Struktur und Dynamik heterotropher Bakteriengemeinschaften ...oops.uni-oldenburg.de/42/1/rinstr07.pdf · bacteria affiliated to the Bacteroidetes phylum, gamma- and delta-Proteobacteria

Struktur und Dynamik

heterotropher Bakteriengemeinschaften im Wattenmeer

und der Deutschen Bucht

Structure and dynamics of heterotrophic bacterial

communities in the German Wadden Sea

and the German Bight

Dissertation

zur Erlangung des akademischen Grades einer

Doktorin der Naturwissenschaften (Dr. rer. nat.)

der Fakultät V Mathematik und Naturwissenschaften

der Carl von Ossietzky Universität Oldenburg

vorgelegt von

Beate Rink

geboren am 23.01.1974 in Bremerhaven

http://docserver.bis.uni-oldenburg.de/publikationen/dissertation/2007/rinstr07/rinstr07.html

http://docserver.bis.uni-oldenburg.de/publikationen/dissertation/2007/rinstr07/rinstr07.html

Erstgutachter : Prof. Dr. Meinhard Simon

Zweitgutachter: Prof. Dr. Heribert Cypionka

Eingereicht am:

Disputation am:

Für Rosemarie

Erklärung

Teilergebnisse dieser Arbeit sind als Beiträge bei den genannten Fachzeitschriften eingereicht

oder werden eingereicht. Mein Beitrag an der Erstellung der verschiedenen Manuskripte wird

im Folgenden erläutert:

Rink, B., Seeberger, S., Martens, T., Duerselen, C. D., Simon, M., und Brinkhoff, T. (2006)

Effects of a phytoplankton bloom in a coastal ecosystem on the composition of bacterial

communities (Eingereicht bei Aquat. Microb. Ecol.)

Etablierung und Spezifitätstest der Roseobacter spezifischen PCR durch S. S. unter Anleitung

von B. R. und T. B (Diplomarbeit, 2003). Durchführung der spezifischen PCR und DGGE,

der Klonierung und Sequenzierung durch B. R. Statistische Auswertung und Erstellung der

phylogenetischen Stammbäume durch B. R. Erstellung der ersten Fassung des Manuskripts

durch B. R., Überarbeitung durch T. B., B. R. und M. S.

Rink, B., Martens, T., Fischer, D., Lemke, A., Grossart, H. P., Simon, M., und Brinkhoff, T.

(2006) Tidal effects on coastal bacterioplankton (In Vorbereitung zum Einreichen bei

Limnol. Oceanogr.)

Planung und Durchführung der Probenahme 2005 durch B. R. Durchführung der spezifischen

PCR und DGGE sowie der RNA Untersuchungen und CARD-FISH durch B. R. Statistische

Auswertung und Erstellung der phylogenetischen Stammbäume durch B. R. Erstellung der

ersten Fassung des Manuskripts durch B. R., Überarbeitung durch T. B., B. R. und M. S.

Stevens, H., Brinkhoff, T., Rink, B., Vollmers, J., und Simon, M. (2006) Diversity and

abundance of Gram-positive bacteria in a tidal flat ecosystem (Eingereicht bei Environ.

Microbiol.)

Durchführung der spezifischen CARD-FISH und DGGE Untersuchungen von J. V. unter

Anleitung von B. R. und T. B (Leistungsnachweis, 2005). Überarbeitung des Manuskriptes

von B. R., T. B. und M. S.

Rink, B., Brinkhoff, T., Ziegelmüller, K., und Simon, M. (2006) High regional variability of

bacterial communities in the German Bight, North Sea (Eingereicht bei Aquat. Microb.

Ecol.)

Planung und Durchführung der Probenahme 2002 von Mirko Lunau und B. R. Planung und

Durchführung der Probenahme 2003 von B. R. Molekularbiologische Untersuchungen (PCR,

DGGE, Klonierung, Sequenzierung), statistische Auswertung und Erstellung der

phylogenetischen Stammbäume durch B. R. Erstellung der ersten Fassung des Manuskripts

durch B. R., Überarbeitung durch T. B., B. R. und M. S.

Tagungsbeiträge

Rink B, Stevens H, Simon M, Brinkhoff T (2006) Stability of Microbial Communities Within

Different Time Scales in a Tidal Flat Ecosystem. Posterbeitrag, International

Symposium Microbial Ecology (ISME-11), Wien, Österreich, 20-25 August

Rink B, Brinkhoff T, Simon M (2004) Bacterial communities reflect different regional

properties of the German Bight. Vortrag, VAAM-Meeting Braunschweig, 28-31 March

Rink B, Kruse M, Seeberger S, Stevens H, Brinkhoff T, Simon M (2004) Seasonal and spatial

differences in the composition and abundance of bacterial communities in the German

Bight of the North Sea. Posterbeitrag, International Symposium Microbial Ecology

(ISME-10), Cancun, Mexico, 22-27 August

Simon M, Selje N, Schledjewski R, Rink B, Grossart HP (2004) Diversity and substrate

turnover of bacterioplankton communities in the Gulf of Aqaba, Red Sea. Posterbeitrag,

International Symposium Microbial Ecology (ISME-10), Cancun, Mexico, 22-27

August

Rink B, Lunau M, Seeberger S, Stevens H, Brinkhoff T, Grossart H-P, Simon M (2003)

Diversity patterns of aggregate-associated and free-living bacterial communities in the

German Wadden Sea. In Rullkötter J. (ed.), BioGeoChemistry of Tidal Flats -

Proceedings of a Workshop held at the Hanse Institute of Advanced Study, Delmenhorst

(Germany), 14- 17 May. Forschungszentrum Terramare, Wilhelmshaven, Berichte Nr.

12, 96-98. ISSN 1432-797X.

Lunau M, Rink B, Grossart H-P, Simon M (2003) How to sample marine microaggregates in

shallow and turbid environments? - Problems and solutions. In Rullkötter J. (ed.),

BioGeoChemistry of Tidal Flats - Proceedings of a Workshop held at the Hanse

Institute of Advanced Study, Delmenhorst (Germany), 14-17 May. Forschungszentrum

Terramare, Wilhelmshaven, Berichte Nr. 12, 85-88. ISSN 1432-797X.

Rink B, Brinkhoff T, Simon M (2002) Completing the picture of natural habitats: The use of

specific Primersets in DGGE. Posterbeitrag, VAAM Meeting, Berlin, 23-26 March

Zusammenfassung

Im Wattenmeer unterliegen die Organismen hochdynamischen Prozessen. Eine flache

Wassersäule und der Einfluß der Gezeiten sorgen für starke Strömungen und hohe

Resuspensionsraten. Auch der tidale Ein- und Ausstrom von Wassermassen aus der Nordsee

in das Rückseitenwatt beeinflusst das System. Während in Herbst- und Wintermonaten

sedimentologische Faktoren überwiegen, ist im Frühjahr und Sommer ein deutlicher Einfluss

biologischer Größen nachweisbar. Im Rahmen des interdisziplinären Forschungsprojekts

„Biogeochemie des Watts“, in das diese Arbeit eingebunden ist, wurden große Varianzen

innerhalb des Schwebstoffaufkommens sowie in bakterieller Aktivität und Abundanz auf

saisonaler Ebene sowie im Tidenzyklus beschrieben.

In der vorliegenden Arbeit wurde untersucht, inwiefern tidale und saisonale Faktoren die

Struktur der ansässigen Bakteriengemeinschaften in der Wassersäule beeinflussen.

Weiterführend wurde untersucht, ob die im Wattenmeer detektierten Phylotypen

standortspezifisch oder auch in anderen Gebieten der Deutschen Bucht nachweisbar sind.

Im Wattenmeer fand die Beprobung in der Otzumer Balje im Rückseitenwatt von Spiekeroog

statt. Im ersten Teil dieser Arbeit wurden zur Untersuchung des Zusammenhangs von

Bakteriengemeinschaften und Phytoplankton wöchentlich Proben genommen und mittels

gruppenspezifischer DGGE (Denaturierende Gradienten Gelelektrophorese) und statistischer

Methoden untersucht. Im zweiten Teil wurden neben saisonalen auch tidale Vorgänge

beleuchtet. Die Probenahme fand im Herbst, Frühjahr und Sommer in einstündigem und

dreistündigem Probenahmeraster statt. Die Bakteriengemeinschaften wurden mittels

gruppenspezifischer DGGE für alpha-Proteobakterien, Bacteroidetes und Roseobacter

sowohl DNA- als auch RNA basiert untersucht. Zusätzlich wurden FISH (Fluoreszens in situ

Hybridisierung) und die hoch sensitive CARD-FISH (Catalyzed Reporter Deposition-FISH)

eingesetzt und somit erstmalig die Abundanzen einzelner Bakteriengruppen in der

Wassersäule des Wattenmeeres dargestellt.

In einer vorangegangenen Arbeit wurden im Wattenmeer bemerkenswert viele gram-positive

Bakterien isoliert, was zu der Annahme führte, dass diese Bakteriengruppe eine besondere

Stellung in diesem Habitat einnimmt. Zur Vervollständigung der Daten wurde im dritten Teil

dieser Arbeit die CARD-FISH eine Actinobakterien-spezifische Sonde eingesetzt und

zusätzlich eine spezifische DGGE entwickelt, um Abundanz und phylogenetische Vielfalt der

Actinobakterien im Watt zu untersuchen. Die Probenahme hierzu wurde an verschiedenen

Standorten im Spiekerooger Rückseitenwatt durchgeführt.

Im vierten Teil wurden im Sommer 2002 und 2003 verschiedene Standorte der Deutschen

Bucht an der Küstenzone, vor Helgoland und in der offenen Nordsee beprobt. Die

Bakteriengemeinschaften wurden mit spezifischer DGGE für alpha-Proteobakterien und

Bacteroidetes untersucht. Zur weiteren Beschreibung der Ökologie an den untersuchten

Standorten wurden zusätzlich hydrologische, mikrobiologische und partikuläre Parameter

bestimmt.

Zusammenfassend ergaben sich aus diesen Arbeiten folgende Hauptaussagen:

• Im Wattenmeer sind die Bakteriengemeinschaften in der Wassersäule im

Wesentlichen aus alpha- und gamma-Proteobakterien sowie Bacteroidetes

zusammengesetzt. Darüber hinaus sind beta-Proteobakterien abundant auf

Aggregaten. Hierbei bilden frei lebende und Aggregat-assoziierte Bakterien distinkte

Gemeinschaften sowohl im Wattenmeer als auch in der Deutschen Bucht. Die Struktur

der frei lebender Bakteriengemeinschaften besteht hauptsächlich aus wenigen

dominanten Phylotypen der Roseobacter Gruppe. Ihre Zusammensetzung ist saisonal

und räumlich stabil. Die Struktur der Aggregat-assoziierten Bakterien zeigt grössere

Artenvielfalt als bei frei lebenden Bakterien und unterliegt deutlicher räumlich-

zeitlichen Einflussfaktoren. Hier dominieren Phylotypen innerhalb der Bacteroidetes,

gamma- und delta-Proteobakterien.

• Saisonale Einflüsse auf die Bakteriengemeinschaften sind in den produktiven

Frühjahrs- und Sommermonaten erkennbar. Insbesondere Aggregat-assoziierte

Bakterien der Roseobacter-Gruppe und Bacteroidetes unterliegen biologischen

Einflussfaktoren wie Phytoplanktonblüten. Tidale Einflüsse auf bakterielle Aktivität

und Abundanz werden nur geringfügig und nicht systematisch durch Änderungen in

der Zusammensetzung der Bakteriengemeinschaften reflektiert.

• Actinobakterien stellen knapp 5% des Bakterioplanktons im Wattenmeer. Ihre

Abundanz und Zusammensetzung im Süßwasserbereich unterscheidet sich von den

marinen Standorten, wobei frei lebende und Aggregat-assoziierte Actinobakterien

distinkte Gemeinschaften bilden. Aus dem Wattenmeer isolierte Stämme zeigen hohe

Anpassungsfähigkeit anhand breiter Substrat- und Salinitätsspektren.

• Insgesamt wird das organische Material im Wattenmeer von wenigen dominanten

Bakterienarten umgesetzt, die ganzjährig auftreten und hoch angepasst sind. In

produktiven Jahreszeiten treten darüber hinaus weitere, spezialisierte Bakterienarten

auf, die in kurzen Zeitskalen von Änderungen der Zusammensetzung des organischen

Materials, z. B. durch absterbendes Phytoplanton, profitieren.

Summary

In the German Wadden Sea, organisms are influenced by highly dynamic processes. A

shallow water column and tidal impact cause strong currents and high resuspension rates. The

introduction of North Sea water masses also influences the Wadden Sea System. While

sedimentological factors prevail in autumn and winter months, biological processes dominate

in spring and summer. Within the research group “Biogeochemistry of tidal flats”, in which

this thesis is included, tidal and seasonal variations of suspended matter appearance and

bacterial activity and abundance were described.

Hence, the focus of this thesis was to investigate the extend of tidal and seasonal impacts on

the structure of resident bacterial communities in the water column. Furthermore, we

determined if phylotypes detected in the German Wadden Sea are site-specific or detectable at

other locations in the German Bight as well.

Sampling was performed in the backbarrier tidal flat system of Spiekeroog in the German

Wadden Sea. In the first part of this work, samples were taken weekly to investigate

correlations of the bacterial communities and phytoplankton by group-specific DGGE

(Denaturing gradient gel electrophoresis) and statistical methods. In the second part, in

addition to seasonal also tidal processes were focussed. Sampling was performed in autumn,

spring and summer hourly and in three hour intervals. The bacterial communities were

investigated by group-specific DGGE (Denaturing gradient gel electrophoresis) for alpha-

Proteobacteria, Bacteroidetes and the Roseobacter group. In addition, FISH (Fluorescense in

situ hybridization) and the highly sensitive CARD-FISH (Catalyzed reporter deposition-

FISH) were applied to determine abundances of individual bacterial groups in the water

column of the German Wadden Sea.

In a former study, remarkably high numbers of different gram-positive Bacteria were isolated

which led to the assumption that this bacterial group exhibits an exceptional position in this

habitat. To complete these data, CARD-FISH with Actinobacteria-specific probes was

applied and a specific DGGE was established to determine abundances and phylogenetic

variety of Actinobacteria in the Wadden Sea. Samples were taken at different sites in the

backbarrier tidal flat system of Spiekeroog.

In the last part of this work, different locations at the coastal line, near Helgoland and offshore

were investigated in the German Bight in summer 2002 and 2003. The bacterial communities

were analysed by specific DGGE for alpha-Proteobacteria and Bacteroidetes. To describe the

ecology of the sampling sites hydrological, microbiological and particulate parameters were

determined additionally.

The major findings of this thesis can be summarized as follows:

• The bacterial communities in the water column of the German Wadden Sea are mainly

composed of alpha- and gamma-Proteobacteria and Bacteroidetes. In addition, beta-

Proteobacteria are abundant on aggregates. Free-living and aggregate-associated

bacteria form distinct communities in the German Wadden Sea and in the German

Bight as well. The structure of free-living bacterial communities is mainly composed

of few dominant phylotypes affiliated to the Roseobacter group. Their composition is

stable on seasonal and spatial scales. The structure of aggregate-associated bacteria

shows higher richness compared to free-living bacteria and is influenced by spatial-

temporal impacts to a greater extend. Aggregate-associated bacteria are dominated by

bacteria affiliated to the Bacteroidetes phylum, gamma- and delta-Proteobacteria.

• Seasonal influences on the bacterial communities are detectable in the highly

productive spring and summer months. Especially aggregate-associated Roseobacter

and the Bacteroidetes follow biological impacts e. g. phytoplankton blooms. Tidal

influences on bacterial activities and abundances are only marginally and not

systematically reflected by changes of the bacterial community composition.

• Actinobacteria represent about 5% of the Wadden Sea bacterioplankton. Their

abundance and composition differs between the fresh water and marine sites, and free-

living and aggregate-associated bacteria form distinct communities. Strains isolated

from the Wadden Sea show high adaptation qualities on the basis of broad substrate

and salinity ranges.

• The organic matter in the Wadden Sea is mediated by few dominant bacterial species

which are present throughout the year and are highly adapted. In productive seasons,

specialised bacteria appear additionally which benefit from the changes of the organic

matter composition, e. g. decaying phytoplankton, on small time-scales.

Inhaltsverzeichnis

Zusammenfassung

Summary

I. Einleitung 1

I.1 Kleine Lebewesen, große Wirkung – Marine heterotrophe Bakterien im

globalen Stoffkreislauf 2

I.2 Who´s who – Die Zusammensetzung der Bakteriengemeinschaften 5

I. 3 Geographie und Ökologie der Untersuchungsgebiete 8

I.3.1 Die Nordsee und die Deutsche Bucht 8

I.3.2 Das Wattenmeer 11

I.4 Zielsetzungen der Arbeit 15

I.5 Literatur 16

II. Effects of a phytoplankton bloom in a coastal ecosystem on the composition

of bacterial communities 20

Abstract 22

Introduction 23

Materials and Methods 24

Results 28

Discussion 32

Literature cited 36

III. Tidal effects on coastal bacterioplankton 48

Abstract 51

Introduction 52


Results 56

Discussion 60

References 66

IV. Diversity and abundance of Gram-positive bacteria in a tidal flat ecosystem 81

Abstract 83

Introduction 84

Results 85

Discussion 88

Experimental procedures 93

References 98

V. High regional variability of bacterial communities in the German Bight,

North Sea 114

Abstract 116

Introduction 117


Results 121

Discussion 124

References 130

VI. Schlussbetrachtung und Ausblick 144

Danksagung

Kurzbiographie

Abkürzungsverzeichnis

CARD-FISH catalyzed reporter deposition-FISH

Chl a Chlorophyll a

CO2 Kohlendioxid

DAPI 4´,6´-Diamidino-2-phenylindol hydrochlorid

DGGE Denaturierende Gradienten Gelelektrophorese

DNA desoxy ribonucleic acid

DOC dissolved organic carbon

DOM dissolved organic matter

DSM, DSMZ Deutsche Sammlung von Mikroorganismen und Zellkulturen

et al. et alii

FISH fluorescence in situ hybridization

FL free living

HT high tide

LT low tide

ml Milliliter

MT mean tide

NCBI National Center for Biotechnology Information

n. a. not available

n. d. not determined

PA particle attached

PCR polymerase chain reaction

PIC particulate inorganic carbon

POC particulate organic carbon

psu practical salinity unit

rRNA ribosomal ribonucleic acid

SPM suspended particulate matter

1

I.

Allgemeine Einleitung

Kapitel I Einleitung

2

I.1 Kleine Lebewesen, große Wirkung – Marine heterotrophe Bakterien im globalen

Stoffkreislauf

Bei der Betrachtung der Gesamtgröße der Weltmeere erscheint es zunächst kaum vorstellbar,

dass mikroskopisch kleine Lebewesen den Großteil des Umsatzes organischen Materials im

Wasser bewirken sollen. Berücksichtigt man allerdings, dass in einem tausendstel Liter

bereits durchschnittlich 1-3 Mio. Bakterien vorhanden sind, ist offenbar, warum sich die

Forschung seit mehr als zwei Jahrzehnten bemüht, diese höchst bemerkenswerten Lebewesen

besser kennen zu lernen. Bakterieller Abbau und Remineralisierung wirken sich auf den

Stoffkreislauf aller Elemente aus (Schlegel, 1992; Madigan et al., 2003), wobei der

Kohlenstoffkreislauf große Bedeutung nicht zuletzt für klimatische Veränderungen besitzt. So

hat die Aktivität mariner phototropher und heterotropher Bakterien sowohl durch die

Fixierung als auch durch den Ausstoß von CO2 Einfluss auf das Weltklima (Smith und

Hollibaugh, 1993; Wollast, 1993; Falkowski et al., 1998), so dass der marinen mikrobiellen

Ökologie im Zuge der globalen Erwärmung immer größere Bedeutung beigemessen wird.

Auch das genetische Potential der marinen Bakterien, das durch moderne Methoden zwar

detektiert, aber bei weitem noch nicht entschlüsselt wurde, gibt der Wissenschaft Rätsel auf

(Venter et al., 2004). Die zukunftsträchtige und viel versprechende Vision einer möglichen

medizinischen oder biotechnologischen Nutzung mariner Mikroorganismen bietet daher ein

weiteres großes Interessensgebiet der ökologischen Forschung.

