMENTS

The bacterial community in sediments from Chile was dominated by members of the

phylum Proteobacteria, with minimal presence of the phylum Firmicutes. However, as

soon as the sediments were exposed to the mineral medium and a temperature of 30ºC,

Firmicutes took over the entire microbial community. Many bacteria from marine

Discussion

131

sediments, e.g., phyla Proteobacteria, Chloroflexi are most likely non-spore forming

microorganisms that may live in a dormant state in sediments, meanwhile other

microorganisms may be in non-vegetative state as spores. Spores may be as abundant as

vegetative cells in deep marine sediments (Lomstein et al 2012). Furthermore, the ratio

of spores to vegetative cells was found to increase with depth (Fichtel et al 2007).

Interestingly, deeper sediments have been observed to have greater microbial

‘culturability’ (Parkes et al 2014), which might be due to higher numbers of spores in

deeper sediment layers. Isolated bacterial members from the marine subsurface belong

to the phyla Actinobacteria, Firmicutes, Proteobacteria and Bacteroidetes. Spore-

forming microorganisms like those from phyla Actinobacteria and Firmicutes may out-

compete marine Dehalococcoidia or any other group when grown in a rich or even any

medium due to its fast growth and the quick response of spore to the favourable

conditions of a nutrient-rich medium.

All isolated colonies from deep-agarose dilution tubes of sediments originating from

Chile site 7155, belonged to the phylum Firmicutes. Similarly, the whole microbial

community evolved into Firmicutes along time in sediment cultures. Together these

results suggest that sediments from Chile site 7155 contained spores which quickly

germinated upon incubation in favourable medium, however, not for the other Chilean

sediments, i.e., site 7165, and the other sediments from other locations, i.e., Århus or

Ireland. Isolation and cultivation of Firmicutes was also observed by other studies and

the great majority of the cultivated bacteria from marine sediments so far belong to the

phylum Firmicutes (D'Hondt et al 2004, Köpke et al 2005, Parkes et al 2009, Süβ et al

2004).

Conclusion

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5 CONCLUSION

This PhD work aimed to gain more insights into microbial life in marine sediments,

with the focus to understand the bacterial class of the Dehalococcoidia, phylum

Chloroflexi, which is widely distributed and highly abundant in marine sediments.

Specifically, the ecological role of marine Dehalococcoidia was investigated within this

study. For that, experiments based on the cultivation of marine sediments (ex situ study)

and the distribution of marine Dehalococcoidia in marine sediments associated to

environmental parameters, i.e., depth, geographical location, geochemistry (in situ

study) were carried out.

Within the ex situ study, the cultivation of various marine sediments with different

potential electron acceptors was carried out aiming for an identification of a specific

respiration mode performed by marine Dehalococcoidia. The different electron

acceptors tested here included halogenated (chlorinated and brominated organohalides)

and non-halogenated compounds (sulphate, manganese(IV), and iron(III)). Halogenated

compounds were selected as known members of the class Dehalococcoidia, such as

Dehalococcoides mccartyi, strictly depend on organohalide respiration. Cultivation of

marine sediments may be challenging when the indigenous microorganisms inhabiting

them may be piezophilic, or have extremely slow metabolic rates. However, the

cultivation experiments carried out within this study showed that marine

Dehalococcoidia can be cultivated in the laboratory at atmospheric pressures and at

temperatures of 30ºC. A medium containing hydrogen as an electron donor and acetate

as carbon source allowed Dehalococcoidia growth and maintenance in reduced anoxic

minimal medium. It has been suggested that microorganisms inhabiting the marine

subsurface may divide on geological time scales (i.e., thousands to millions of years),

however, Dehalococcoidia growth was observed after months of incubation, similarly

to other anaerobic microorganisms, when cultivated with a reduced and anoxic

minimum medium. Results from the cultivation of marine Dehalococcoidia indicated

that they may use terminal electron acceptors other than the organohalides used here,

which included chlorinated and brominated aliphatics and/or aromatics, suggesting that

a respiratory mode other than organohalide respiration is likely. From all the tested

organohalides, only 1,2,3-TCB was dehalogenated to 1,3-DCB in sediment cultures.

However, in-depth investigations of the enrichment cultures over four generations

indicated no role for members of the Dehalococcoidia in the dehalogenation. On the

contrary, enrichment of members of the family Peptococcaceae, phylum Firmicutes,

was attained.

Conclusion

133

Additionally, the cultivation of marine Dehalococcoidia with other potential electron

acceptors such as sulphate, iron(III), manganese(IV), and humic acids indicated that for

some sediments, i.e., from Århus and Ireland, sulphate promoted an increase in marine

Dehalococcoidia 16S rRNA gene copy numbers, however, not for sediments from

Chile, where none of the tested electron acceptor promoted a sustained Dehalococcoidia

growth with time. Thus, marine Dehalococcoidia may have diverse respiratory modes.