Die Nahrungsquelle heterotropher Bakterien, organischer Kohlenstoff, liegt in der

Wassersäule in gelöster (dissolved organic carbon, DOC) oder in partikulär gebundener Form

(particulate organic carbon, POC) vor.

DOC umfasst bis zu 95% des Gesamtkohlenstoffs der Weltmeere (Hedges, 1992) und wird

nach Zusammensetzung und bakterieller Verfügbarkeit in eine labile und eine refraktäre

Fraktion unterteilt (Søndergaard und Middelboe, 1995). Die Entstehung von gelöstem

organischen Material (dissolved organic matter, DOM) ist durch einen komplexen Kreislauf

gekennzeichnet, in dem im Wesentlichen Phytoplankton, Bakterien und Viren eine Rolle

spielen (Abb. 1). Anhand der Darstellung wird deutlich, dass Bakterien DOM direkt

aufnehmen und verwerten und somit die größte Bedeutung für die Umsetzung des DOC

besitzen. Durch das daraus resultierende Wachstum und den Fraß durch Protozoen gelangt der

Kohlenstoff dann indirekt in das Nahrungsnetz höherer Organismen. Die Entstehung des

DOM ist abhängig von verschiedenen Faktoren, da die am Kreislauf beteiligten Organismen


3

räumlich-zeitlichen Gegebenheiten unterliegen. Phytoplankton bildet bei ausreichender

Nährstoffversorgung und günstigen Licht- und Temperaturverhältnissen Blüten aus, die in

direkter Form zu der Absonderung („Leakage“) von DOM führen kann (Bjørnsen, 1988).

Nach Absterben der Blüte werden durch Lysis der Zellen ebenfalls gelöste Stoffe freigesetzt.

Darüber hinaus entsteht bei diesem Vorgang Detritus (partikuläre Zellreste), der durch

Bakterien hydrolysiert und somit dem DOM Pool zugeführt wird.

Neben dem DOC stellt in Aggregaten angereicherter POC eine weitere wichtige organische

Kohlenstoffquelle dar (Alldredge, 1979) und bildet die Grundlage für komplexe Lebens-

gemeinschaften, die sich im Weiteren aus Phytoplankton, Protozoen, Bakterien und Pilzen

zusammensetzen (Alldredge and Silver, 1988). An Aggregate angeheftet, weisen sie,

verglichen mit frei suspendierten Mikroorganismen, wesentlich höhere Zelldichten auf

(Simon et al., 2002). Je nach Ursprung des partikulären Materials variieren der organische

Anteil sowie dessen Zusammensetzung aus Kohlenhydraten und Proteinen, die den Bakterien

als Substrat dienen (Azam und Cho, 1987; Smith et al., 1995, Azam und Cho, 1987; Biddanda

und Benner, 1997). Auch hier ist das Phytoplankton, abhängig von seiner

Artenzusammensetzung, hauptsächlicher Nährstofflieferant (Smith et al., 1995); weitere

Bestandteile von Aggregaten sind hochrefraktäres oder auch anorganisches Material, z. B.

resuspendiertes Sediment (Eisma, 1993). Die Freisetzung der Nährstoffe durch Bakterien

erfolgt durch die Ausscheidung hydrolytischer Ektoenzyme, die partikulär gebundene

Abb. 1: Kreislauf gelösten organischen Kohlenstoffs in der Wassersäule (dissolved organic matter, DOM; modifiziert nach Riemann, 2001)


4

Makromoleküle in Oligo- und Monomere spalten (Madigan, 2003). Die hydrolysierten Stoffe

werden teils von den Aggregat-assoziierten Bakterien selbst verwertet, teils diffundieren sie

jedoch auch in das Umgebungswasser und stehen somit den frei suspendierten Bakterien und

anderen planktischen Organismen zur Verfügung (Smith et al., 1992). Durch die

Substrataufnahme wachsen die Bakterien und bilden somit Biomasse, die wiederum

Zooplankton als Nahrungsquelle dient. Durch diesen Stoffkreislauf, der als microbial loop

bezeichnet wird, werden Nährstoffe aus abgestorbenen Tier- und Pflanzenresten (Detritus) für

höhere Trophiestufen wieder verfügbar (Azam et al., 1983).

In flachen Küstenzonen, Wattsystemen und Ästuaren unterscheidet sich die Situation im

Vergleich zu den offenen Ozeanen durch ein sehr hohes Schwebstoffaufkommen. Die

Schwebstoffe werden hier durch Flüsse oder von den angrenzenden Landgebieten eingetragen

und sind zum Teil hohen Scherkräften ausgesetzt, die durch die flache Wassersäule und

Tidenhub entstehen. Dadurch sind die Aggregate in den Küstenzonen wesentlich kleiner und

häufiger (Lunau et al., 2006) und besitzen, verglichen mit Schwebstoffen in küstenfernen

Gebieten, einen geringeren organischen Anteil (Postma, 1981; Lunau et al., 2006). Durch

ständige Turbulenz werden die Aggregate fortwährend resuspendiert und somit in Schwebe

gehalten, was ebenfalls Auswirkungen auf die angehefteten Bakterien hat. So weisen

Aggregat-assoziierte Bakterien in schwebstoffreichen Gewässern wesentlich höhere

Enzymaktivitäten und Biomasse auf, und können sogar bis zu 95% der Gesamtaktivität der

suspendierten Bakterien ausmachen (Crump et al., 1998; Crump & Baross, 2000).

Auch die Bakteriengemeinschaften können in Küstenzonen anders zusammengesetzt sein als

in zulaufenden Flüssen oder im offenen Meer. So bilden sich entweder Mischformen von

Süßwasser- und marinen Bakteriengemeinschaften (Rappé et al., 2000), oder auch distinkte

Bakteriengemeinschaften von Süßwasser, Brackwasser und marinem Milieu aus (Selje et al.,

2003, Crump et al. 1999).

Diese Zusammenhänge verdeutlichen, in welchem Umfang Bakterien das gesamte

Nahrungsnetz beeinflussen und dass freilebende und Aggregat-assoziierte Bakterien

vollkommen unterschiedliche Lebensbedingungen vorfinden. Daher ist eine differenzierte

Untersuchung beider Lebensgemeinschaften essentiell, um die ökologischen Zusammenhänge

in der Wassersäule verstehen zu können.


5

I.2 Who´s who – Die Zusammensetzung der Bakteriengemeinschaften

Da Bakterien unter dem Mikroskop und in der Kultivierung nur sehr wenige Unterschiede

anhand von Zellmorphologie und Wachstum aufweisen, wurde die Artenvielfalt von

natürlichen Bakteriengemeinschaften lange Zeit unterschätzt. Darüber hinaus bot die

Kultivierung nur bedingt Einblick in das Vorkommen und die Häufigkeit von Bakterienarten,

da die Bedingungen, die Bakterien im Labor vorfinden, nicht den natürlichen Gegebenheiten

entsprachen. Einige Bakterienstämme oder auch phylogenetische Gruppen konnten leicht

unter künstlichen Bedingungen angereichert werden und wurden somit auch häufiger in

verschiedenen Habitaten nachgewiesen, während sich andere Bakterien nur unter bestimmten

Voraussetzungen kultivieren ließen oder bis heute unkultiviert bleiben. Daher ergaben sich

große Unterschiede zwischen mikroskopisch und durch Kultivierungsansätze ermittelte

Zellzahlen („great plate count anomaly“, Staley & Konopka, 1985).

So brachte die Einführung molekularbiologischer Methoden, die auf dem Vergleich des

Erbguts anhand der ribosomalen RNA beruhten, neue Einblicke in die mikrobielle Ökologie

und die phylogenetischen Zusammenhänge (Woese et al., 1987). Bis heute stellt die

hochkonservierte 16S rRNA bzw. der 16S rRNA Genabschnitt eine wesentliche Grundlage

für die Untersuchung von Bakteriengemeinschaften dar. Die Vervielfältigung und

Sequenzierung von Genen ermöglichte es, Bakteriengenome und Phylogenie unabhängig von

Kultivierungserfolgen zu erforschen (Saiki et al, 1988; Sanger et al., 1977). Gängige

Methoden zur Detektion sind z.B. die Denaturierende Gradienten Gelelektrophorese (DGGE;

Muyzer et al., 1993), Restriktionsfragment Längen-Polymorphismus (RFLP; Marsh, 1999),

oder die rDNA Intergenic Spacer Analysis (RISA). Die Quantifizierung von

Bakteriengruppen oder auch –arten kann durch Fluoreszenz in situ Hybridisierung (FISH;

Giovannoni et al, 1988; Amann et al., 1990) bzw. Catalyzed Reporter Deposition-FISH

(CARD-FISH; Pernthaler et al., 2002) sowie mittels Realtime PCR (Heid et al., 1996)

erfolgen. Heute werden Kultivierungsansätze und kultivierungsunabhängige Methoden sowie

Aktivitätsmessungen kombiniert, um möglichst viele Informationen über die Mikrobiologie

eines Habitats zu gewinnen.

Durch den Einsatz dieser Methoden konnte die Struktur der am Stoffumsatz beteiligten

Bakterien, die vorher als „Black Box“ betrachtet wurden, weiter aufgeklärt werden

(Giovannoni & Rappé 2000). So stellte sich heraus, dass insbesondere die gram-negativen

Proteobakterien sowie Bacteroidetes bedeutende Gruppen innerhalb des marinen


6

heterotrophen Bakterioplanktons bilden. Darüber hinaus wurden u.a. methylotrophe

Bakterien, Planctomycetales und die gram-positiven Actinobakterien in marinen Habitaten

nachgewiesen.

Innerhalb der Proteobakterien wurden die gamma-Proteobakterien lange Zeit als die

dominanteste Gruppe des marinen Bakterioplanktons angenommen, da sich Vertreter dieser

Gruppe leicht unter Laborbedingungen isolieren ließen. Über kultivierungsunabhängige

Methoden fand man jedoch heraus, dass die meisten weltweit nachgewiesenen Phylotypen

distinkte Cluster bildeten, die wiederum keine Isolate beinhalteten (Giovannoni und Rappé,

2000). Mittlerweile konnten teilweise auch für diese Cluster mit gezielten

Anreicherungsversuchen einzelne Isolate gewonnen werden (Cho und Giovannoni, 2004), so

dass die Erforschung der ökologischen Funktion dieser Organismen weiter voranschreiten

kann. Physiologisch betrachtet sind gamma-Proteobakterien fakultativ anaerobe und

chemoheterotrophe Organismen, die häufig Oberflächen-assoziiert vorkommen und somit im

Sediment sowie auf Aggregaten eine zentrale Rolle einnehmen.

Die alpha-Proteobakterien sind ebenfalls weltweit verbreitet und zumeist durch die

Subgruppen Sphingomonas und Roseobacter vertreten. Weitere große Bedeutung besitzen die

hoch spezialisierten Cluster SAR 11 (Rappé et al. 2002) und SAR 116 innerhalb der alpha-

Proteobakterien. Die chemoorganotrophen Roseobacter wurden bisher ausschließlich im

marinen Milieu nachgewiesen und stellen dort habitatabhängig bis zu 50% der gesamten

alpha-Proteobakterien. Einige Vertreter gehören zu den aeroben anoxygenen phototrophen

Bakterien und sind somit auch in der Lage, Photosynthese zu betreiben. Aktuell werden große

Forschungsprojekte zur Genomentschlüsselung dieser Organismen1 durchgeführt, die das

Potential dieser hoch interessanten und vielfältigen Gruppe weiter aufklären sollen.

Die aeroben oder fakultativ anaeroben, chemoorganotrophen Bacteroidetes bilden die dritte

große Gruppe innerhalb des marinen Bakterioplankton. Sie sind hoch divers und leben in der

Wassersäule sowohl frei suspendiert als auch Aggregat-assoziiert. Ihre

Stoffwechselphysiologie ist äußerst vielfältig, doch es hat sich gezeigt, dass besonders schwer

abbaubare, hochmolekulare Substanzen bevorzugt von Bacteroidetes abgebaut werden

können, z. B. Chitin oder Cellulose (Cottrell und Kirchman, 2000). Darüber hinaus sind sie

häufig beweglich und können auf Oberflächen gleiten, so dass durch diese Eigenschaften

1 Auch andere Bakteriengruppen, die in marinen Habitaten von Bedeutung sind, werden derzeit durch große Genomprojekte erforscht (z. B. Moran et al., 2004). Die Untersuchung des genetischen Potentials von Organismen führt neben der Entschlüsselung bisher unbekannter Gene auch zur Entdeckung neuer Stoffwechselwege oder biotechnologisch nutzbarer Substanzen (Fusetani, 2000). Man kann daher annehmen, dass die Ozeane ein riesiges Potential bisher unentdeckter Ressourcen bietet, deren Erforschung im Zuge interdisziplinärer Projekte immer mehr in den Vordergrund tritt.


7

angenommen wurde, dass sie besonders auf Aggregaten eine große Bedeutung für

Stoffumsatzprozesse einnehmen.

Diese Zusammenhänge zeigen, dass das Verständnis über die Vorgänge der

Remineralisierung durch die Strukturaufklärung der beteiligten Bakteriengemeinschaften

wesentlich verbessert werden konnte. Darüber hinaus können lokale Gegebenheiten einzelner

Habitate jedoch übergeordnet Einfluss auf die Zusammensetzung der

Bakteriengemeinschaften und somit auch auf die Effektivität des Stoffumsatzes nehmen.


8

I. 3 Geographie und Ökologie der Untersuchungsgebiete

I.3.1 Die Nordsee und die Deutsche Bucht

Die Nordsee liegt auf dem europäischen Kontinentalschelf und wird begrenzt von den

Britischen Inseln und dem Europäischen Kontinent (Abb. 2). Sie ist mit einer

durchschnittlichen Tiefe von 93 m ein flaches Schelfmeer und durch verschiedene

angrenzende Land- und Wasserregionen beeinflusst. Salines Atlantikwasser dringt im Norden

zwischen der schottischen und der norwegischen Küste sowie südlich durch den Ärmelkanal

in die Nordsee. Der größte Eintrag von Süßwasser erfolgt über den Skagerrak aus der Ostsee

und durch verschiedene große Flüsse, die in die Nordsee münden (z. B. von deutscher Seite

die Flüsse Rhein, Ems, Weser, Elbe und Eider). Der mittlere Salzgehalt ist demnach mit

durchschnittlich 15 – 25 Promille an den Küstengebieten geringer als in der offenen Nordsee,

wo durchschnittlich 32 – 35 Promille vorherrschen (Alongi, 1997). Aufgrund der

Amphidromie in der südlichen Nordsee (Defant, 1923) fließt das Wasser in der deutschen

Bucht entgegen dem Uhrzeigersinn, wodurch auch der Transport von partikulären und

gelösten Stoffen sowie von planktischen Organismen beeinflusst wird.

Die deutsche Bucht, der südliche Teil der Nordsee, reicht von Jütland in Dänemark über die

Friesischen Inseln (Nord-, Ost-, und Westfriesische Inseln) bis zur niederländischen Grenze

im Westen. Im Nordwesten wird sie begrenzt von der Doggerbank, einer flachen Sandbank

Abb. 2: Geographische Lage der Nordsee und der Deutschen Bucht sowie des Nationalparks Deutsches Wattenmeer von der ostfriesischen zur nordfriesischen Küste (modifiziert nach http://www.bsh.de)


9

innerhalb der Nordsee, die durch große Fischvorkommen insbesondere für die Fischerei eine

wesentliche Rolle spielt. Das Küstengebiet der Deutschen Bucht bildet das Deutsche

Wattenmeer, eine einzigartige Flachwasserzone, die sich hinter den Friesischen Inseln

erstreckt (vgl. I.3.2).

Grundlage des Nahrungsnetzes in der Nordsee bilden einzellige Algen, das Phytoplankton,

welches im Jahresverlauf mehrere Blüten ausbildet (Alongi, 1997). Im Frühjahr (März bis

April) führen steigende Temperatur- und Lichteinstrahlung sowie hohe

Nährstoffkonzentrationen dazu, dass sich eine Blüte aus Kieselalgen (Diatomeen,

Bacillariophyceae) ausbildet (Drebes, 1974). Durch Nährstofflimitierung und sukzessiven

Fraßdruck durch Zooplankton endet die Blüte zumeist im Juni, bis im Spätsommer eine

zweite, meist weniger intensive Phytoplanktonblüte entsteht (Alongi, 1997). Während der

Blüte scheiden die Diatomeen gelösten organischen Kohlenstoff aus, der von Bakterien

genutzt werden kann (Ittekott et al. 1981). Auch nach dem Zusammenbruch einer

Phytoplanktonblüte profitieren heterotrophe Bakterien vom nährstoffhaltigen Lysat. Dies

kann sich in vermehrter bakterieller Aktivität, Abundanz und Veränderungen in der

Artenzusammensetzung ausdrücken (Reinthaler et al., 2005; Smith et al, 1995, Riemann et al.,

2000, Fandino et al., 2001).

Seit Mitte der 1950er Jahre wurde in der Nordsee ein stetiger Anstieg von Nährstoffen

bedingt durch anthropogene Einflüsse gemessen, der dazu geführt hat, dass die Nordsee stark

eutrophiert ist (Alongi, 1997). Dies führte zu Verschiebungen sowohl in der

Artenzusammensetzung als auch in der Biomasse des Phytoplanktons und resultierte in der

Einschränkung der gesamten Artenvielfalt in der Nordsee. Da die Nährstoffe zumeist über die

Zuflüsse in die Nordsee eingetragen werden, sind erhöhte Konzentrationen von Phosphat,

Nitrat oder auch von Schwermetallen als Gradient von der Küste in die offenen Gewässer zu

beobachten. Darüber hinaus ist durch die globale Erwärmung auch eine Erwärmung der

Wassersäule in der Deutschen Bucht um durchschnittlich 1,1°C beobachtet worden (Wiltshire

and Manly, 2004). Die damit verbundene Verschiebung der Algenblüten könnte durch eine

Veränderung der temperaturabhängigen Rahmenbedingungen entstanden sein. Der Zustand

der Nordsee wird daher seit Jahrzehnten durch verschiedene Institutionen in Monitoringserien

untersucht, um die Nutzung und Belastung der Gewässer zu überwachen und den Lebensraum

zu schützen (Bundesamt für Seeschiffahrt und Hydrographie, BSH; http://www.bsh.de).


10

Untersuchungen zu den Bakteriengemeinschaften in der Nordsee liegen von verschiedenen

Autoren vor. So untersuchten Eilers et al. (2001) die Kultivierbarkeit von Nordseebakterien

bei Helgoland. Es konnten hauptsächlich alpha- und gamma-Proteobakterien sowie

Bacteroidetes isoliert werden. FISH-Zählungen mit spezifischen Sonden ergaben, dass diese

Gruppen ebenfalls einen großen Teil der Bakteriengemeinschaften in der Nordsee darstellen.

Große saisonale Unterschiede in der Hybridisierbarkeit waren erkennbar, die zeigten, dass in

den biologisch hoch produktiven Sommermonaten wesentlich höhere Effizienz erreicht wurde

als in den Wintermonaten. Gerdts et al. (2004) gaben eine Übersicht der Aktivität, Abundanz

und saisonale Veränderungen bakterieller Gemeinschaften, die mit verschiedenen Methoden

als Langzeitmonitoring bei Helgoland durchgeführt wurden. Besonders in den produktiven

Sommermonaten ergaben sich deutliche Änderungen in der Aktivität und Zusammensetzung.

In der südlichen Nordsee wurden saisonal bakterielle Respiration, Artenreichtum

(„Richness“) und Biomasseproduktion entlang von Transekten (Reinthaler et al., 2005) sowie

in Abhängigkeit von Phytoplanktonblüten untersucht (Reinthaler & Herndl., 2005). Es zeigte

sich, dass die bakterielle Biomasseproduktion saisonal stark variiert und korreliert ist mit der

Primärproduktion.

Diese Zusammenhänge verdeutlichen die Notwendigkeit, durch weitere Erforschung der

relevanten Bakterienarten in der Nordsee Schlüsselorganismen zu erkennen und zu

beschreiben. Anhand solcher Indikatororganismen könnten sowohl Änderungen der

ökologischen Gegebenheiten sowie detaillierte Aussagen über Stoffumsatz und äußere

Einflüsse möglich sein. Die Erforschung der Wechselwirkungen zwischen physiko-

chemischen und biologischen Kräften stellt eine essentielle Brücke dar zum Verständnis der

Umwelt und der Bedeutung für das gesamte Ökosystem.


11

I.3.2 Das Wattenmeer

Die südöstliche Nordseeküste besteht aus einem besonderen ökologischen Lebensraum, dem

europäischen Wattenmeer (Abb. 2). Es bildet mit einer Gesamtfläche von ca. 7500 m2 und

einer Gesamtlänge von 500 km die größte zusammenhängende Wattfläche der Welt und reicht

von Den Helder (Niederlande) bis Esbjerg (Dänemark). Es ist gekennzeichnet durch eine hohe

Artenvielfalt und wurde daher 1985 zum Nationalpark erklärt. Der deutsche Teil des

Wattenmeeres wird unterteilt in das Niedersächsische-, das Hamburgische- und das

Schleswig-Holsteinische Wattenmeer. Die Friesischen Inseln sind dem Wattenmeer in

Richtung Nordsee vorgelagert und bilden so eine natürliche Begrenzung.

Als Lebensraum ist das Wattenmeer stark durch die Gezeiten geprägt, wodurch große Teile

des Watts in regelmäßigem Abstand trocken fallen und daher extreme Lebensbedingungen

bieten. Nach Lozan et al. (1994) kann das Watt in vier Ablagerungsbereiche unterteilt werden:

a) Das Sublitoral: Ständig von Salzwasser bedeckte Flächen, z. B. Seegat, Wattrinnen

b) Das Eulitoral: Bereiche, die bei Hochwasser überflutet sind und bei Niedrigwasser

trockenfallen, z. B. Wattflächen zwischen Inseln und Festland

c) Das Supralitoral: Nur bei hochauflaufender Flut von Salzwasser bedeckt, z. B.

Salzmarschen der Inseln und des Festlands

d) Die Dünen: Keine Überspülung mit Salzwasser

Die im Watt lebenden Organismen müssen daher eine hohe Anpassungsfähigkeit besitzen, da

durch die zeitweise Exponierung der Wattfläche und eine insgesamt flache Wassersäule starke

Temperaturschwankungen entstehen. Auch Salinitätsschwankungen sind sehr ausgeprägt,

insbesondere bei starken Regenfällen und an Flußmündungen. Der Austausch der

Wasserkörper zwischen der Nordsee und dem Wattenmeer geschieht über die Seegatten,

Durchlässe zwischen den Inseln, in denen bei jeder Ebbe und Flut sehr hohe

Strömungsgeschwindigkeiten von bis zu 2 m s-1 erreicht werden.

Durch die Eutrophierung der Nordsee insbesondere an den Küsten (vgl. Abschnitt I.3.1) sind

seit den 70er Jahren diverse Projekte zur Beobachtung der Stoffflüsse im Wattenmeer

durchgeführt worden (Baretta and Ruardij, 1988; Cadée, 1984; de Wilde und Beukema,

1984). Um die komplexen Zusammenhänge zwischen der Hydrographie, der Biologie und


12

anthropogenen Einflüssen im Wattenmeer zu studieren, wurden darüber hinaus umfassende

interdisziplinäre Forschungsprojekte ins Leben gerufen. Diese sind zusammenfassend für das

nordfriesische Wattenmeer von Gätje und Reise (1998) sowie für das Spiekerooger

Rückseitenwatt von Dittmann (1999) veröffentlicht worden und bilden die Grundlage für das

Verständnis der Wattenmeerökologie. In beiden Werken wurden Entstehung, Geologie,

Nährstoffkonzentrationen und Stoffflüsse sowie Flora und Fauna untersucht. Eine wesentliche

Rolle nahmen Phyto- und Zooplankton ein; das Bakterioplankton wurde zwar als relevant

erachtet, erschien jedoch als „Black Box“, da nur die Abundanz des gesamten

Bakterioplanktons gemessen wurde. Die Artenzusammensetzung wurde in diesen Arbeiten

nicht berücksichtigt.

Im Rahmen der DFG-geförderten interdisziplinären Forschergruppe „BioGeoChemie des

Watts“, die 2001 ins Leben gerufen wurde, sind weitere Erkenntnisse über die Vorgänge im

Wattenmeer gewonnen worden. Hierzu wurde ein Messpfahl im Spiekerooger Rückseitenwatt

(Standort Otzumer Balje; http://www.icbm.de/watt) errichtet, über den ein fortwährendes

Monitoring der physiko-chemischen Parameter durchgeführt wird. Zusätzlich wurden

konzertierte Meßkampagnen sowie regelmäßige Beprobungen von Tagesgängen

durchgeführt. Durch diese umfangreichen Datensätze manifestierte sich die Bedeutung

biologischer Vorgänge auch im sediment-dominierten Wattenmeer. Besonders zu

Niedrigwasser während des Tages waren biologische Einflüsse erkennbar, sowie saisonal zu

produktiven Jahreszeiten, in denen Phytoplanktonblüten auftraten (Grossart et al., 2004;

Lunau et al., 2006). So wurden im Gegensatz zu den Herbst- und Wintermonaten

beispielsweise im Mai und Juni erhöhte bakterielle Biomasseproduktion und Abundanz

gemessen. Partikelabundanz und –größe verhielt sich gegenläufig, indem in den

Wintermonaten höhere Abundanzen kleinerer Partikel detektiert wurden, im Frühjahr und

Sommer jedoch größere Aggregate in kleinerer Anzahl. Es ist daher anzunehmen, dass der

Einfluß von Phyto- und Bakterioplankton auf Aggregation und Disaggregation durch

Ausscheidung klebriger Substanzen (TEP, transparente Exopolymere; EPS,

Exopolysaccharide; Passow, 2002; Bhaskar et al., 2005) auch in Wattsystemen von

elementarer Bedeutung ist.

Auf tidaler Ebene konnten regelmäßig wiederkehrende Signaturen des suspendierten

partikulären Materials (SPM) nachgewiesen werden, die mit weiteren partikulären Parametern

wie partikulärem organischem Kohlenstoff (POC) und Chlorophyll a korrelierten. Die

Abundanz partikel-assoziierter Bakterien verhielt sich in der Gesamtprobe weitestgehend


13

konstant, wie auch schon in anderen Arbeiten gezeigt (Stevens et al., 2005). Die Auftrennung

in eine absinkende und eine frei schwebende Fraktion zeigte jedoch deutliche Unterschiede in

der Besiedelung (Lunau et al., 2004). Diese Hinweise deuten darauf hin, dass sich die

distinkten Bakteriengruppen unterschiedlich verhalten und Einfluss nehmen.

Erste Untersuchungen zur Zusammensetzung der bakteriellen Gemeinschaft im Watt wurden

1998 durch Llobet-Brossa und Kollegen im Wattenmeersediment durchgeführt. Mittels

Fluoreszenz In Situ Hybridisierung (FISH) wurden sulfatreduzierende Bakterien sowie

Bacteroidetes im Jadebusen nachgewiesen (Llobet-Brossa et al. 1998, 2002). Im Rahmen der

Forschergruppe „BioGeoChemie des Watts“ wurden umfangreiche Untersuchungen auch in

tieferen Sedimentschichten an mehreren Probenahmeorten im Spiekerooger Rückseitenwatt

durchgeführt (Mußmann et al., 2005; Köpke et al., 2005; Willms et al., 2006).

Kultivierungsansätze ergaben eine hohe Artenvielfalt, und die Isolate konnten den

phylogenetischen Gruppen der Proteobakterien, Bacteroidetes, Fusobakterien,

Actinobakterien und Firmicutes zugeordnet werden. Die molekularbiologischen

Untersuchungen ergaben ein ähnliches Spektrum innerhalb der phylogenetischen Gruppen

und die zusätzliche Detektion methanogener Archaeen. Beide Untersuchungen ergaben

Zusammenhänge sowohl zu Aktivitätsmessungen als auch zu sedimentologischen Parametern

und geben somit deutliche Hinweise auf eine ökologische Bedeutung der nachgewiesenen

Stämme und Phylotypen.

Auch in der Wassersäule des Spiekerooger Rückseitenwatts wurden umfassende

Untersuchungen über Bakteriengemeinschaften von Stevens et al. (2005a, b) durchgeführt. In

den Jahren 1999 bis 2000 wurden monatlich Proben genommen, in denen mittels DGGE

nachgewiesen werden konnte, dass hier distinkte Bakteriengruppen existieren: Auf dem

Wattsediment, auf Schwebstoffen, und frei lebend in der Wassersäule. In jedem dieser

Kompartimente waren Phylotypen nachgewiesen worden, die sich ausschließlich in diesem

Lebensraum befanden, sowie Schnittmengen zwischen den einzelnen Gruppen. Vor allem auf

Schwebstoffen bildeten die nachgewiesenen Phylotypen eine Mixtur aus Sediment- und

freilebenden Bakterien. Im Allgemeinen wurden Phylotypen verschiedener Proteobakterien

(alpha-, beta-, gamma- und delta-Proteobakterien), Bacteroidetes und Gram-positive

Bakterien gefunden. Saisonale Veränderungen wurden vorwiegend in den Sommermonaten

während oder nach Phytoplanktonblüten beobachtet. Ein parallel durchgeführter umfassender


14

Kultivierungsansatz ergab nur wenige Übereinstimmungen zu den molekularbiologischen

Ergebnissen.

Die monatliche Probenahme in der Wassersäule war somit geeignet, einen ersten Einblick in

den mikrobiologischen Lebensraum Wattenmeer zu geben, gab jedoch keinen Aufschluss

über die Zeitskala, in der sich Veränderungen der bakteriellen Zusammensetzung ereigneten.

Ebenfalls waren bestimmte Bakteriengruppen unterrepräsentiert, von denen sich in

Kultivierungsansätzen (z. B. Gram-positive Bakterien) sowie anhand von FISH-Zellzahlen

(Bacteroidetes) gezeigt hat, dass sie einen großen Anteil der Bakteriengemeinschaft im

Wattenmeer bilden.

Die weitere Aufklärung der Zusammensetzung der Bakteriengemeinschaften auf Ebene

einzelner, als häufig erkannter phylogenetischer Gruppen ist daher dringend erforderlich.

Ebenso stellt sich die Frage, wie die Bakteriengemeinschaften in kleineren Zeitskalen

beeinflusst werden und inwiefern sich die oszillierenden SPM-Signaturen im Tidenzyklus auf

einzelne Bakteriengruppen auswirken. Es ergaben sich daher folgende Zielsetzungen für die

vorliegende Arbeit:


15

I.4 Zielsetzungen der Arbeit

Ziel der Arbeit war die detaillierte Analyse der frei lebenden und aggregat-assoziierten

Bakteriengemeinschaften im Wattenmeer und der Deutschen Bucht mittels DGGE und FISH

anhand von 16S rRNA und 16S rRNA Genabschnitten. Um auch unterrepräsentierte

Bakteriengruppen erfassen zu können, wurden in allen Arbeiten nicht nur Bacteria-, sondern

auch gruppenspezifische Oligonukleotide (PCR-Primer) verwendet.

Ein Teilaspekt dieser Zielsetzung war die Untersuchung der Zusammenhänge zwischen

Phytoplanktonblüten und Veränderungen in den Bakteriengemeinschaften beider

Kompartimente im ostfriesischen Wattenmeer mit einem engen Probenahmeraster (Kapitel

II).

Einen weiteren Aspekt stellte die Untersuchung tidaler Einflüsse auf die

Bakteriengemeinschaften in Abhängigkeit von saisonalen Aspekten dar (Kapitel III).

Aufgrund der extremen Verhältnisse, die im Wattenmeer herrschen (vgl. Abschnitt I.3.2), war

die Anwendung hoch sensitiver Methoden erforderlich, die zunächst etabliert und getestet

werden mussten. So wurde die DGGE nicht nur DNA, sondern auch RNA basiert

durchgeführt; neben der FISH wurde für einen Vergleich ebenfalls die CARD-FISH Methode

angewendet.

Anhaltspunkte über außergewöhnlich hohe Abundanzen von Gram-positiven Bakterien im

Wattenmeer (Stevens, 2004) erforderten molekularbiologische Untersuchungen, um die

Relevanz dieser Aussage zu untermauern (Kapitel IV). Mit Hilfe einer spezifischen DGGE

sowie CARD-FISH Untersuchungen konnten zusätzliche Hinweise über Zusammensetzung

und Abundanz von Gram-positiven Bakterien an verschiedenen Standorten im Wattenmeer

nachgewiesen werden.

Eine weitere Zielsetzung stellte die Untersuchung von Bakteriengemeinschaften im Watten-

meer und der Deutschen Bucht dar, um den Austausch von Wassermassen und die räumliche

Verteilung der dominierenden Bakteriengruppen darzustellen (Kapitel V). Hierzu wurden

zwei Messkampagnen im Sommer 2002 und 2003 mit umfassenden Probenahmen

durchgeführt, um lokale Gegebenheiten und ihren Einfluss auf die Bakteriengemeinschaften

zu erfassen.


16

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Alldredge, A. L., and M. W. Silver. 1988. Characteristics, dynamics, and significance of marine snow. Prog Oceanogr 20:41-82

Amann, R. I., L. Krumholz, and D. A. Stahl. 1990. Fluorescent-oligonucleotide probing of whole cells for determinative, phylogenetic, and environmental studies in microbiology. J Bacteriol 172: 762-770

Azam, F., T. Fenchel, J. G. Field, J. S. Gray, L. A. Meyer-Reil, and T. F. Thingstad. 1983. The ecological role of water-column microbes in the sea. Mar Ecol Prog Ser 10:257-263

Azam, F., and B. C. Cho. 1987. Bacterial utilization of organic matter in the sea. In Ecology of Microbial Communities, Cambridge University Press, 261-281

Baretta, J., and P. Ruardij. 1988. Tial flat estuaries. Berlin: Springer-Verlag.

Bhaskar, P., H. P. Grossart, N. Bhosle, and M. Simon. 2005. Production of macroaggregates from dissolved exopolymeric substances (EPS) of bacterial and diatom origin. FEMS Microbiol Ecol 53: 255-264

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II.

Effects of a phytoplankton bloom in a coastal ecosystem

on the composition of bacterial communities

Kapitel II Effects of a phytoplankton bloom on bacterial communities

21

Effects of a phytoplankton bloom in a coastal ecosystem on the composition

of bacterial communities

Beate Rink 1, Susanne Seeberger 1, Torben Martens 1, Claus-Dieter Duerselen 2,

Meinhard Simon 1, Thorsten Brinkhoff 1*

1 Institute for Chemistry and Biology of the Marine Environment (ICBM), University of

Oldenburg, P.O. Box 2503, D-26111 Oldenburg, Germany

2 AquaEcology, Marie-Curie-Str. 1, D-26129 Oldenburg, Germany

*Corresponding author. E-mail: [email protected]

Running head: Effects of a phytoplankton bloom on bacterial communities

KEY WORDS: Free-living and particle-attached bacteria, Bacteroidetes, Roseobacter,

phytoplankton, DGGE


22

ABSTRACT: We studied the composition of free-living and aggregate-associated bacterial

communities during the course of the phytoplankton succession in spring and early summer in

the German Wadden Sea, a tidal flat ecosystem in the southern North Sea. We applied the

DGGE approach based on PCR amplified 16S rRNA gene fragments, and, in addition to

Bacteria-specific primers, used primers specific for alpha-Proteobacteria, the Roseobacter

clade, and the Bacteroidetes phylum. Even though the application of Bacteria- and alpha-

Proteobacteria-specific primers detected some changes, changes were most pronounced with

the Roseobacter- and Bacteroidetes-specific primer sets. They were supported by a

correspondence analysis, which showed a highly significant correlation of the DGGE banding

patterns of the Roseobacter specific PCR with the composition of the phytoplankton. This

indicates that changes of the phytoplankton composition in this habitat are not reflected by the

patterns of the most abundant or most readily amplifiable phylotypes. The findings rather

suggest that few, specialized heterotrophic bacteria are most responsive to the organic matter

supplied by senescent phytoplankton and that the main part of organic matter in the German

Wadden Sea is utilized by generalists. Sequence analyses of excised bands revealed a high

diversity for the Bacteria- and Bacteroidetes-specific approaches. The bacterial community

detected by the alpha-Proteobacteria-specific primer set, however, was mainly composed of

bacteria affiliated to the Roseobacter clade.


23

INTRODUCTION

Today it is well established that heterotrophic bacteria are an important component of and

key players in the biogeochemical cycling of elements and the flux of energy in aquatic

ecosystems. Depending on the ecosystem and on various environmental and biotic factors the

composition of the bacterial communities involved may exhibit distinct differences and

variations in time and space. Temperature preferences certainly select for certain bacterial

taxa but little direct information is available within this context. The most important factor for

selecting specific bacterial groups is supply by specific monomeric and polymeric

components of the dissolved organic matter (DOM) pool and of inorganic nutrients such as

phosphate, ammonium or nitrate. It has been shown that alpha-Proteobacteria prefer

monomers such as amino acids and N-acetyl-glucosamine, whereas Cytophaga/Flavobacteria

(now Sphingobacteria/Flavobacteria) of the Bacteroidetes phylum prefer polymers such as

chitin and protein, and gamma-Proteobacteria amino acids and proteins (Cottrell & Kirchman

2000). Various mesocosm studies have shown that distinct DOM components via direct

supply or the experimental induction of phytoplankton blooms select for specific bacterial

subcommunities or populations (LeBaron et al. 1999, Pinhassi et al. 2004, Riemann et al.

2000, Schäfer et al. 2001). The specific organic matter profile of various algae appears also to

be an important selection factor for distinct bacterial communities and populations evolving in

the phycosphere of algae (Grossart 1999, Grossart et al. 2005, Schäfer et al. 2002). In fact,

alpha-Proteobacteria, in particular the Roseobacter clade, and the Bacteroidetes appear to be

most responsive to inputs of phytoplankton-born DOM (Fandino et al. 2001, Grossart et al.

2005, Pinhassi et al. 2004, Riemann et al. 2000, Schäfer et al. 2001).

It is also well established that the community composition of particle-associated (PA)

bacteria differs from that of free-living (FL) bacteria. Several studies have shown that

Sphingobacteria and Flavobacteria preferentially colonize particles whereas alpha- and

gamma-Proteobacteria mainly dwell in free-living marine bacterial communities (Fandino et

al. 2001, Grossart et al. 2005, Simon et al. 2002). Our knowledge on the development and

succession of specific subcommunities and populations within PA bacterial communities

during phytoplankton blooms, however, is still fragmentary.

Experimental studies are important to elucidate single factors affecting the composition

of bacterial communities. As the aim of such studies is to better understand how the

composition of bacterial communities is controlled at ambient, but much more complex

conditions it is important to complement these studies by appropriate field observations. Such


24

studies have been carried out in various ecosystems and shown that the composition of

bacterial communities undergoes temporal changes during phytoplankton blooms (Fandino et

al. 2001, Larsen et al. 2004, Yager et al. 2001). These changes often reflect the changing

environmental conditions and DOM supply and also indicate which bacteria are mainly

involved in the biogeochemical cycling of elements and flux of energy. Denaturing gradient

gel electrophoresis (DGGE) of PCR-amplified 16S rRNA gene fragments using Bacteria-

specific primers (Muyzer et al. 1993) has been proven to be a powerful tool to assess the

composition and temporal changes of bacterial communities. Using Bacteria-specific primers

for this approach appears to be selective against the Bacteroidetes group (Cottrell &

Kirchman 2000, Selje et al. 2005, but see Castle & Kirchman 2004). Therefore, and to obtain

a more detailed insight into the composition of bacterial communities and their major players,

it is desirable to apply primers targeting specifically important groups such as Bacteroidetes

and alpha-Proteobacteria.

The aim of our study was to investigate the composition of free-living and aggregate-

associated bacterial communities during the course of the phytoplankton succession in spring

and early summer in the Wadden Sea, a tidal flat ecosystem of the southern North Sea. Based

on previous studies, we hypothesized that the expected bacterial response to the

phytoplankton succession would be reflected most pronounced by alpha-Proteobacteria and

the Sphingobacteria/Flavobacteria group. Therefore, we applied the DGGE approach and, in

addition to Bacteria-specific primers, primers specific for alpha-Proteobacteria, the

Roseobacter clade, and the Bacteroidetes group.