Therefore, an approach per clade, instead of per class, may be more useful in order to

decipher the respiratory mode of specific groups within the class Dehalococcoidia. In

addition, high Dehalococcoidia 16S rRNA gene copy numbers in sediment cultures

amended with no electron acceptor indicated that a respiratory mode other than the

tested in the current study may also be likely.

Within the in situ study, the distribution of marine Dehalococcoidia in sediments of the

Baffin Bay, in the Arctic, was investigated. Several sites at distinct geographical

locations within the Baffin Bay were analysed for its geochemistry and microbial

composition. The focus was placed on the identification of natural occurring conditions

promoting Dehalococcoidia abundance. These natural conditions may include biotic

factors, i.e., Dehalococcoidia may be associated to the presence of other bacterial

groups, or abiotic factors, i.e., Dehalococcoidia may be associated to specific

environmental parameters such as depth or sediment geochemistry. Dehalococcoidia

members were present in all sediment sites and depths from different geographical

locations and different sediment geochemical conditions within the Baffin Bay,

indicating a wide distribution. The highest abundance of Dehalococcoidia was at shelf

sites, which were richer in organic matter than basin sites. However, Dehalococcoidia

accounted for the greatest proportion of the total bacterial 16S rRNA gene copy

numbers (sometimes higher than 50%) in deeper core sections where low Bacteria were

found, particularly at the central deep basin of the Baffin Bay, indicating that

Dehalococcoidia members are bacteria that are resilient to burial.

When associations of microbial distributions in relation with geochemical parameters in

the Baffin Bay were studied, Dehalococcoidia and particularly the clade GIF-9,

correlated positively with organic matter and negatively with sulphate and

manganese(II) concentrations. Other bacteria that showed similar correlations to

geochemical parameters were members of the candidate phyla “JS1” and “OP8”. In fact,

higher relative abundance of JS1 and OP8 were observed at sites and depths where

Dehalococcoidia relative abundance was also higher. These depths and sites were

grouped (by hierarchical clustering) in a cluster containing near-surface sediments from

Conclusion

134

all sites, and deeper layers from shelf sites. A bacterial phylum that correlated

differently to the geochemical parameters in the Baffin Bay was the Proteobacteria.

More specifically, the class Betaproteobacteria positively correlated to manganese(II)

and negatively to organic matter. Additionally, about 40% of the members belonging to

the class Alphaproteobacteria correlated positively to sulphate and another 40% to

iron(II) concentration.

Altogether, the identification of natural conditions in the Baffin Bay promoting higher

Dehalococcoidia relative abundance (positive correlation to organic matter, and

negative to sulphate and manganese), and quantified abundance (higher

Dehalococcoidia 16S rRNA gene copy numbers in the shelf as measured with qPCR),

and intrinsic characteristics observed (high resilience at sites and depths where low

overall bacteria were found), together with the cultivation with various electron

acceptors described in the current PhD thesis, provide a better understanding of the

ecological role of marine Dehalococcoidia in marine sediments.

Appendix

135

6 APPENDIX

6.1 APPENDIX 1

The medium was prepared from sterile concentrated solutions (20 ml mineral Widdel

solution; 10 ml trace metal SL9 solution; 5 ml sodium acetate 1 M and 50 µl of the

redox indicator resazurin) and brought up to 1 litre with Milli Q water. The content of

the stock solutions used in the medium and the respective components and

concentrations is the following:

6.1.1 Mineral solution “Widdel solution”

Stock solution 50x (Widdel 1980):

KH2PO4 10 g l-1

NH4Cl 13.5 g l-1

NaCl 50 g l-1

MgCl2 x 6H2O 20.5 g l-1

KCl 26.0 g l-1

CaCl2 x 2H2O 7.5 g l-1

Sterilization by autoclaving at 121ºC for 40 min.

6.1.2 Trace metal solution “SL-9 solution”

Stock solution 100x (Tschech and Pfennig 1984):

H2O 500 ml

(CH₂CO₂H)₃, (NTA) 12.8 g

FeCl2 x 4H2O 2 g

ZnCl2 70 mg

MnCl2 x 2H2O 80 mg

H3BO3 6 mg

CoCl2 x 6H2O 190 mg

CuCl2 x 2H2O 2 mg

NiCl2 x 6H2O 24 mg

Na2MoO4 x 2H2O 36 mg

NaOH added up to pH 6.0

H2O added up to 1000 ml

Sterilization by autoclaving at 121ºC for 40 min.