MATERIALS AND METHODS

Sample collection and processing. Surface water samples were collected weekly by

bucket from shipboard at high tide from 12 April to 29 June 2000 in the Backbarrier tidal flat

ecosystem of the German Wadden Sea near Spiekeroog Island (53° 44.4 N, 7° 41 E). This is a

mesotidal ecosystem characterized by high loads of suspended particulate matter (SPM). For

further details see Stevens et al. (2005a) and Lunau et al. (2006). For analysis of SPM and the

particulate carbon fractions 0.5-1 L of seawater was filtered onto pre-combusted (2 h at

550°C) and pre-weighed glass fiber filters (GF/F, Whatman, USA) and stored at –20°C in the

dark until further processing. For DGGE analysis, 250 ml of seawater were pre-filtered onto

5.0 µm polycarbonate-filters (Nuclepore) to obtain the fraction of aggregate-associated and


25

subsequently onto 0.2 µm polycarbonate-filters to obtain that of free-living bacteria. Filters

were stored at –20°C in the dark until further processing. For enumeration of bacterial and

phytoplankton cells 100 ml of water sample were fixed with formaldehyde (final

concentration 2% vol/vol) or Lugol and stored at 4°C. Hydrographic data (temperature,

salinity, pH, and oxygen) were measured by probes (LF 196, pH192, OXI 196, WTW,

Weilheim, Germany).

SPM dry weight, particulate carbon fractions. Filters were dried for 1 hour at 110°C

and weighed on a micro-balance (Sartorius, Germany). Total particulate carbon (TC) and

particulate inorganic carbon (PIC) were determined after high temperature combustion and

titration of the CO2 produced against Ba(ClO4)2. Particulate organic carbon (POC) was

calculated as the difference of TC and PIC. For further details see Stevens et al. (2005a).

Bacterial and algal cell counts. Abundance of free-living and aggregate-associated

bacteria was enumerated after DAPI (4´-6-diamidino-2-phenylindole) staining by

epifluorescence microscopy at 1000x magnification according to Crump et al. (1998). To

distinguish particle-attached and free-living bacteria, seawater was fractionated by filtration

onto 5.0 µm and subsequently onto 0.2 µm polycarbonate-filters. To reduce the background

fluorescence by inorganic matter filters were counter-stained with an acridine orange solution

(0.1%). Lugol-fixed phytoplankton samples were enumerated by inverted microscopy.

Phytoplankton was identified on the species level when possible. For estimating

phytoplankton biomass cell numbers were multiplied by cell carbon. The latter was estimated

from measured cell sizes of individual cells converted to carbon according to empirical

carbon/cell volume conversion factors from the Biologische Anstalt Helgoland (J. Berg,

unpubl. results).

Nucleic acid extraction. The isolation of genomic DNA was performed by phenol-

chloroform extraction after bead beating as described earlier with slight modifications (Selje

& Simon, 2003). The precipitation was done overnight at –20°C using isopropanol. The DNA

was resuspended in molecular grade water (Eppendorf, Germany) and stored at –20°C until

further processing.

Primer sets. PCR amplification of 16S rRNA gene fragments was performed with primer

pairs specific for Bacteria (GC 341F and 907RM), the Bacteroidetes phylum (GC-CF319f

and 907RM), alpha-Proteobacteria (GC 341F and ALF968r), and the Roseobacter clade

within alpha-Proteobacteria (GC ROSEO536Rf and GRb735r). Primer sequences and

references are given in Table 1. ’GC’ indicates that a GC clamp was added to the primer

(Muyzer et al. 1993). For the primer GC ROSEO536Rf the following GC clamp was used: 5’-


26

CGCCCGCCGCGCCCCGCGCCCGTCCCGCCGCCCCCGCCCG-3’. For the sequences of

the other GC clamps used in this study see the references cited in Table 1. Specificity of the

primers used for Bacteroidetes was described earlier (Jaspers et al. 2001, Kirchman 2002).

The oligonucleotide probe ALF968r (Neef, 1997), used as reverse primer for alpha-

Proteobacteria, was tested theoretically using the BLAST function of the NCBI server

(http://www.ncbi.nlm.nih.gov). Search results for this primer sequence revealed up to 10%

matches to other phylogenetic groups with 100% sequence similarity for the first one hundred

matches. The primer set used for the Roseobacter-group was tested theoretically with the

whole database of the ARB software package (Ludwig et al. 2004) and recently published

sequences present in GenBank (www.ncbi.nlm.nih.gov) of cultivated and uncultivated

organisms affiliated with the Roseobacter clade. In total 183 sequences affiliated with this

group were considered. Specificity was also tested in PCR assays using several described

species as positive and negative controls (Table 2), and 25 isolates affiliated with the

Roseobacter group, taken from our culture collection.

PCR amplification of 16S rRNA gene fragments. PCR amplifications were performed

with an Eppendorf Mastercycler (Eppendorf, Hamburg, Germany) as follows: One µl of

template was added to 49 µl of PCR mixture containing 1 U of Sigma RedTaqTM polymerase

and 5 µl 10 x RedTaqTM PCR buffer (Sigma, Deisenhofen, Germany), bovine serum albumin

(10 mg ml-1), 250 µM of each deoxynucleotide triphosphate, 2.1 µM MgCl2, and 20 pmol of

each primer. The PCR protocol for the Bacteria-specific primer set was performed as

described previously (Brinkhoff & Muyzer, 1997). Amplification of the 16S rRNA gene

fragments of alpha-Proteobacteria was performed under the same conditions with an

annealing temperature of 65°C for 10 cycles and subsequently 55°C for 20 cycles.

Roseobacter-specific PCR conditions were 5 cycles at 65°C and 25 cycles with an annealing

temperature of 63°C. For highest specificity, a maximum of 30 cycles is recommendable at

this step. PCR with the primer set specific for Bacteroidetes was performed as described

previously (Jaspers et al. 2001). Four µl of the amplification products were analyzed by

electrophoresis in 2% (w/v) agarose gels and stained with ethidium bromide (1 µg ml-1)

(Sambrook et al. 1989). For subsequent sequencing analysis PCR products were purified by

using the Qiaquick PCR purification kit (Qiagen Inc., Chatsworth, Calif.).

DGGE analysis of PCR products. DGGE was performed with the D-Code system (Bio-

Rad Laboratories, Inc.). For gene fragments of Bacteria and alpha-Proteobacteria, the

protocol described by Brinkhoff & Muyzer (1997) was used. For 16S rRNA gene fragments

obtained with the primer pair GC-CF319f and 907 RM the gradient was modified to 15 to


27

85% denaturant. DGGE analysis of Roseobacter 16S rRNA gene fragments was performed

with 20 to 70% denaturant and 9% (wt/vol) polyacrylamide content. After electrophoresis, the

gels were stained with SYBR Gold (Molecular Probes, Inc.) and photographed using a

BioDoc Analyze Transilluminator (Biometra, Göttingen, Germany). Bands were excised with

a scalpel sterilized with ethanol and transferred to sterile Eppendorf caps. Fifty µl of water

(molecular grade, Eppendorf, Germany) were added and the samples were stored at –20°C.

Cloning. Twenty four DGGE bands (GWS-e1-FL to GWS-e13-PA, GWS-c3-FL, GWS-

c16-PA, GWS-c9-PA, GWS-c10-PA, GWS-c18-PA and GWS-a10-PA to GWS-a13-PA,

GWS-a4-FL, GWS-a8-FL) were cloned using the pGEM®-T Vector System II (Promega,

Madison, USA) following the instruction manual. Clones with inserts were picked,

resuspended in molecular grade water (Eppendorf, Germany) and screened by DGGE to

check if the insert position matches the position of the corresponding DGGE band. Adequate

clones were amplified and subsequently sequenced using the primers pUC/M13f and

pUC/M13r (Messing, 1983) with an annealing temperature of 48°C.

Sequencing and phylogenetic analysis. PCR products were sequenced using the

DYEnamic Direct cycle sequencing kit (Amersham Life Science, Inc.) and a Model 4200

Automated DNA Sequencer (LI-COR, Inc.). Sequencing primers were 341F and 907RM for

direct sequencing of DGGE bands, or M13 primers as described above for cloned bands

labeled with IRDyeTM800. For all sequences, at least 400 bp were determined. Phylogenetic

affiliation of the sequences was compared to those in GenBank using the BLAST function of

the NCBI server (http://www.ncbi.nlm.nih.gov/BLAST/). Phylogenetic trees were constructed

using the ARB software package (Ludwig et al. 2004, http://www.arb-home.de). The

backbone tree was calculated with the maximum likelihood method using sequences with a

minimum length of 1300 bp including type strains of the selected phylogenetic groups. For

tree calculation, positions were excluded at which less than 50% of all sequences showed the

same residues to avoid uncertain alignments. Sequences with less than 1300 bp were added to

the backbone tree with the maximum parsimony method using the same filter. As an

outgroup, 16S rRNA gene sequences of seven type strains belonging to Cyanobacteria were

used.

The sequences obtained in this study are available from GenBank under accession no.

DQ080919 to DQ080962.

Statistics. Cluster analyses of DGGE banding patterns were performed using Gel

Compar II, version 2.5 (Applied maths, Kortrijk, Belgium). Calculations were done curve

based using Pearson correlation and UPGMA. A correspondence analysis of the DGGE


28

banding patterns and the phytoplankton composition was performed using ADE-4

(Thioulouse et al. 1997). To analyze the bacterial community structure, we exported the raw

data of the cluster analysis and generated a matrix based on the specific band heights. For

phytoplankton, we used relative species abundance. A modified correspondence analysis was

performed row weighted on a biplot scale. After calculation of the COA for each community

a Coinertia analysis was performed to connect the data. A permutation test based on the

Monte Carlo method was calculated using the Coinertia test (– Fixed D; number of random

matching: 1000).

RESULTS

Environmental conditions and SPM properties

From the start of the study period in mid-April until 10 May 2000 the water temperature

continuously increased from 8-17 °C (Fig. 1A). Thereafter it fluctuated between 17 and 13

°C. Salinity ranged between 29 and 32‰ (Fig. 1A) and SPM dry weight from 80-120 mg l-1

in April and May, but increased to 160 mg L-1 on 14 June (data not shown). POC

concentrations varied between 0.8 mg L-1 on 3 May and 4.7 mg L-1 on 26 April (Fig. 1B).

They steadily increased from 3 to 17 May and from 24 May to 14 June.

Phytoplankton and bacterial dynamics

The phytoplankton consisted exclusively of diatoms and few dinoflagellates. From 12

April to 3 May diatom cell numbers strongly decreased from 6.5x103-1.2x103 L-1 but

thereafter continuously increased until 24 May (Fig. 1C). After the decline of this bloom in

late May only low numbers remained. Whereas the initial bloom on 12 April exhibited a high

diversity and evenness, the bloom in May became more and more dominated by Guinardia

delicatula, constituting 70% of algal cell numbers and biomass on 24 May (Fig. 1C). One

week later, when diatom cell numbers declined to ~30% of the previous week, abundance of

Guinardia delicatula had strongly decreased while Pseudonitzschia pungens constituted 50%

of the cell numbers. At the onset of the study phytoplankton constituted 50% of POC, but on

26 April only 4%. Thereafter during the Guinardia bloom, phytoplankton carbon

continuously increased to 13% on 24 May (Fig. 1B).


29

Cell numbers of FL bacteria increased from 12 April until 3 May from 2.4x106-4.0x106

ml-1 and fluctuated thereafter around this value (Fig. 1D). Cell numbers of PA bacteria were

lower and ranged from 0.5-1.1x106 ml-1, accounting for 11 to 22% of total bacterial numbers.

Specificity of the Roseobacter primer set

Comparison of 16S rRNA gene fragments present in our ARB database revealed that the

forward primer GC ROSEO536Rf matched 131 of a total of 183 target sequences affiliated to

the Roseobacter clade. Forty three Roseobacter sequences had no or incomplete information

at the target site of the primer, and 8 sequences of uncultured Roseobacter-affiliated

organisms showed up to three mismatches to the primer sequence. Sulfitobacter pontiacus

(Acc. no. Y13155) had one mismatch at position 17 of the primer sequence. Reverse primer

GRb735r targeted 133 sequences after insertion of a wobble (G/A) at Escherichia coli

position 752. Forty four Roseobacter sequences had no or incomplete information at the target

site of the 16S rRNA gene. Six sequences of target bacteria had up to three mismatches:

Roseobacter sp. J8W (AF026462; two mismatches), Roseobacter sp. J2W (AF026462, three

mismatches), Roseobacter sp. KT11117 (AF173971, one mismatch), Adriatic 72 (AF030780,

one mismatch, two non-defined bases), Sulfitobacter pontiacus (Y13155, one mismatch) and

GWS-BW-H66M (AY515422, one mismatch). The non-target sequences of Rhodovulum

iodosum and clone SAR102 (Acc. no. L35460) had no mismatches to the primer sequence.

Considering all respective sequences in the ARB data base, the use of both primers paired

resulted in at least one mismatch to all other phylogenetic groups.

PCR results showed that the specificity and sensitivity of the Roseobacter primer set was

very high. With one step down from 65°C -63°C and 1 U of Taq polymerase, 0.2 ng genomic

DNA µl-1 of Roseobacter gallaeciensis was detectable. DNA of the non-target organism

Paracoccus aminophilus (1 mismatch to the target sequence) was detected down to 2 ng µl-1.

Specificity was higher but less sensitive under the same conditions with 0.5 U of polymerase,

detecting 0.1-1 ng DNA µl-1 of R. gallaeciensis and 20 ng DNA µl-1 of P. aminophilus. To

determine a possible sequence preference of the primer set, a DNA mixture of both organisms

with equal DNA amounts was amplified and the PCR products were analyzed using DGGE.

By this approach only amplicons of R. gallaeciensis could be detected (data not shown). This

notion suggests that the amplification of non-target organisms is suppressed under the chosen

PCR and DGGE conditions.


30

DGGE banding patterns

The DGGE analyses with the various primer sets showed distinctly different banding

patterns of the FL as well as PA bacterial communities (Fig. 2A-D). The Bacteria-specific

primer set yielded 12 to 15 bands per lane in the PA bacterial fraction and 12 to 18 bands in

the FL bacterial fraction (Fig. 2A). Changes in the banding patterns occurred mainly during

the Guinardia bloom in May, showing a slight increase of band numbers in the FL fraction on

17 May and the appearance of a strong band in the PA fraction (GWS-e11-PA). The cluster

analysis yielded distinct clusters for the FL and PA bacterial communities (Fig. 3A). Only

during the Guinardia bloom between 10 and 24 May the PA bacteria clustered separately and

closer with the FL bacterial community. Correspondence analysis did not yield a significant

correlation with the phytoplankton composition. The aggregate sample of 31 May was

reamplified from a former PCR product and showed reduced band numbers compared to the

other samples. Hence, the fingerprint of this sample appeared as an outgroup in the cluster

analysis and was not regarded for the further discussion.

The Bacteroidetes–specific banding patterns revealed 7-12 and 9-18 amplicons per lane

in the FL and PA bacterial fractions, respectively (Fig. 2B). Low numbers of 7-9 bands

occurred in the FL bacterial fraction before and after the Guinardia bloom and higher

numbers of 10-12 bands during the bloom. In contrast, the number of bands in the PA

bacterial community was high before and after and decreased during the bloom. The cluster

analysis showed a distinct cluster of the FL bacterial community, excluding the dates towards

the end of the Guinardia bloom, when banding patterns clustered together with those of the

PA bacterial community during the bloom (Fig. 3B). Furthermore, the latter fraction exhibited

clearly different patterns before and after the bloom. A correspondence analysis revealed a

moderate correlation of the banding patterns with the composition of the phytoplankton

(P=0.067).

DGGE banding patterns specific for alpha-Proteobacteria showed 7-8 and 8-13 bands per

lane in the FL and PA bacterial communities, respectively (Fig. 2C). Most of the bands were

permanently present but a few bands in both fractions occurred in the course of the bloom

(GWS-a11-PA, GWS-a6-FL, and GWS-a5-FL). The cluster analysis showed generally rather

complex patterns and that PA bacteria during the Guinardia bloom clustered together with FL

bacteria (Fig. 3C). The correspondence analysis did not yield a significant correlation of the

banding patterns with the phytoplankton composition.


31

The Roseobacter-specific DGGE banding patterns showed 5-8 bands per lane in the FL

bacterial fraction and 8-18 bands in the PA bacterial fraction (Fig. 2D). Quite a few bands

were permanently present in both fractions, but additional bands occurred during the decline

of the bloom in April and the Guinardia bloom in May, mainly in the PA bacterial fraction.

The cluster analysis yielded complex patterns with several subclusters both of FL and PA

associated bacterial fractions. A distinct subcluster comprised the banding patterns of both

fractions during the Guinardia bloom (Fig. 3D). The correspondence analysis showed a

highly significant correlation of the banding patterns with the composition of the

phytoplankton (P=0.03).

Phylogenetic affiliation

The sequence analysis of excised bands revealed a high diversity of the obtained

phylotypes for the 16S rRNA gene fragments of the Bacteria- and Bacteroidetes-specific

approaches (sequences obtained with Bacteria-specific primers were designated GWS-e and

sequences obtained with Bacteroidetes-specific primers as GWS-c; Figs. 4A, B). The

bacterial community detected by the alpha-Proteobacteria-specific primer set (sequences

GWS-a) was mainly composed of bacteria belonging to the Roseobacter clade. Most

phylotypes of this group, detected by the Bacteria- and alpha-Proteobacteria-specific primer

set, clustered within the recently described WAC I cluster (Stevens et al. 2005b) or RCA

cluster (Selje et al. 2004). The primer set used for alpha-Proteobacteria turned out to be not

specific, as sequencing results revealed that two sequences affiliated to delta-Proteobacteria

(GWS-a12-PA, GWS-a13-PA) and one to Bacteroidetes (GWS-a8-FL). In contrast, although

the primer GC-CF319f used for specific amplification of 16S rRNA gene sequences of

bacteria belonging to Bacteroidetes is known to be unspecific (Kirchman et al. 2003), all our

phylotypes of the sequenced bands fell into this phylum.

During the Guinardia bloom, DGGE derived phylotypes belonging to the WAC I cluster

dominated the FL bacterial fraction. DGGE band GWS-e7-FL was present during the bloom.

This phylotype was closely related to GWS-a6-FL and GWS-a5-FL (sequence differences

<0.8%, Fig. 4A) which were also present only during the Guinardia bloom. While these

organisms seem to be highly responsive to the phytoplankton composition, other members of

the WAC I cluster were present during the whole investigation period, e.g. GWS-e6-FL (Fig.

4A). This phylotype is closely related to DGGE band GWS-FL-3, which was persistently

detected throughout the year in the Wadden Sea, indicating that this organism is well adapted

to highly variable biotic and environmental conditions in this habitat (Stevens et al. 2005a). In


32

the PA bacterial fraction chloroplast DNA (GWS-e11-PA) represented the most significant

change within the community detected by the Bacteria-specific primer set. Sequencing of

other conspicuous bands was not possible, as the diffuse bands in the upper part of the gel

could not be reamplified.

One of the Bacteroidetes–specific phylotypes appeared during 24 May to 14 June at the

end of the Guinardia bloom both in the FL and PA bacterial fractions (GWS-c6-FL, GWS-c5-

FL and GWS-c15-PA). This phylotype is closely related to GWS-e9-FL in the FL bacterial

fraction (Fig. 4B). BLAST results revealed that the closest related sequence of these bands is

DGGE band GWS-AG-8, which was detected on aggregates in June 2000 in the same area

(Stevens et al. 2005a). Other phylotypes affiliated to the Bacteroidetes were present during

the whole investigation period. DGGE band GWS-c8-PA was detected in the PA bacterial

fraction from April to June and is closely related to strain T15 (AY177723, 99% similarity,

502/505 bp), isolated from the same habitat in October 1999 (Brinkhoff et al. 2004). The

phylotype was also detected in a seasonal sampling campaign in this habitat from 2000 to

2002 using GC-CF319f and 907RM for DGGE analysis (S. Seeberger, unpublished results).

DISCUSSION

Our results indicate that the composition of the bacterial communities in the Wadden Sea

underwent changes during the phytoplankton succession in spring and early summer. These

changes were detected as the disappearance of DGGE bands and the appearance of new ones,

were most pronounced during the Guinardia bloom and its decline in May, and occurred in

the FL as well as in the PA bacterial communities. Even though the application of Bacteria-

and alpha-Proteobacteria-specific primers in the DGGE approach detected some of these

changes, they were detected most clearly with the Roseobacter- and Bacteroidetes-specific

primer sets and supported by a correspondence analysis. Whereas the number of bands of the

FL bacterial fraction within the Bacteroidetes increased during the Guinardia bloom it

decreased in the PA bacterial fraction. Within the Roseobacter clade, the number of bands of

the PA bacterial fraction increased during the decline of both blooms, in late April and late

May. Hence our results show that the bacterial communities respond to the changing

phytoplankton community and organic matter field on a rather specific phylogenetic level and

call for applying class- and subclass-specific primer sets in the DGGE approach for such

investigations.


33

Our investigation complements mesocosm experiments which obtained similar findings

(LeBaron et al. 1999, Pinhassi et al. 2004, Riemann et al. 2000, Schäfer et al. 2001) and

demonstrate that changes in FL as well as PA bacterial communities during the development

of phytoplankton blooms also occur and can be detected at ambient conditions in a natural

ecosystem. Our results, however, go beyond the mentioned experimental studies by showing

in a much more detailed way the different responses of bacteria affiliated to the Roseobacter

clade and the Bacteroidetes.

To achieve these results we applied sets of published primers for all bacterial target

groups and modified specific probes for the Roseobacter clade from earlier studies to

optimize its specificity (Table 1). In addition, we developed a PCR to achieve highest

specifity for this primer set. As expected from the BLAST search, the results revealed that the

alpha-Proteobacteria-specific primer set was not specific. We detected two sequences

affiliated to delta-Proteobacteria and one to the Bacteroidetes phylum, indicating that

sequencing of bands is essential when applying this primer set. However, the great majority of

the bands sequenced affiliated to alpha-Proteobacteria and exclusively to the Roseobacter

clade, underscoring the significance to apply a primer set specific for this clade. In this study,

the Bacteroidetes-specific primer set, in fact, was specific, as all sequenced bands affiliated to

the respective target group. The fact that most changes within the bacterial communities were

only detected by applying primer sets for a lower phylogenetic level indicates that these

changes do not affect the most abundant or most readily amplifiable phylotypes.

Although various prominent bands were visible using the group-specific primer set, only

two Bacteroidetes affiliated phylotypes could be detected with the Bacteria-specific primer

set. Only one of these two phylotypes was also detected with the group-specific primer set

(GWS-e9-FL, Fig. 4B) suggesting that the Bacteria-specific primer set discriminates the

Bacteroidetes affiliated bacteria, as has been reported previously (Cottrell & Kirchman,

2000). In contrast, we had no indication of a biased amplification of phylotypes affiliated to

alpha-Proteobacteria by the Bacteria-specific primer set. Seven of the 17 sequences of this

subclass were amplified by the Bacteria-specific primer set and all except one sequence

(GWS-e10-FL) were very closely related or similar to those amplified by the alpha-

Proteobacteria-specific primer set (Fig. 4A). However, the latter and the Roseobacter-

specific primer set yielded a much better resolution and detected substantially more

phylotypes with a presumably lower abundance.

The decreasing number of bands of Bacteroidetes in the PA bacterial fraction detected

during the Guinardia bloom and its decline indicates the formation of a more specialized


34

environment on suspended particles and aggregates to which fewer bacteria of this

phylogenetic group were able to adapt. The number of bands of the Roseobacter clade,

however, remained rather unchanged and even increased at the decline of the bloom on 31

May, indicating that the niche diversity for this bacterial group did not decrease.

In contrast to aggregates, the number of bands of FL bacterial phylotypes belonging to

the Bacteroidetes increased during the bloom, indicating that the DOM supply became more

diverse, presumably including a variety of polymers released from growing and decaying

diatoms and solubilizing phytodetrital aggregates. Two of the newly occurring phylotypes

cluster together (GWS-c6-FL, GWS-c5-FL) and also together with other phylotypes which

occurred on 14 June (GWS-e9-FL) in the FL bacterial fraction and in the PA bacterial fraction

(GWS-c15-PA, Fig. 4B). These phylotypes are closely related to phylotypes which were

retrieved from the associated bacterial communities of two diatoms [SB-42-DB (Schäfer et al.

2001), Flo-21 (Grossart et al. 2005)], suggesting that they are particularly adapted to the

organic matter profile of diatoms.

Our results show that organisms of the Roseobacter clade and the Bacteroidetes are most

responsive to the changing organic matter field during the phytoplankton blooms. This is in

line with other studies (Dang & Lovell, 2002, Lebaron et al. 1999, Riemann et al. 2000,

Fandino et al. 2001, Pinhassi et al. 2004, Grossart et al. 2005) and thus indicates that members

of these two bacterial groups appear to be particularly adapted to such conditions, at least in

temperate waters. The significance of the Bacteroidetes affiliated bacteria in consuming

complex and polymeric DOM in marine systems is well known and fairly well understood,

mainly because of their specific properties to hydrolyze polymers (Cottrell & Kirchman,

2000; Kirchman, 2002). The significance of the Roseobacter clade is much less understood.

Some members of this clade exhibit aerobic anoxygenic photosynthesis but the significance of

this metabolic pathway at ambient conditions and varying trophic state is still unclear

(Schwalbach & Fuhrman, 2005). Other members of this clade appear to be involved in the

decomposition of DMS (Moran et al. 2003). Roseobacter strains have been isolated from FL

as well as PA bacterial communities and quite a few of them exhibit antibiotic and quorum

sensing properties (Long & Azam 2001, Gram et al. 2002; Grossart et al. 2004). Hence, it

appears conceivable that varying adaptive properties make this clade well suitable to dwell

successfully in marine systems. More work, however, is needed to better understand the

success of this clade at a physiological and genetic level.

The combined application of Bacteria- and group-specific primer sets revealed that a

hierarchical structure exists in the bacterial communities, both in the FL as well as the PA


35

fractions. The Bacteria-specific primer set detected mainly those phylotypes which constitute

the main and often dominant components of the bacterial communities, persisting throughout

most of the time and thus comprising bacteria able to adapt to rather variable environmental

conditions and exhibiting a rather generalistic life style. These phylotypes include members of

the RCA-cluster (Selje et al. 2004), and the WAC I cluster (Stevens et al. 2005b) of the

Roseobacter clade. In contrast, the group- and clade-specific primer sets detect, besides some

of these generalistic phylotypes, others which are probably less abundant but appear at

distinct environmental and biotic conditions, such as during certain periods of phytoplankton

blooms. The phylotypes detected by these primer sets reflect in a much more sensitive way

these changing conditions and thus allow a more detailed analysis of bacterial communities at

varying environmental conditions. The application of Bacteria-specific primer sets appears to

be appropriate to study the main components of bacterial communities and their variability at

greatly varying environmental conditions such as in salinity gradients (Selje & Simon, 2003;

Troussellier et al. 2002), PA vs. FL bacterial communities (Stevens et al. 2005a), or in

manipulated mesocosms (Lebaron et al. 1999, Pinhassi et al. 2004, Riemann et al. 2000). In

other cases when more subtle variations or discrimination against specific target groups may

occur, this approach appears not sensitive enough to comprehensively detect these changes.

Then the application of more specific primer sets is a valuable tool to detect these changes

which are an important indication of distinct responses of the bacterial communities to their

changing environment.

Acknowledgements. We appreciate the hospitality and assistance of the RV Senckenberg

crew. We thank B. Kuerzel and R. Weinert for dry weight analyses and H.-P. Grossart for

valuable discussions. This work was supported by the Deutsche Forschungsgemeinschaft

(DFG) within the research group “BioGeoChemistry of Tidal Flats” (FG 432 TP5).


36

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39

Table 1. Primers used in this study. _______________________________________________________________________________________________________________

Primer Sequence (5´- 3´) E. coli 16S Target group Reference rRNA position

_______________________________________________________________________________________________________________

GC-341F CCTACGGGAGGCAGCAG 341 – 358 Bacteria (Muyzer et al. 1993) 907RM CCGTCAATTCMTTTGAGTTT 907 – 924 Universal (Muyzer 1998) GC-CF319f GTACTGAGACACGGACCA 319 – 336 Bacteroidetes (Manz et al. 1996) ALF968r GGTAAGGTTCTGCGCGTT 968 – 985 alpha-Proteobacteria (Neef 1997) GC-ROSEO536Rf CGGAGGGGGTTAGCGTTG 536 – 553 Roseobacter clade (Brinkmeyer et al. 2000) GRb735r a GTCAGTATCGAGCCAGT(G/A)AG 735 – 754 Rhodobacter group (Giuliano et al. 1999)

_______________________________________________________________________________________________________________ a modified

Table 2. Phylogenetic affiliation of strains and species used for the specificity test of the Roseobacter primer set. Strains are from the culture collection of our lab or the DSMZ: German collection for cell cultures and microorganisms. DSMZ strain numbers are given in parenthesis. Class Strain or species (DSM No.) Acc. No.

alpha-Proteobacteria a TL AY177716

(Roseobacter clade) T5 AY177712 T11 AY177714 TY AY841788 D1 AY841770 D4 AY841771 HP12 AY841769 HP14w AY841773 HP29w AY239008 HP30 AY239009 HP32 AY841774 HP37 AY239010 HP44 AY841765 HP47 AY841776 HP50 AY841778 ROS2 AY841779 ROS4 AY841780 ROS7 AY841781 ROS8 AY841782 AP-27 AY145564 H43-35 AY841784 H55 AY841765 GWS-BW-H22M AY515407 GWS-BW-H66M AY515422 GWS-BW-H71M AY515423 Roseobacter gallaeciensis (12440) Y13244 Roseobacter denitrificans (7001) M59063 Ruegeria algicola (10251) X78315 Ruegeria gelatinovorans (5887) D88523 Roseovarius tolerans (11457) Y11551 Leisingera methylohalidivorans (14336) AY005463 Sulfitobacter pontiacus (10014) Y13155


40

Table 2 cont. Class Strain or species (DSM No.) Acc. No.

alpha-Proteobacteria b Paracoccus aminophilus (8538) Y16929

beta-Proteobacteria b Aquaspirillum delicatum (11558) AF078756

Burkholderia pyrrocinia (10685) AB021369 Sphaerotilus natans (6575) L33980 gamma-Proteobacteria

b Pseudomonas putida (548) AF094741

Pseudeoalteromonas atlantica (6839) X82134 Fundibacter jadensis (12178) AJ001150 delta-Proteobacteria

b Desulfococcus multivorans (2059) AF418173

Desulfobulbus mediterraneus (13871) AF354663 Pelobacter venetianus (2394) U41562 Flavobacteria

b Muricauda ruestringensis (13258) AF218782

Bacilli b Bacillus marinus (1297) AJ237708

Bacillus subtilis (7, 10) AJ276351 Lactobacillus plantarum (20205) n. a. Actinobacteria

b Streptomyces violaceoruber (40701) n. a.

Streptomyces glaucescens (40155) D44092 Streptomyces antibioticus (40715) n. a. Arthrobacter nicotinovorans (420) X80743 a positive control b negative control n. a. : not available


41

Figure legends

Fig. 1. Temperature and salinity (A), total particulate carbon, particulate organic carbon

(POC) and phytoplankton carbon as % of POC (B), phytoplankton cell counts and species

composition (C), and abundance of particle-attached and free-living bacteria (D) in the

German Wadden Sea from 12 April to 29 June 2000.

Fig. 2. DGGE fingerprints of the free-living (FL) and particle-attached (PA) bacterial

communities of the German Wadden Sea from 12 April to 29 June 2000 using primer sets for

16S rRNA genes of Bacteria (EUB; A), Bacteroidetes (CFB; B), alpha-Proteobacteria (ALF;

C) and the Roseobacter clade (ROS; D). The numbered arrows mark excised and sequenced

bands. Because of the small fragment size of the Roseobacter amplicons the DGGE bands

were not excised for sequencing.

Fig. 3. Cluster analyses of the DGGE banding patterns of particle-attached (PA) and free-

living (FL) Bacteria (EUB; A), Bacteroidetes (CFB; B), alpha-Proteobacteria (ALF; C) and

the Roseobacter clade (ROS; D) using UPGMA. The similarity matrix was calculated using

Pearson correlation.

Fig. 4. Phylogenetic trees of Proteobacteria (A) and the Bacteroidetes phylum (B) calculated

with Maximum-Likelihood based on 16S rRNA gene fragments. Sequences obtained in this

study are highlighted in bold.


42

0

2

4

6

12 April 26 April 10 May 24 May 07 June 21 June

Date

Cel

l C

oun

ts m

l-1

*106

attached

free-living

0

2

4

6

8


mg

l-1

0

20

40

60

%

C total

POC

% C phyt./C org.

A

B

C

D

Cel

lCou

nts

l-1x1

03

5

10

15

20


°C

25

30

35

‰

TemperatureSalinity

Cel

lCou

nts

ml-1

x105

Date

mg

l-1

8

6

4

2

%

60

40

20

0

Cryptophyceae sp.

Dinophyceae

Pennate diatoms

Thalassionema nitzschioides

Raphoneis amphiceros

Pseudonitzschia pungens

Plagiogrammopsis vanheurckii

Cylindrotheca closterium

Centric diatoms

Thalassiosira punctigera

Guinardia delicatula

8

6

4

2

6

4

2

0

0

00

2

4

6


Date

Cel

l C

oun

ts m

l-1

*106

attached

free-living

0

2

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8


mg

l-1

0

20

40

60

%

C total

POC

% C phyt./C org.

A

B

C

D

Cel

lCou

nts

l-1x1

03

5

10

15

20


°C

25

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35

‰

TemperatureSalinity

Cel

lCou

nts

ml-1

x105

Date

mg

l-1

8

6

4

2

%

60

40

20

0

Cryptophyceae sp.

Dinophyceae

Pennate diatoms






Centric diatoms



Cryptophyceae sp.

Dinophyceae

Pennate diatoms






Centric diatoms



8

6

4

2

6

4

2

0

0

0

Fig. 1. Rink et al.


43

April May June April May June

12 26 03 10 17 24 31 14 29 12 26 03 10 17 24 31 14 29Std. Std.

EUB

CFB

ALF

ROS

A

B

C

D

FL PA

Fig. 2. Rink et al.


44

Fig. 3. Rink et al.

ROS

ALF

CFB

EUB

A

B

C

D


45

Fig. 4A. Rink et al.

A

gamma

beta

RCA

WAC I

alpha


46

B

Fig. 4B. Rink et al.

47

III.

Tidal effects on coastal bacterioplankton

Kapitel III Tidal effects on coastal bacterioplankton

48

Tidal effects on coastal bacterioplankton

Beate Rink, Torben Martens, Doreen Fischer, Andreas Lemke, Hans-Peter Grossart, 1

Meinhard Simon, and Thorsten Brinkhoff *

Institute for Chemistry and Biology of the Marine Environment (ICBM),

University of Oldenburg, P.O. Box 2503, D-26111 Oldenburg, Germany

1 Present address: Institute of Freshwater Ecology and Inland Fisheries, Department of

Limnology of Stratified Lakes, Alte Fischerhuette 2, D-16775 Stechlin, Germany

Running head: Tidal effects on coastal bacterioplankton

Key words: Aggregates, bacteria, CARD-FISH, DGGE, tidal flat, Wadden Sea

__________________________________________________________________________

* To whom correspondence should be addressed ([email protected]).


49

Acknowledgements

We appreciate the hospitality and assistance of the RV Senckenberg crew. We gratefully

acknowledge K. Ishii and R. Amann for the introduction of the CARD-FISH method and M.

Mußmann for valuable methodical discussions. We thank B. Kuerzel and R. Weinert for Chl

a and dry weight analyses and R. Reuter and T. Badewien for the supply of temperature and

salinity data for July 2005. This work was supported by the Deutsche

Forschungsgemeinschaft (DFG) within the research group “BioGeoChemistry of Tidal Flats”

(FG 432-5).

Abstract

Tidal flats are highly dynamic and productive ecosystems, strongly influenced by

hydrodynamic forces and tidal events. We examined the impact of the tide on the composition

of free-living (FL) and particle-attached (PA) bacterioplankton in the Wadden Sea, southern

North Sea, in November 1999, May 2000 and July 2005 complementing the study of Grossart

et al. (2004) which focused mainly on the dynamics of suspended particulate matter (SPM)

and bacterial abundance. FISH, CARD-FISH and DGGE fingerprints with primer sets for

Bacteria, Bacteroidetes, alpha-Proteobacteria and the Roseobacter Clade based on 16S rRNA

genes and 16S rRNA were applied. In addition, suspended particulate matter (SPM),

particulate organic carbon (POC), chlorophyll a (Chl a), bacterial cell counts and bacterial

protein production rates (BPP) were measured. The introduction of North Sea water in the

Wadden Sea system was shown by changes of the water temperature, salinity values and

oxygen saturation during high tides (HT). In spring and summer, particulate organic carbon

(POC) ratio and cell specific bacterial production was increased and altered with the tide.

Surprisingly, these strong variations had only slight effect on the composition of the bacterial

communities. FISH counts revealed abundance of Bacteroidetes, alpha- and gamma-

Proteobacteria. On particles, also beta-Proteobacteria were detected with 15.29% DAPI in

May 2000 and 8.19% DAPI in July 2005. The Roseobacter Clade constituted almost all FL

but only one third of the PA alpha-Proteobacteria. In spring and summer, FISH counts

showed higher variations within the bacterial groups compared to autumn and increased

abundances of PA alpha-Proteobacteria. A clear relationship between these variations and the

tide was not found. DGGE banding patterns of 16S rRNA gene fragments were highly stable

even on the group-specific level. At higher resolution based on cDNA amplicons, higher

richness was detected but only few variations related to the tide appeared. Phylogenetic

analysis revealed that prominent bands were affiliated to the RCA, the WAC I and the


50

SAMMIC cluster. These clusters were found in previous studies of the Wadden Sea and are

distributed worldwide in temperate and polar marine regions. Our results suggest that the

stability of the abundant bacterial groups is a consequence of the strong changes within the

Wadden Sea ecosystem selecting highly adapted species which persist even on a long-term

scale.

Introduction

Tidal flats are coastal ecosystems appearing in temperate and tropical regions and are

composed of sandy or muddy sediment. They are strongly exposed to the tide and thus form a

very stressful environment for the indwelling organisms. Furthermore, tidal flats provide

complex substrates due to high loads of suspended particulate matter (SPM) deriving from the

land and the sea, and belong to the most productive ecosystems in the world (Dittmann,

1999). The German Wadden Sea, a tidal flat system in the southern part of the North Sea, is

the largest tidal flat system worldwide comprising an area of 7500 km2. Several monitoring

studies examined the status of this ecosystem on spatial, diurnal and seasonal scales. It is a

well studied area in terms of suspended particulate matter flux and particle size distribution

(Eisma and Li 1993; van Leussen 1996; Behrends und Liebezeit 1999; Mikkelsen 1998;

Fugate and Friedrichs 2003; Grossart et al. 2004; Lunau et al. 2006), primary production (van

Duyl et al. 1999; Tillmann et al. 2000; Niesel and Günther 1999) and several planktonic and

benthic organisms (Günther, 1999; Dittmann, 1999), but the main mediators for organic

matter decomposition, the bacterial communities, remained disregarded.

Correlation of SPM composition and -variation and microbial activities in the Wadden Sea

were investigated by Grossart et al. (2004) during two tidal cycles in November 1999 and

May 2000. Strong changes in several parameters were detected due to differences in tidal

currents and subsequent introduction of North Sea water masses into the Wadden Sea

ecosystem at high tides (HT). In November 1999, water temperatures and salinity were higher

during HT. SPM and PA bacteria were correlated and fluctuated during the tidal cycle

dependent from current velocities. In May 2000, water temperatures were higher but oxygen

saturation was increased during HT due to the North Sea water influence. Cell specific

bacterial production increased with rising tide indicating response of the bacterial community

to tidal changes. Also seasonal influences were detected particularly for the POC ratio and FL

cell counts which were increased in May 2000. These observations substantiated the

assumption that the tide may strongly affect the composition of pelagic bacterial

communities. So far, only few studies focused on short-term influences on bacterioplankton.