6.1.3 Bicarbonate solution

For the bicarbonate solution, 7.06 g of NaHCO3 were dissolved in 84 ml of anaerobic

and sterile Milli Q water, which was previously saturated with CO2, sealed under a

headspace atmosphere of CO2. It was sterilized by autoclaving.

Appendix

136

6.1.4 Vitamin 7 solution

Stock solution of 2000x (Adrian 1999, Widdel 1980):

4-Aminobenzoic acid 40 mg l-1

D (+)-Biotin 10 mg l-1

Niacin 100 mg l-1

Ca-D (+) pantothenate 50 mg l-1

Pyridoxine hydrochloride 150 mg l-1

Thiamine chloride-di-hydrochloride 100 mg l-1

Cyanocobalamin 100 mg l-1

Sterilization by filtering using a Minisart filter with a 0.2 μm pore size (Sartorius), and

preserved in the dark at 4ºC

Addition of 0.5 ml of vitamin solution to 1 litre of medium

6.1.5 Titanium (III) citrate solution as reducing agent

The titanium (III) citrate solution was prepared according to (Zehnder and Wuhrmann

1976). For that, an anoxic sodium citrate solution (1 M; 10 ml) was mixed with a

titanium (III) chloride solution (15%; 5.14 ml) under a stream of nitrogen. The final

solution was adjusted to a pH of 7.0 with sodium carbonate (added as solid) and diluted

with anoxic water to obtain a final concentration of 0.1 M of titanium (III) and 0.2 M

citrate. The final titanium (III) citrate solution was filter-sterilized, and stored in a sterile

brown-glass vial with biogon (80% N2, 20% CO2 v/v) in the vial headspace, which was

kept in the dark.

6.1.6 Iron sulphide solution as reducing agent

For the preparation of the iron sulphide (FeS) solution, two solutions were prepared and

then mixed. The first solution was Na2S in a concentration of 400 mM. The second

solution was FeCl2 in a concentration of 10 mM. Both solutions were filter-sterilized by

using a Minisart filter with a 0.2 μm pore size (Sartorius). Then, 12 ml of the FeCl2

solution was supplied to a sterile anoxic glass vial, and 0.3 ml of the Na2S solution was

added. The FeS solution was always mixed before used. All reducing agent solutions

were prepared shortly before used.

Appendix

137

6.1.7 Medium composition and concentration

Substance Concentration

Salts mM

KH2PO4 1.5

KCl 7.0

NH4Cl 5.0

NaCl 17.1

MgCl2 x 6H2O 2.0

CaCl2 x 2H2O 1.0

Buffer mM

NaHCO3 30.0

Trace elements µM

Iron, FeCl2 x 4H2O 10.060

Boron, H3BO3 0.098

Manganese, MnCl2 x 2H2O 0.494

Cobalt, CoCl x 6H2O 0.799

Nickel, NiCl2 x 6H2O 0.088

Copper, CuCl2 x 2H2O 0.012

Zinc, ZnCl2 0.514

Molybdenum, Na2MoO4 x 2H2O 0.149

+ NTA as a complexing agent

Vitamins µM

4-Aminobenzoic acid (PABA), C7H7NO2 0.15

D (+) – Biotin (B7), C10H16N2O3S 0.02

Niacin (B3), C6H5NO2 0.41

Ca-D (+)-pantothenate (B5), C18H32CaN2O10 0.10

Pyridoxine hydrochloride (B6), C8H12ClNO3 0.36

Thiamine (B1), C12H17N4OS 0.15

Cyanocobalamin (B12), C63H88CoN14O14P 0.04

Reducing agent

Titanium (III) citrate ~1.4 mM in regard to Ti(III)

Sulphide / L-cysteine 0.3 mM / 0.2 mM

Redox indicator

Resazurin 0.5 mg l-1

Appendix

138

6.2 APPENDIX 2

Cores (divided into one-metre sections) investigated in this study from the sampled

sediment sites during the ARK XXV/3 expedition at the Baffin Bay. Photos are

courtesy of Dr Thomas Pletsch from BGR-Hannover, Germany.

Site 363 Site 365

Appendix

139

Site 371 Site 383

Appendix

140

Site 387 Site 389

Appendix

141

Site 391 Site 453

Appendix

142

Site 486 Site 488

Appendix

143

6.3 APPENDIX 3

Figure 55. Iron oxide (Fe2O3; upper panel) and manganese oxide (MnO; lower panel) percentages in the

mineral solid phase of sediments from the Baffin Bay. Shown are depth profiles of three sites that were

selected, one site per area (Northern Greenlandic shelf, central deep basin, Southern slope), for elemental

composition of the mineral fraction in sediment samples analysed at the BGR-Hannover.

References

144

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