51

Pernthaler and Pernthaler (2005) described variations in the proliferation of North Sea

bacteria during a tidal cycle. Diurnal changes in pigment concentrations derived from bacteria

in Baltic Sea water were examined by Koblizek et al. (2005). In dilution cultures of North Sea

bacteria, the impact of UV on bacterial communities during a 24 hours period was

investigated using DGGE profiles (Winter et al., 2001).

In previous studies, the main interest of Wadden Sea investigations focused on the

composition and abundance of sediment bacteria. Llobet-Brossa et al. (1998) showed

dominance of Bacteroidetes and sulfate-reducing bacteria in the sediment surface layer. At

two different sampling sites of Wadden Sea sediment, Mussmann et al. (2005) isolated

several delta-Proteobacteria. In the water column, the Wadden Sea bacterioplankton was

recently examined on a qualitative level by Stevens et al. (2005a, b) and Rink et al. (2006)

using DGGE. It was demonstrated that many bands persisted over long-term periods, but also

variations in the banding patterns were detected in spring and summer months. Rink et al.

(2006) showed correlation of phytoplankton composition changes and alteration in the

bacterial community composition using group-specific primer sets. As these studies were

performed on a monthly or weekly scale, the influences of tidal currents and the fast exchange

of different water bodies in the Wadden Sea on the bacterioplankton remained unknown.

Thus, the aim of this study was to examine the impact of the tide on the composition and

abundance of the bacterial communities in the Wadden Sea with respect to seasonal aspects.

With this study, we complete the results by Grossart et al. (2004) with detailed bacterial

community composition analysis based on the same sampling campaigns and an additional

sampling in July 2005.


52

Materials and methods

Sample collection and processing– Samples were taken in November 1999, May 2000 and

July 2005 from shipboard in the major channel of the backbarrier tidal flat system of the

German Wadden Sea (53° 44.9´N, 07° 40.0´E). The sampling period was 19 hours in

November 1999, 22 hours in May 2000 and 12 hours in July 2005. In November 1999 and

May 2000, surface water was collected every hour and filtrated for suspended particulate

matter (SPM), chlorophyll a (Chl a), particulate organic carbon (POC) and bacterial cell

counts according to Grossart et al. (2004). In July 2005, samples for SPM, Chl a, POC and

bacterial cell counts were taken and processed according to Lunau et al. (2006). Hydrographic

data of July 2005 were measured by a CTD probe (SeaCat 19plus, Seabird, Washington,

USA).

For DGGE analyses, one hundred ml of seawater were filtered at high tides, mean tides

and low tides, fractionated onto polycarbonate-filters (diameter 47 mm, Nuclepore) and stored

at -20°C in the dark. A pore size of 5.0 µm was used to obtain particle-attached (PA) and 0.2

µm to obtain free-living (FL) bacteria. For Fluorescent In Situ Hybridization (FISH) and

Catalyzed Reporter Deposition (CARD)- FISH analysis, 2 - 4 ml were filtered onto 5.0 µm

polycarbonate-filters (diameter 25 mm, Nuclepore) and 2 ml of the filtrate were subsequently

filtered onto 0.2 µm polycarbonate-filters. After fixation with paraformaldehyde (4% w/v) for

one hour filters were stored at -20°C in the dark until further processing. Sediment cores in

November 1999 and May 2000 were taken with Plexiglas tubes (36 mm diameter) at low tide

on an intertidal mud flat about 200 m away from the ship. Two mm of the upper surface layer

were sliced, transferred into sterile caps and kept frozen at -20°C.

SPM dry weight, particulate organic carbon and chlorophyll a– Filters for SPM dry

weight were dried for 12 h at 60°C, adapted to room temperature and weighed on a

microbalance. Dry weight was calculated as the difference of filter weight before and after

filtration (500 ml filtration volume, glass fiber filters, GF/F, Whatman). For particulate

organic carbon, filters were exposed to hydrochloric acid fume for 12 h and subsequently

analyzed by a FlashEA 1112 CHN-analyzer (Thermo Finnigan). Chlorophyll a was

determined photometrically after extraction in hot ethanol according to Von Tuempling and

Friedrich (1999). For further details concerning SPM, POC and Chl a analyses see Lunau et

al. (2006).


53

Bacterial cell counts– Bacterial abundance was determined with SybrGreen I as described

by Lunau et al. (2006) with slight modifications. The detachment of particulate bacteria was

performed using methanol (30% v/v) and ultrasonic treatment for 15 min at 35°C. For the

enumeration of bacterial cells, subsamples of 1 ml were filtered onto black polycarbonate

filters (0.2 µm pore size, Poretics). Abundance of particulate bacteria was calculated as the

difference between total and free-living cell counts.

Bacterial Production –The bacterial production rate was estimated by the incorporation of 14C-Leucin (Simon and Azam 1989). Samples were incubated in triplicates with 14C-Leucin

(306 mCi/mmol, Hartmann Analytic, Germany) at a final concentration of 70 nmol l-1 to

ensure saturation of uptake systems. Formalin fixed water samples (2% v/v) were used as

controls. The samples were incubated in 5 ml plastic tubes in the dark at in situ temperature

for up to 1 hour. To avoid sedimentation, incubation was performed on a plankton wheel. The

incubation was linear for at least 1 hour and terminated by the addition of formaldehyde (2%

v/v). After fixation, the samples were filtered onto 0.45 µm nitrocellulose filters (Sartorius,

Germany) and extracted with ice-cold 5% trichloracetic acid (TCA) for 5 min. Subsequently,

filters were rinsed twice with 3 ml ice-cold TCA (5% v/v). To dissolve the filters, 4.5 ml of

scintillations cocktail was added. Vials were shaken vigorously and radioactivity was

determined afterwards in a scintillation counter. Standard deviation of triplicate measurement

was usually <15%.

FISH and CARD-FISH– FISH filters in November 1999 and May 2000 were rinsed with 1

ml phosphate buffered saline (PBS 1x) and dehydrated with 50%, 80% and 100% ethanol.

Subsequently, bacterial cells were hybridized for 5 h and washed 20 min at 46°C following

the protocol of Glöckner et al. (1996). Oligonucleotide probes for various phylogenetic

groups were used in November 1999: EUB 338 (Amann et al. 1990), ALF968, GAM42a

(Manz et al. 1992), CF319a (Cytophaga/Flavobacteria, Manz et al. 1996), ARCH915 (Stahl &

Amann 1991) and SRB385 (Amann et al. 1990). In May 2000, the additional probes BET42a

(Manz et al. 1992) and SRB385db (Rabus et al. 1996) were used. All filters were

counterstained with DAPI (4´,6´-Diamidino-2-phenylindol, 1µg/ml). In July 2005,

hybridizations were performed using the CARD-FISH method following the protocol of

Sekar et al. (2003). After fixation, samples were embedded in agarose (0.2%) and treated with

lysozyme (10mg/ml). Hybridization conditions were 3 h of hybridization at 35°C, 10 min

washing at 37°C and 30 min. amplification at 37°C. The following oligonucleotide probes


54

labeled with horseradish-peroxidase (HRP) were used for CARD-FISH: EUB338, ALF968,

BET42a, GAM42a, ROS536 (Brinkmeyer et al. 2000) and NON338 (Wallner et al. 1993).

Tyramine-HCl was labeled with Fluorescein-5-isothiocyanate (FITC) as described by

Pernthaler et al. (2002). To avoid unspecific accumulation of dye in the cells, the last washing

step in PBS (1x) amended with TritonX-100 (0.05%) was extended to 30 min.

Counterstaining was performed with Vectashield-mounting medium with DAPI (1.5µg/ml;

Vector Laboratories, Peterborough, England).

Nucleic acid extraction– For samples taken in November 1999 and May 2000, genomic

DNA was isolated by phenol-chloroform extraction after bead beating as described by Selje

and Simon (2003). In July 2005, DNA and RNA were co-extracted from the same filter using

phenol-chloroform calibrated with sodium acetate RNA-buffer (50 mM, pH 4.2) containing

EDTA (10 mM) and Polyvinylpolypyrrolidone (PVPP, 2 g l-1). All steps were done under

sterile conditions using Diethylpyrocarbonate (DEPC, 0,1%)-treated ingredients. After

precipitation in isopropanol at -20°C overnight nucleic acids were resuspended in

DNase/RNase-free molecular grade water (Eppendorf, Germany). The RNA was incubated

with DNase I (5 U ml-1) for 1 h at 37°C and precipitated as described above. DNase digestion

was repeated until no DNA contamination could be detected by PCR with primers specific for

bacterial 16S rRNA genes. Samples were stored at -20°C until further processing.

PCR amplification – 16S rRNA gene fragments for subsequent DGGE analyses were

amplified using the primer pair GC 341F and 907RM (Muyzer et al. 1998) in an Eppendorf

Mastercycler (Eppendorf, Hamburg, Germany). In July 2005, the additional primer pairs GC

CF319aF and 907RM, GC 341F and ALF968R, and GC ROS 536F and GRB 735R were used

to investigate the bacterial community composition on a group-specific level. Primer

sequences and PCR conditions were described earlier by Rink et al. (2006). Four µl of the

amplification products were analyzed by electrophoresis in 2% (w/v) agarose gels and stained

with ethidium bromide (1 µg ml-1). For sequencing analysis, PCR products were purified by

using the E.Z.N.A. Microspin Cycle-Pure Kit (Peqlab Biotechnologie GmbH, Erlangen,

Germany) following the instruction manual. To amplify bacterial RNA, the Qiagen RT-PCR

kit (Qiagen, Hilden, Germany) was used following the instruction manual for reverse

transcription of the RNA. Reverse transcription was done directly with bacteria- or group-

specific primer sets as described above. Subsequent PCR was performed in the same reagent

mix directly after transcription using specific conditions for the applied primer set.


55

DGGE analysis of PCR products– DGGE was performed with the INGENY phorU System

(INGENY International BV, Goes, Netherlands) using specific conditions for the applied

primer sets according to Rink et al. (2006). Gels were stained with SYBR Gold (Molecular

Probes, Inc.) after electrophoresis and documented digitally using a BioDoc Analyze

Transilluminator (Biometra, Göttingen, Germany). Bands were excised with a sterile scalpel,

suspended in 50 µl of molecular grade water (Eppendorf, Germany) and stored at –20°C until

reamplification.

Cloning– DGGE bands GWS-TC-a2-PA, GWS-TC-a6-FL, GWS-TC-c4-PA, GWS-TC-e9-

FL, GWS-TC-e11-FL, GWS-TC-e3-SE, GWS-TC-e4-PA, GWS-TC-e1-SE and GWS-TC-e2-

SE were cloned using the pGEM®-T Vector System II (Promega, Madison, USA) following

the instruction manual. Clones with inserts were picked, resuspended in molecular grade

water (Eppendorf, Germany) and screened by DGGE to check if the insert position matches

the position of the corresponding DGGE band. Adequate clones were amplified and

subsequently sequenced using the primers pUC/M13f and pUC/M13r (Messing 1983).

Sequencing and phylogenetic analysis – PCR products were sequenced using the

DYEnamic Direct cycle sequencing kit (Amersham Life Science, Inc.) and a Model 4200

Automated DNA Sequencer (LI-COR, Inc.) using GM8R (5´-TGGGTATCTAATCCT-3´) as

sequencing primer labeled with IRDyeTM800. In addition, DGGE bands were sequenced by

GeneArt (Regensburg, Germany) using the primer 907RM to enhance the sequence quality by

repeat determination. For all sequences, at least 400 bp were determined.

Construction of the phylogenetic trees was performed using the ARB software package

(http:/www.arb-home.de). Calculation of the backbone trees was done with the maximum

likelihood method using sequences of type strains of the selected phylogenetic groups with a

minimum sequence length of 1300 bp. To avoid uncertain alignments, positions at which less

than 50% of all sequences showed the same residues were excluded. Sequences with less than

1300 bp were added to the backbone tree using the maximum parsimony method and the

same filter. Five type strains belonging to Cyanobacteria were used as outgroup.

Nucleotide sequence accession number –The sequences obtained in this study are available

from GenBank under accession no. DQ911822 to DQ911842.


56

Statistics – Cluster analyses of DGGE banding patterns were performed using Gel Compar

II, version 2.5 (Applied maths, Kortrijk, Belgium). Calculations were done curve based using

Pearson correlation and UPGMA.

Results

Hydrographic data–In July 2005 water temperature ranged between 19.1 and 20.1°C

showing slight decrease at HT with incoming North Sea water. Salinity was almost constant

around 32 psu (Fig. 1a).

For hydrographic data as well as results concerning SPM dry weight, POC, Chl a, bacterial

abundance and bacterial production from November 1999 and May 2000 see Grossart et al.

(2004).

SPM dry weight, POC and Chl a–Dry weight of suspended particulate matter in July 2005

ranged from 7.5 to 8.2 mg l-1 at HT and reached the highest value of 36.8 mg l-1 at low tide

(Fig. 1b). Despite highest current values at mean tide, SPM dry weight was average (21 to

29.2 mg l-1). POC values (see supplementary data) were low around LT (0.39 mg l-1) and

increased towards HT (up to 2.15 mg l-1) and constituted 1.1% (LT) to 28.7% (HT) of the

SPM dry weight. The average value was 7.6% dry weight. Chlorophyll a increased from 3.13

µg l-1 at HT to 6.7 µg l-1 around LT and decreased again to 3.76 µg l-1 at high tide (Fig. 1c).

Phaeopigments showed the same tidal dynamic and ranged from 0.30 to 5.12 µg l-1 with the

highest ratio phaeopigments/chlorophyll a of 1.18 at mean tide.

Bacterial abundance and bacterial production– In July 2005, total bacterial cell counts

varied between 1.28 to 3.56 x 106 cells ml-1 (Fig 1d). Free-living (FL) bacteria showed a

minimum of 0.97 x 106 cells ml-1 one hour after the first high tide and increased slightly to a

maximum of 2.30 x 106 cells ml-1 at one hour after mean tide 1 (MT 1). Afterwards, FL cell

numbers decreased again until next mean tide (MT2) and then ranged between 1.46 and 1.75

x 106 cells ml-1 until second high tide. Particle-attached (PA) bacteria were lowest around

high tide with 0.09 to 0.45 x 106 cells ml-1 and highest between MT1 to MT2 (0.96 to 1.33 x

106 cells ml-1) with a slight decrease around LT. In average, PA bacteria accounted for 28% of

total bacterial cell counts. To compare cell densities of PA bacteria, cell counts per mg dry

weight were calculated (see supplementary data). Density of PA cells was highest one hour

after MT and two hours after LT due to lower dry weight values and equal cell counts.


57

Bacterial production (BPP, Fig. 1e) in July 2005 ranged between 0.55 µg l-1 h-1 at both

high tides to 3.33 to 3.64 µg l-1 h-1 during LT and MT2.

FISH counts – FISH counts detected with probe EUB338 were lowest in May 2000 with a

mean value of 56.77% DAPI on particles and 48.42% DAPI in the FL fraction (Table 1, Fig.

2). Highest FISH counts were measured in November 1999 with 73.8% DAPI on particles and

61.24% DAPI in the FL fraction. In July 2005, FISH counts were similar to November with

70.42% DAPI on particles and 59.44% DAPI for free-living bacteria. Highest variation within

the cell counts were detected in May 2000 during a spring phytoplankton bloom, while lowest

variation was observed in November 1999 when hydrodynamic forcing controlled the

processes in the water column (Fig. 2). Generally, a higher percentage (% DAPI) of cells

could be detected on particles compared to the free-living fraction.

Specific probes revealed high numbers of Bacteroidetes, alpha-, beta- and gamma-

Proteobacteria in the Wadden Sea (Table 1, Fig. 2). In November 1999 and May 2000, also

low cell counts of Archaea and sulphate-reducing bacteria were detected (below 5% each).

In May 2000, cell counts of Bacteroidetes were 27.59% DAPI on particles and 13.55%

DAPI in the FL fraction. In November 1999, Bacteroidetes showed lower FISH counts on

particles 20.93% DAPI, but higher numbers (19.16% DAPI) in the FL fraction (mean values).

In July 2005, lowest cell counts of Bacteroidetes were detected with 15.02% DAPI in the PA

fraction and 11.52% DAPI in the FL fraction.

Abundances of alpha-Proteobacteria on particles were lowest in November 1999 with

9.53% DAPI and highest in May 2000 with 30.59% DAPI. In the free-living fraction, alpha-

Proteobacteria contributed 13.76% of the total cell counts in November 1999 but only 7.21 to

7.87% DAPI in spring and summer. In July 2005, the Roseobacter Clade, as part of the alpha-

Proteobacteria, represented 4.34% DAPI on particles and 5.09% DAPI in the FL fraction

contributing nearly 25% of the PA and 71% of the FL alpha-Proteobacteria.

Abundances of gamma-Proteobacteria ranged between 16.49 and 28.83% DAPI on

particles and between 9.83 and 17.12% DAPI in the FL fraction. Hybridization with probe

BET42a revealed 8.19 to 15.29% DAPI counts on particles in spring and summer showing a

certain abundance of PA beta-Proteobacteria in this coastal environment.

DGGE banding patterns and Cluster analysis–DGGE analysis of the tidal cycles in

November 1999, May 2000, and July 2005 revealed distinct bacterial populations for the

sediment surface, the PA and the FL bacterial communities (Fig. 3, 4, 5). In November 1999


58

and May 2000, fingerprints within the different fractions showed almost identical banding

patterns during the tidal cycles (data not shown). Highest band numbers in all seasons were

determined in the PA fraction (24 bands in Nov 1999, 13 bands in May 2000, 25 bands in July

2005) and a direct comparison of November 1999 and May 2000 showed only few variations

in the banding patterns at all (Fig. 3). The banding patterns of the PA bacterial communities

indicate an overlap with those resulted from sediment and the FL bacteria, as both fractions

show bands on the same height as in the PA fraction.

In July 2005, the FL and PA bacterial communities were also distinct as shown by the

DGGE fingerprints and cluster analysis. In addition, high stability of the communities was

observed on the DNA level, however, some bands appeared only in one or two samples, i. e.

at certain times of the tidal cycle (Fig. 4). Analysis with the bacteria specific primers revealed

for the PA fraction, e. g., that samples obtained during the first mean tide (MT1) and LT

showed one additional band (GWS-TC-e6-PA), which probably led to higher similarity of

these two samples in the cluster analysis (Fig. 5). In the FL fraction, one band appeared

exclusively at the first HT (GWS-TC-e8-FL).

The cDNA fingerprints of Bacteria revealed a slightly higher richness for the FL fraction

with up to 23 bands at HT compared to a maximum of 18 bands in the DNA banding patterns.

At HT, two additional bands appeared in the FL fraction which were neither detected in the

DNA fingerprints nor in the other cDNA samples (GWS-TC-e10-FL and GWS-TC-e11-FL).

On the group-specific level, DGGE analysis of Bacteroidetes showed stable DNA based

banding patterns in the PA fraction (Fig. 4, CFB). In the FL fraction, one prominent band

appeared exclusively at MT1 and LT (GWS-TC-c1-FL). The cDNA banding patterns showed

few differences compared to the DNA fingerprints and during the tidal cycle as well. In the

PA fraction, bands no. 2, 3, and 4 (GWS-TC-c2-PA, GWS-TC-c3-PA and GWS-TC-c4-PA)

occurred individually at different times and did not appear in the DNA based banding

patterns. Cluster analysis of the Bacteroidetes fingerprints revealed distinct fractions of PA

und FL fractions except for the second mean tide (MT2) of the cDNA FL fraction, which

showed higher similarity to the PA cluster (Fig. 5). The FL cluster was divided in two

subclusters of DNA and cDNA samples showing high similarity within DNA samples (>90%

Pearson correlation). The PA cluster was also subdivided into DNA and cDNA clusters

except the DNA sample at LT which fell into the cDNA group.

DGGE analysis of the alpha-Proteobacteria showed low richness within the DNA

fingerprints with 14 bands in the PA fraction and 8 bands in the FL fraction (Fig. 4). Slight

differences during the tidal cycle were detected on particles at LT with decreased band


59

intensity of bands at the standard height, also reflected by cluster analysis. The cDNA

banding patterns showed higher richness in both fractions compared to the DNA level. A

maximum of 25 bands was counted at LT on particles and 11 bands were visible at HT in the

FL fraction. These differences between DNA and cDNA resulted in distinct clusters as shown

by cluster analysis for both, PA and FL fraction (Fig. 5). Tidal differences traced back to

alpha-Proteobacteria were detected in the cDNA fingerprints during MT1 and LT (Fig. 4,

GWS-TC-a2-PA, and GWS-TC-a5-PA) or generated by chloroplast phylotypes (GWS-TC-

a3-PA, GWS-TC-a4-PA).

For the Roseobacter Clade, DNA based fingerprints revealed a maximum of 14 bands in

the PA fraction and 11 bands in the FL fraction. The cDNA banding patterns showed

significantly higher richness in the PA fraction (20 bands at LT) compared to DNA

fingerprints, forming distinct clusters (Fig. 5). Slight differences during the tidal cycle were

also found for this group (Fig. 4). While the cluster analysis demonstrated high similarity of

the DNA and cDNA patterns of the FL bacterial communites, the PA samples of the

Roseobacter Clade clustered distinct in DNA and cDNA (Fig. 5).

Phylogenetic affiliation – Sequence analysis of DGGE bands revealed that most of the

prominent bands were affiliated to recently described clusters of Wadden Sea bacteria (WAC

I, Stevens et al. 2005b) or of worldwide distributed bacteria (RCA, Selje et al. 2004;

SAMMIC, Stevens et al. 2005b) as shown in Fig 6A. From the DGGE banding patterns of

Bacteria, phylotypes belonging to the alpha-, gamma- and beta-Proteobacteria were obtained

reflecting the FISH and CARD-FISH results. One phylotype of delta-Proteobacteria and two

sequences of Actinobacteria were detected in the sediment fractions of November and May

(Fig. 3 and 6, GWS-TC-e1-SE, GWS-TC-e2-SE, GWS-TC-e3-SE). In total, four bands were

identified as chloroplasts (Fig. 3: GWS-TC-e5-PA; Fig. 4: GWS-TC-e7-FL, GWS-TC-a3-PA,

GWS-TC-a4-PA). GWS-TC-e11-FL was distantly related to Acidocella aminolytica (Fig. 6A)

and was not related to any sequence found in the Wadden Sea or North Sea before. Two

sequences fell in the WAC I cluster (GWS-e6-FL, GWS-e7-FL), GWS-TC-e9-FL clustered

with the Wadden Sea clone GWS-FL-5 (Stevens et al. 2005a), three sequences belonged to

the RCA cluster (GWS-TC-e6-PA, GWS-e12-PA, GWS-e5-FL) and GWS-e3-FL was related

to clone NAC11-7 (Fig. 6 A).

Two phylotypes of DNA and cDNA fingerprints clustered together within the gamma-

Proteobacteria (GWS-TC-e8-FL, GWS-TC-e10-FL). They were affiliated to

Stenotrophomonas maltophilia and appeared solely during HT in the FL fraction. Two of the


60

sediment derived sequences (GWS-TC-e3-SE, delta-Proteobacteria; GWS-TC-e2-SE,

Actinobacteria) were affiliated to clones which were found in the North Frisian Wadden Sea

sediments (Sylt 19, clones Sylt 21, Fig. 6 A).

Bacteroidetes affiliated phylotypes were exclusively obtained with the specific primer set

(GWS-TC-c1-FL to GWS-TC-c4-PA) confirming the bias against this phylum by the use of

the Bacteria-specific primer set (Kirchman 2002). Within the Bacteroidetes phylum, GWS-

TC-c1-FL represented the most pronounced diurnal change in the FL fraction of the DNA

fingerprints. This phylotype was related to GWS-c14-PA which was detected in May 2000

with specific primer sets for Bacteroidetes (Rink et al. 2006). Both were affiliated to clone

CF60 (AY274866) derived from the Delaware estuary. GWS-TC-c2-PA was detected during

HT in the PA fraction of the cDNA fingerprints and the phylogenetic affiliation showed no

relationship to known Wadden Sea or North Sea organisms. Another cDNA phylotype, GWS-

TC-c3-PA, was related to Lutibacter litoralis, isolated from a tidal flat system in Korea (Choi

and Cho 2006). GWS-TC-c4-PA was related to two novel species, Krokinobacter genikus

(Khan et al. 2006), isolated from marine sediment in Japan, and Dokdonia donghaensis (Yoon

et al. 2005), isolated from seawater of the Korean East Sea.

The application of the ALF968r primer revealed two chloroplast sequences (GWS-TC-a3-

PA, GWS-TC-a4-PA), and three phylotypes of alpha-Proteobacteria. Within the alpha-

Proteobacteria, the cDNA derived phylotypes clustered with Silicibacter lacuscaerulensis

(GWS-TC-a2-PA) and Sphingomonas paucimobilis (GWS-TC-a5-PA).

Discussion

Grossart et al. (2004) demonstrated that bacterial dynamics in the Wadden Sea are

controlled by two major factors: resuspension of sediment and phytoplankton growth.

Another strong influence is the introduction of North Sea water in the tidal basin at high tides

(HT). In November 1999, sediment resuspension was the dominating process in the water

column (Grossart et al. 2004). SPM flux and changes in particulate carbon directly reflected

the tidal dynamics. In May 2000, a strong influence of a phytoplankton bloom was observed

resulting in higher Chl a concentrations and an increase of the POC ratio and free-living

bacterial cells. In July 2005 the Chl a concentrations were lower than during the

phytoplankton bloom in May but typical for summer months in the Wadden Sea (Lunau et al.,

2006). SPM values in July were much lower compared to November and May, probably due

to a lower resuspension rate and lower abundance of phytoplankton, respectively (see


61

Grossart et al. 2004). Bacterial abundance on particles was similar in all seasons following

SPM dynamics. Cell counts of free-living bacteria were lower in July than in May but similar

to November suggesting lower dissolved organic carbon concentrations in the water column

(Grossart et al. 2004). The bacterial production was lowest around HT contrary to May and

other investigations (Lunau et al. 2006). Bacterial production values of May showed not a

tidal, but a diurnal, pattern, with high rates during the day and reduced values at night

(Grossart et al. 2004).

Grossart et al. (2004) showed that free-living bacteria were influenced by SPM

concentrations, total carbon (TC) and particulate organic carbon in November 1999 and May

2000. The abundance of particulate bacteria correlated with SPM, Chl a, total carbon and

particulate organic carbon. Thus, we investigated the composition of the bacterial

communities in the Wadden Sea by FISH and DGGE expecting strong changes of the

populations following the tidal dynamics of these parameters. By contrast, the fluctuation of

particulate carbon, SPM, Chl a and bacterial cell counts was not reflected by changes within

the phylogenetic groups during tidal cycles in all investigated seasons as shown by the FISH

results. Even the application of the highly sensitive CARD-FISH method in July 2005 did not

show any systematic impact of the tide on the bacterial community composition. In all

seasons, alpha- and gamma-Proteobacteria as well as Bacteroidetes were most abundant in

both fractions. This is in line with other studies from the North Sea (Eilers et al. 2000) and the

Weser estuary (Selje et al. 2003). The seasonal comparison of the FISH results revealed very

low variation within the bacterial groups during the tidal cycle in November in contrast to

May and July (Fig. 2). Largest variations and highest group-specific FISH counts were

obtained in May 2000 on particles. The exceeding of the group-specific over the EUB338

counts indicated high activity of the PA bacteria, as the signal strength of directly

fluorochrome-labeled oligonucleotide probes depends on the ribosomal RNA content of the

cells (Schut 1994). The SPM in May was enriched with phytoplankton derived POC which

intensely stimulated bacterial degradation processes of the organic matter as suggested by the

decoupling of activity parameters and tidal dynamics (Grossart et al. 2004). This may be one

reason for the fact that despite large variations within the bacterial groups, no clear

relationship was found between the FISH counts and the tide.

In spring and summer, beta-Proteobacteria were also investigated and very high

abundances were detected on particles. In May 2000, they constituted even 15.29% of the

total PA bacterial community (mean value, Table 1). This is in line with FISH counts of a

study showing that this group constituted 6% of DAPI cell counts in the marine section of the


62

nearby Weser estuary (Selje et al. 2003) and findings of several other authors (Rappé et al.

2000; Beja et al. 2002). High abundance of beta- Proteobacteria in this habitat was supported

by detection of phylotype GWS-e8-FL in May 2000, which clustered with other phylotypes

derived from the Wadden Sea (GWS-e4-FL, Rink et al. 2006; GWS-FL-6, Stevens et al.

2005) and the Weser estuary (DC11-51-11, Selje et al. 2005). The latter phylotype was

obtained from a 10-5 dilution step on marine medium even suggesting abundance of beta-

Proteobacteria in this saline environment.

DGGE results demonstrated that the composition of the bacterial communities in

November, May and July showed almost no changes during the tidal cycles. Sequencing of

prominent bands revealed that most of these phylotypes were affiliated to previously

described phylogenetic clusters, e. g. the RCA and the WAC I cluster within the alpha-

Proteobacteria (Selje et al. 2004; Stevens et al. 2005b) and the SAMMIC cluster within the

gamma-Proteobacteria (Stevens et al. 2005b). Organisms affiliated with the RCA cluster are

globally distributed in temperate and polar regions. They live exclusively in marine

environments freely suspended in the water column. The SAMMIC cluster comprises

organisms living on surfaces in marine environments. Bacteria of the SAMMIC cluster were

permanently detected on particles and on the sediment surface in the Wadden Sea during a

seasonal study (Stevens et al. 2005a). In parallel, Bowman et al. (2005) described the same

cluster with phylotypes from polar and temperate marine sediments detecting abundances of

up to 4% of the total bacterial community by quantitative RealTime-PCR. The distribution

and abundance of WAC I affiliated organisms is not yet clarified, but phylotypes belonging to

this cluster were also present in the Wadden Sea during all seasons (Stevens et al. 2005a).

Thus, bacteria belonging to these phylogenetic groups are not affected by short-term changes

generated by the tide as confirmed by our results.

In contrast to these permanently present organisms, some phylotypes were not detected

during the whole tidal cycles but in single samples (Fig. 4). The application of group-specific

primer sets revealed additional bands present at specific points of time. Although the use of

specific primer sets enhances the resolution of microbial studies (Abell & Bowman 2005;

Gich et al. 2005; Rink et al. 2006) the bacterial communities were remarkably stable even on

the specific level indicating low influence of the tide on the investigated groups.

Most tidal changes were detected by the RNA approach showing significantly higher

richness compared to the DNA fingerprints. The rRNA content of metabolically active

bacteria is higher than in dormant cells (Poulsen et al. 1993) and correlates with bacterial

growth rates (Delong et al. 1989). Thus, rRNA is suggested as activity indicator and reflects


63

responses of bacterial communities to environmental changes more pronounced than the

rRNA gene. Higher sensitivity concerning species diversity and appearance of community

shifts have been reported before, using rRNA based fingerprinting methods for marine

bacterioplankton (Schäfer et al. 2001; Moeseneder et al. 1999), drinking water supply systems

(Eichler et al. 2006) and bacterial assemblages of mariculture biofilter systems (Cytryn et al.

2005). Our study provides the first RNA based insights into bacterial communities of the

Wadden Sea tidal flat system, showing that most of the cDNA derived phylotypes, which

occured exclusively at specific points of the sampling, were not closely related to previously

described Wadden Sea organisms. Within the alpha-Proteobacteria, two of these phylotypes

were affiliated to described species (GWS-TC-a5-PA and GWS-TC-a2-PA, Fig. 6A).

Phylotype GWS-TC-a6-FL, which appeared at rising tide (MT2) was related to clone ZD0117

derived from the North Sea during a phytoplankton bloom (Zubkov et al. 2002). Largest

differences between DNA and cDNA fingerprints were reflected by the Roseobacter specific

fingerprints in the PA fraction. The richness within the cDNA banding patterns showed

almost two-fold increase compared to DNA. These findings were reflected by distinct clusters

of DNA and cDNA as revealed by cluster analysis. Hence, our results suggest that

significantly more particle-attached organisms are active than previously indicated by DNA

based methods and that the additional application of RNA based fingerprinting methods on a

group-specific level is essential to detect small-scale processes within microbial communities.

Relationships between additionally appearing bands and tidally generated processes were

hardly detected. In July 2005, bands GWS-TC-e8-FL and GWS-TC-e10-FL were pronounced

at HT in the FL fraction of DNA and cDNA samples. At HT, increased salinity, %POC values

and lower water temperature indicated influence of water from the open North Sea. The two

above mentioned phylotypes were closely related to Stenotrophomonas maltophilia (Fig. 6A,

96-97% sequence similarity). The genus Stenotrophomonas is widespread in terrestrial and

limnic habitats, but was also found in different marine samples, several times associated with

algae, sponges or dinoflagellates (e. g. Hagstrom et al. 2000; Seibold et al. 2001; Sfanos et al.

2005). Seibold et al. (2001) found a phylotype of Stenotrophomonas associated with the

dinoflagellate Noctiluca scintillans obtained from plankton hauls taken at Helgoland Roads

(German Bight, North Sea). Thus it is possible that the additional DGGE bands we observed

in our study derived from a Stenotrophomonas sp. introduced to the Wadden Sea with the

incoming tide. The same probably holds true for band GWS-TC-e11-FL (Fig. 4), which

belongs to the SAR116 cluster (Giovannoni et al. 1990) and band GWS-TC-a6-FL, which

appeared at rising tide (MT2) and was related to a clone obtained from the open North Sea


64

during a phytoplankton bloom (Zubkov et al. 2002). Appearance of these phylotypes only at

HT or rising tide, however, indicates low significance of the organisms within the Wadden

Sea ecosystem.

Cunha et al. (2001) showed that the activity of marine bacteria exposed to brackish water

increased significantly while bacteria of the estuary were less active when exposed to marine

water. Thus, the detection of differences at HT or rising tide may also be a single response of

marine bacteria to high concentrations of suspended substrates in the Wadden Sea.

Band GWS-TC-e6-PA (RCA cluster) disappeared in July 2005 at HT and MT2 (rising

tide) in parallel to the incoming North Sea water, but only in the particle fraction. This effect

was also detected in May 2000, but not in November 1999 (data not shown). In spring and

summer, SPM concentrations were lower compared to November with inversed tidal

dynamics (Grossart et al., 2004; this study). In May and July the inflow of North Sea water

resulted in lower amounts of SPM at HT suggesting that the disappearance of this band may

be due to a dilution effect. This assumption is supported by data showing much lower

numbers of RCA related phylotypes in the open North Sea compared to the Wadden Sea (H.

Giebel, University of Oldenburg, unpublished results). As no corresponding bands were

visible in the cDNA pattern of the particle fraction (Fig. 4), the significance of this organism

for SPM degradation in the Wadden Sea is also questionable.

On the group-specific level, GWS-TC-c1-FL appeared at MT1 and LT in the FL fraction

of the Bacteroidetes in the DNA and cDNA banding patterns. Closest relative was clone

GWS-c14-PA found in the Wadden Sea during a phytoplankton bloom in May 2000 (Rink et

al. 2006). This indicates that the organism is resident and active in the Wadden Sea. The

ecological function, however, remains unknown as this organism was found on particles

(Rink et al. 2006) and free-living (this study) as well.

Several other bands appeared on the group specific level at MT1 and LT (Fig. 4: GWS-TC-

c3-PA, GWS-TC-c4-PA, GWS-TC-a2-PA, GWS-TC-a5-PA) what might be explained by

resuspension of material from the sediment surface but this remains speculation.

The fingerprints of the Roseobacter Clade, which constituted a high fraction within the

alpha-Proteobacteria (approx. 25% on particles and 71% in the FL fraction as shown by

CARD-FISH results), were most stable even on the RNA level. Despite significantly higher

richness on particles (see above), almost no changes in the community composition were

observed for this group during the tidal cycle. Thus, our results indicate that the combined

application of group-specific primer sets and DNA/RNA based fingerprinting is sufficient for

the reliable examination of bacterial communities, i. e. to detect small changes within the


65

bacterial communities. In the tidal flat ecosystem of the Wadden Sea, however, only very few

changes could be observed and thus connection of these data with processes generated by the

tide were only sparely visible.

Overall our results demonstrate that even application of very sensitive investigation

methods like CARD-FISH and rRNA based DGGE analyses with specific primer sets

revealed highly stable FL and PA bacterial communities, almost not influenced by strong tidal

effects of this dynamic ecosystem. Seasonal changes and phytoplankton blooms may result in

larger tidal variations of group-specific abundances and the appearance of additional

organisms on a short time-scale but do not seem to influence the composition of the bacterial

communities significantly (Stevens et al. 2005a; Rink et al. 2006). Thus, we conclude that the

recurrent short- and long-term changes in the Wadden Sea resulted in the selection of highly

adapted organisms and lead to an exceptional stability of the bacterial communities in this

ecosystem. Further investigations are now required to clarify how the abundant bacteria are

involved in the microbial degradation processes and what makes them superior to other

organisms.


66

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Figure legends

Fig. 1. Temperature and salinity (a), dry weight (DW) and percent particulate organic carbon

(%POC) (b), chlorophyll a (chl a), phaeopigments and phaeopigment/chl a ratio (c),

abundance of total, particle-attached (PA) and free-living (FL) bacteria (d), and bacterial

protein production (BPP) (e) in the Wadden Sea during a tidal cycle in July 2005. X-axis:

Time of high tide (HT), mean tide (MT1/2) and low tide (LT).

Fig. 2. FISH counts for samples taken in November 1999, May 2000 and July 2005 during

tidal cycles at high tide, mean tide, and low tide. Results of July 2005 were obtained using

CARD-FISH. Calculation includes all values of single counting grids to display outliners

(dots). Box-Whisker-Plots show the 25th/75th percentile (box), the mean (dashed line), the

median (solid line) and the 5th/95th percentile (error bars). Abbreviations: EUB (EUB338),

CFB (CFB319a), ALF (ALF968), GAM (GAM42a), BET (BET42a), ROS (ROS536), ARCH

(ARCH915), SRB (SRB385), SRBdb (SRB385db), NON (NON338).

Fig. 3. DGGE fingerprints of sediment surface (SE), particle-attached (PA), and free-living

(FL) bacterial communities of the Wadden Sea during tidal cycles in November 1999 (Nov)

and May 2000 using primer sets for 16S rRNA genes of Bacteria. The arrows mark excised

and sequenced bands. Bands sequenced in this study include “TC” for tidal cycle, bands

named GWS-eX- PA/FL refer to sequences of Rink et al. (2006). Std. = standard.

Fig. 4. DGGE fingerprints of free-living (FL) and particle-attached (PA) bacterial

communities during a tidal cycle in July 2005, using primer sets for 16S rRNA genes of

Bacteria (EUB), Bacteroidetes (CFB), alpha-Proteobacteria (ALF) and the Roseobacter

clade (ROS). “DNA” marks banding patterns obtained from 16S rRNA gene fragments and

“cDNA” refers to banding patterns derived from 16S rRNA after reverse transcription. HT =

high tide, MT1/2 = mean tide, LT = low tide. The numbered arrows mark excised and

sequenced bands. Because of the small fragment size of the Roseobacter amplicons (approx.

200 bp) the DGGE bands were not excised for sequencing. Std. = standard.

Fig. 5. Cluster analyses of DGGE banding patterns (July 2005, Fig. 4) of particle-attached

(PA) and free-living (FL) Bacteria (EUB), Bacteroidetes (CFB), alpha-Proteobacteria (ALF)

and the Roseobacter clade (ROS). “DNA” marks banding patterns obtained from 16S rRNA


71

gene fragments and “cDNA” refers to banding patterns derived from 16S rRNA after reverse

transcription. Samples were taken during a tidal cycle at high tide (HT), mean tide (MT1/2)

and low tide (LT). The similarity matrix was calculated using UPGMA and Pearson

correlation. Std. = standard.

Fig. 6. Phylogenetic trees of Proteobacteria and Actinobacteria (A) and Bacteroidetes (B)

calculated with Maximum-Likelihood based on 16S rRNA gene fragments. Sequences

obtained in this and one earlier study partially including the same samples [Rink et al. (2006),

compare Fig. 3] are highlighted in bold. Alpha, delta, gamma and beta = different

phylogenetic groups of Proteobacteria, WAC I = Wadden Sea Alpha Cluster (Stevens et al.

2005), RCA = Roseobacter Clade Affiliated (Selje et al. 2004), SAMMIC = Surface Attached

Marine MICrobes (Stevens et al. 2005). “GWS” (German Wadden Sea) indicates clones

obtained from the same habitat in this and earlier studies. The percentage of sequence

divergence is indicated by the scale bars.

Supplementary Data

Fig. I. Particulate organic carbon (POC), ratio of POC per dry weight (DW) and density of

particle-attached bacteria at high tide (HT), mean tide (MT) and low tide (LT) in July 2005.

Fig. II. DGGE fingerprints of sediment surface, particle-attached, and free-living bacterial

communities of the Wadden Sea during tidal cycles in November 1999 and May 2000 using

primer sets for 16S rRNA genes of Bacteria.


72

Table 1. FISH counts of November 1999, May 2000 and July 2005 during tidal cycles taken

at high tide (HT), mean tide (MT) and low tide (LT) in percent. Results of July 2005

were obtained using CARD-FISH. n = number of analysed subsamples, std dev =

standard deviation

Table. 1. Rink et al.


73

Fig. 1. Rink et al.


74

Fig. 2. Rink et al.

Fig. 3. Rink et al.

GWS-TC-e5-

GWS-e3-

GWS-e6-

GWS-e7-

GWS-e5-

GWS-e8-

GWS-e13-

GWS-TC-e4-

GWS-e12-

GWS-TC-e3-

GWS-TC-e1-

GWS-TC-e2-

No Ma No Ma No MaSt St

SE PA FL


75

EUB

H MT

L MT

H MT

L MT

H MT

L MT

H MT

L MT

PA FL PA FL

DNA cDNA

CFB

ALF

ROS

Std. Std.

GWS-TC-e6-GWS-TC-e7-

GWS-TC-e8-GWS-TC-e9-

GWS-TC-e10-

GWS-TC-e11-

GWS-TC-c1-

GWS-TC-c2-GWS-TC-c3- GWS-TC-c4-

GWS-TC-a1-

GWS-TC-a2-GWS-TC-a3-

GWS-TC-a4-

GWS-TC-a5-

GWS-TC-a6-

Fig. 4. Rink et al.


76

Fig. 5. Rink et al.

EUB

CFB

ALF

ROS

Pearson correlation [0.0%-100.0%]


77


alpha

delta

gamma

Actinobacteria

WAC I

RCA

SAMMIC

beta

A

Roseobacter


78

Fig. 6B. Rink et al.

B


79

Fig. I. Rink et al.

Supplementary Data


80

Fig. II. Rink et al.

81

IV.

Diversity and abundance of Gram-positive bacteria

in a tidal flat ecosystem

Kapitel IV Diversity and abundance of Gram-positive bacteria

82

Diversity and abundance of Gram-positive bacteria in a tidal flat ecosystem

Heike Stevens, Thorsten Brinkhoff, Beate Rink, John Vollmers, and Meinhard Simon*


University of Oldenburg, Germany

Running title: Gram-positive bacteria in tidal flats

Key words: Gram-positive bacteria, Actinobacteria, Firmicutes, tidal flats, dilution cultures,

fluorescence in situ hybridization, DGGE

___________________________________________________________________________

*Corresponding author

Mailing address: ICBM, University of Oldenburg, PO Box 2503, D-26111 Oldenburg,

Germany. Phone: +49 (0) 441 / 798-5361. Fax: +49 (0) 441 / 798-3438. E-mail:

[email protected]


83

Abstract.

Gram-positive (Gram+) bacteria recently have been identified as important components of

freshwater ecosystems and are also present in marine environments. However, their

quantitative significance and possible role in the latter systems is still little studied, in

particular in coastal regions. Therefore, we investigated the abundance and composition of

Gram+ bacteria in the Wadden Sea, a tidal flat ecosystem in the German Bight of the North

Sea. Applying fluorescence in situ hybridization we found that Actinobacteria constitute 4-7%

of total bacteria in the Wadden Sea and slightly higher proportions in a freshwater drainage

channel connected to the sea by a sluice. The application of DGGE of 16S rRNA gene

fragments after amplification by an Actinobacteria-specific primer set and subsequent

sequencing showed that the composition of the actinobacterial community in the Wadden Sea

was distinctly different from that in the freshwater system. A clone library of 103 clones

yielded 8 Gram+ phylotypes which are related closely to other marine phylotypes including the

Marine Actinobacteria Clade but also to freshwater phylotypes. We applied dilution cultures,

enriched with various biopolymers for isolating bacteria from the bulk water, suspended

aggregates, the oxic surface and oxic/anoxic transition zone of the sediment, Marine Broth and

Fucus vesiculosus extracts. Fifty three isolates affiliated to seven families of the order

Actinomycetales and 9 isolates to the family Bacillaceae. The salinity range (1 to 45‰ NaCl)

and growth optimum of fourteen strains from various families showed that all except one strain

exhibited a rather broad range of sustained growth from 1 to >20‰ NaCl and several strains

exhibited an optimum of >10‰ NaCl. The results indicate that the Gram+ bacterial community

in the Wadden Sea is surprisingly diverse and consists mainly of indigenous species which

appear to be well adapted to the environmental conditions of this coastal ecosystem.


84

Introduction

In the recent past, more and more evidence accumulated which indicated that Gram-positive

(Gram+) bacteria, in particular Actinobacteria, are of hitherto unknown significance in aquatic

ecosystems (Bull et al., 2005). Studies in freshwater systems showed a surprising diversity and

abundance of Actinobacteria even though the specific ecological role remains to be unveiled

(Glöckner et al., 2000; Hahn et al., 2003; Stepanauskas et al., 2003; Warnecke et al., 2004;

Haukka et al., 2005; Allgaier and Grossart, 2006). It has been known for more than 60 years

that Gram+ bacteria also exist in marine environments, but they were thought not to be

indigenous but introduced from terrestrial habitats (Zobell and Upham., 1944; Goodfellow and

Haynes, 1984). Since the mid-1990s, studies using culture-independent but also refined

culture-dependent methods indicated that Gram+ bacteria reveal an unexpected diversity in

marine bacterioplankton communities (Fuhrman et al., 1993; Rappé et al., 1997; Suzuki et al.,

1997; Rappé et al., 1999; Fuchs et al., 2005) as well as in marine sediments and a surprisingly

high abundance of up to 13% (Jensen and Fenical, 1995; Moran et al., 1995; Urakawa et al.,

1999; Mincer et al., 2002; Maldonado et al., 2005; Pathom-aree et al., 2006). Phylogenetic

analyses of the phylotypes and isolates obtained in these studies implied that, in fact,

indigenous marine Gram+ bacteria exist. Some of the Gram+ bacteria obtained from marine

habitats fall into distinct “marine” clusters, only distantly related to clusters comprising also

Gram+ bacteria from freshwater and soil. The Marine Actinobacteria Clade, (Rappé et al.,

1999), which includes the “BD1-5 cluster” (Fuhrman et al., 1993), a deeply branching cluster

within the Actinobacteria, comprises exclusively marine bacterioplankton phylotypes. The

MAR 1 cluster (Mincer et al., 2002) consists of Actinobacteria isolates from tropical and

subtropical marine sediments, but is branching not as deeply as the Marine Actinobacteria

Clade.

Quantitative studies on marine Gram+ bacteria are still scarce. A biomarker study on the basis

of the composition of phospholipid ester-linked fatty acids (PLFA) indicated that Gram+

bacteria are major components of bacterial communities in sediments of a eutrophic bay

(Rajendran et al., 1994). Actinobacteria constituted up to 5% of total bacteria in shallow

marine sediments and <1.4% in an Arctic deep-sea sediment as determined by dot blot and

fluorescence in situ hybridization (Moran et al., 1995; Llobet-Brossa et al., 1998; Ravenschlag

et al., 1999). In the Sargasso Sea and the Arabian Sea, Actinobacteria have been identified as

substantial components of the bacterioplankton (Fuchs et al., 2005; Morris et al., 2005). In the

Delaware estuary, USA, Actinobacteria constitute a decreasing fraction of total

bacterioplankton numbers and of the proportions assimilating glucose and polysaccharide with


85

increasing salinity (Elifantz et al., 2005; Kirchman et al., 2005). It is of particular interest to

reveal the significance, composition and abundance of Gram+ bacterial communities in coastal

tidally affected ecosystems and whether in these systems indigenous Gram+ bacteria do exist

These systems are severely understudied with respect to Gram+ bacteria as compared to other

marine systems (Bull et al., 2005).

The Wadden Sea is a shallow and nutrient-rich tidally affected coastal ecosystem of the

southern North Sea stretching from the Netherlands (Den Helder) to Denmark (Esbjerg). Due

to the pronounced tidal dynamics and inputs of organic and inorganic nutrients from land,

freshwaters, and the North Sea it can be considered as a melting pot in which microbial

processes are of major significance (Dittmann, 1999; Poremba et al., 1999). In the recent past

extensive work on microbial processes and on the composition of bacterial communities in the

bulk water, on suspended aggregates and in the sediment of the Wadden Sea has been carried

out (e.g. Llobet-Brossa et al., 1998; Köpke et al., 2005; Stevens et al., 2005a, 2005b; Lunau et

al., 2006; Wilms et al., 2006).

This study investigated the composition and abundance of the Gram+ bacterial community in

the Wadden Sea. We applied cultivation-based approaches, using enrichment and dilution

cultures amended with a variety of biopolymers, as well as cultivation-independent approaches

using Gram+-specific CARD-FISH (CAtalyzed Reporter Deposition Fluorescence In Situ

Hybridization, Sekar et al., 2003), DGGE (Denaturing Gradient Gel Electrophoresis, Muyzer

et al., 1993) and clone library construction. The salt requirements for growth of representative

isolates were examined by determining their growth adaptation to the ambient salinity range.

RESULTS

Isolation

Seventeen of 63 bulk water isolates (27%) obtained from dilution cultures of the May sample

were Gram+ strains. Isolates affiliating to α- and γ-Proteobacteria constituted proportions of

44.4% and 20.6%, respectively, and those affiliating to Bacteroidetes 7.9%. In October, the

majority of strains affiliated to α -Proteobacteria (31.0%), Actinobacteria (30.2%) and γ -

Proteobacteria (24.0%). Including the Firmicutes isolates (n=6) 45 of the 129 October strains

(34.9%) were Gram+ bacteria, of which 35.6% originated from the bulk water, 20% from

aggregates and the transition zone each and 24.4% from the oxic layer of the sediment. In

other studies of which Gram+ isolates were included in the phylogenetic analysis, only the


86

bulk water (Bruns et al., 2003; Selje et al., 2005) or aggregates (Grossart et al., 2004) were

investigated.

Gram+ isolates were obtained with all substrates used. The only isolate obtained with

laminarin also affiliated to Gram+ bacteria. The highest MPN dilution steps which yielded

isolates, was 10-8. These isolates were derived from bulk water in May with casein and from

the sediment transition zone in October with MB as substrate and affiliated to Actinobacteria

and Firmicutes, respectively. Overall most Gram+ isolates obtained in May and October were

isolated with alginate (n=11), followed by agar (n=9), MB and starch (n=8). Only few isolates

were retrieved from dilution cultures amended with casein (n=2) and palmitate (n=4).

Dilution steps 10–1, 10–2 and 10–4 yielded >10 and the others not more than 7 Gram+ isolates

(Fig. 2).

Salinity-dependent growth

All strains except one exhibited a rather broad salinity range of sustained growth and 9 strains

grew equally well from 1 to >20‰ NaCl (Table 1) and thus appeared well adapted to the

salinities occurring in the Wadden Sea. Two strains grew significantly better at salinity ranges

>10‰ (GWS-BW-H260, GWS-BW-H252). One strain, affiliating to Bacilli, grew

significantly better at salinities <5‰.

CARD-FISH

Hybridization efficiencies varied from 48.9 to 67.7% as indicated by the ratio of numbers of

EUB338 positive over DAPI cell counts and subtracting the non-EUB338 numbers which

remained below 1.4% (Table 2). Gram+ bacteria were detected at all five locations and

proportions ranged from 4.2 to 8.0% of DAPI cell counts (Table 2). The highest proportion

occurred in the freshwater location behind the sluice.

DGGE analysis

The banding patterns of the Actinobacteria–specific DGGE revealed distinct communities of

the freshwater and the marine locations (Fig. 3). The marine samples were rather similar with

one prominent band occurring in all samples. However, the particle-associated and free-living

bacterial fractions exhibited also different bands. Band numbers ranged between 4 in the

particle-associated bacterial fraction of station 4 and 9 in the free-living bacterial fraction of


87

station 2. Sequencing of the excised bands yielded only actinobacterial 16S rRNA phylotypes,

indicating the specificity of the primers.

Clone library

Altogether 103 different clones were sequenced . Two of them were identified as chimera.

Eight of the remaining clones (7.8%) affiliated to Gram+ bacteria and the majority to γ- and

δ-Proteobacteria.

Phylogenetic affiliation

A total of 94 Gram+ 16S rRNA gene sequences was phylogenetically analyzed (Fig. 4A and

B, Tab. 3), 82 from isolates, 8 from the clone library and 4 from DGGE bands. To avoid an

overestimation of diversity, sequences with similarities of ≥99% were merged into 15

“sequence-groups” (I – XV) resulting in a total of 58 different sequence types (Tab. 3, Fig. 4).

Sixty-three sequences from isolates, 7 from clones and 4 from DGGE bands affiliated to

Actinobacteria. The sequences mainly clustered into seven families of the order

Actinomycetales, the Microbacteriaceae, Micrococcaceae, Mycobacteriaceae, Nocardiaceae,

Nocardioidaceae, Promicromonosporaceae, Pseudonocardiaceae, and the

Sanguibacteraceae. Most sequences group with the Microbacteriaceae (18 isolates, 1 clone),

with the Micrococcaceae (16 isolates) and with the Nocardioidaceae (15 isolates). Because of

their low sequence similarity to the next relative, Streptomyces cinnabarus and candidatus

“Microthrix parvicella”, clone K39 and DGGE bands GWS-DG2, GWS-DG1, GWS-FL-8,

GWS-DG3 could only be classified on the class-level (Table 3). The latter three phylotypes

clustered together with various phylotypes of uncultured marine Actinobacteria. Clone GWS-

K46 and isolates GP-5 and GP-6 belong to the order Actinobacteria with next related species

of the subclass Frankinae. Clones GWS-K72, GWS-K105 and GWS-K112 clustered within

the Marine Actinobacteria Clade.

Sixteen of the 58 Gram+ sequence types (28%) obtained from the Wadden Sea exhibit

a closest relative of marine origin, 37 of other environments, such as freshwater, soil, plants,

or endophytic habitats. For 5 sequence types no information is available on the source of

isolation of the next related sequence (Table 3). Our strains affiliating with the

Micrococcaceae were mainly isolated from bulk water, isolates affiliating with the

Nocardioidaceae from bulk water and aggregates. For all other sequences and isolates no


88

relationship exists between phylogeny and habitat or substrate from which they were

obtained.

Eight of our strains, 10 from other studies of the same habitat and one clone (K48, not

shown in Fig. 4B) affiliate to the phylum Firmicutes. Fifteen of the isolates were merged into

five sequence groups. All isolates affiliate to the order Bacillales and, except sequence group

XIV (GP-13, GP-14) and isolate GWS-TZ-H232, fall within the family Bacillaceae.

Sequence group XIV affiliates to the Planococcaceae. The clone K48 affiliates within the

class Clostridia to the “Peptostreptococcaceae” (family name not validly published), with the

next relative Fusibacter paucivorans (Table 3). Isolates affiliating to the Firmicutes were

never obtained from aggregates, but from the sediment layers and the bulk water. They were

obtained from assays performed with MB, agar, alginate, and stearine.

Discussion

Our results show that Gram+ bacteria constitute a substantial fraction of the bacterial

community in the Wadden Sea and that they are indigenous to this environment. They

comprise around 5% of total bacterial numbers and are a major component of its so far

cultivated fraction. However, the isolated strains were only distantly related to most

phylotypes retrieved from the clone library and from the Actinobacteria-specific DGGE

bands. Most of the Gram+ isolates tested grew at a rather wide salinity range including 20 and

30‰ NaCl, thus indicating that they are well adapted to growing in the Wadden Sea , in

which salinities of 26 to 33 psu occur (http://las.physik.uni-oldenburg.de/wattstation). Even

though it seems possible that Gram+ bacteria are also washed in from terrestrial run off and

by releasing drained fresh water through the dike sluices these bacteria do not appear to

constitute prominent members of the marine Gram+ bacterial community. The DGGE bands

of the freshwater sample were distinctly different from the marine ones and only one isolated

strain exhibited a distinct growth optimum at salinities <5‰. Further, several clones of the

clone library affiliated to the Marine Actinobacteria Clade (Rappé et al., 1999), and the

sequence of one prominent DGGE band detected at all marine stations (GWS-DG3) affiliated

to marine Actinobacteria phylotypes and most closely to a phylotype detected in the

northwestern coastal Pacific (Fig. 4; Table 3; Morris et al., 2006).

Gram+ bacteria comprised 27 and 35% of all isolates retrieved in May and October

1999, respectively, and the majority affiliated to Actinobacteria. Thus, Gram+ bacteria

constitute one of the two major cultivable phylogenetic classes of isolates in the Wadden Sea.


89

Because the Gram+ bacteria were obtained with various substrates we assume that the

frequent isolation of these organisms was not due to a cultivation bias but rather a result of the

various biopolymers used as enrichment substrates. Even though most of the isolates were

retrieved from low dilution steps, several isolates from various habitats were obtained from

10-6 to 10-8 dilutions, suggesting that some of them are significant constituents of the bacterial

community in this ecosystem. This is particularly true for sequence groups I

(Microbacteriaceae) and X (Nocardioidaceae). Three of the strains within group I were

obtained from 10-6 or higher, and 6 of the 7 strains in group X from 10-4 or higher (Fig. 4).

This suggestion is supported by the finding that the recently described Aeromicrobium

marinum, isolated from a 10-4 dilution culture retrieved from the Wadden Sea and the first

described marine species within the family Nocardioidaceae, constitutes up to 1% of total

bacteria in the water column (Bruns et al., 2003).

Several other recent studies also isolated Actinobacteria from various marine

environments, but in most cases they constituted lower fractions of all isolates as compared

with our results. In studies from various regions including the German Bight of the North Sea,

the Baltic Sea and the Oregon coast of the Pacific, which examined the diversity of isolates

from bacterioplankton samples and enrichments with various substrates, between <1 and 15%

of all isolates affiliated to Actinobacteria (Suzuki et al., 1997; Eilers et al., 2000; Hagström et

al., 2000; Uphoff et al., 2001). Du et al. (2006) reported that Gram+ bacteria were a

prominent fraction of the pigmented cultivable bacterial community in Chinese estuaries and

were also present in the coastal sea, even though to much lower proportions. In various

marine sediments, ranging from shallow coastal regions to the deepest ocean, Gram+ bacteria

of a rather wide diversity have been isolated in recent studies (Mincer et al. 2002; Köpke et

al., 2005 ; Maldonado et al., 2005; Pathom-aree et al., 2006). These notions indicate that

Gram+ bacteria constitute a greater fraction of the so far cultivable proportion of the bacterial

community in coastal environments than previously assumed.

The fraction of Actinobacteria we assessed by CARD-FISH is in the same range as

that reported from the Delaware Bay, USA (Kirchman et al., 2005). These authors found that

Actinobacteria constitute decreasing proportions of 20-30% of total bacterial numbers to 5%

from the freshwater end of the estuary to the marine Bay. This study did not examine the

phylogenetic composition of the Gram+ bacterial community in the Bay. According to our

DGGE and sequencing results, however, it appears most likely that the composition of the

Gram+ bacterial community in the Bay was different from the freshwater section and that the

decreasing proportion of this phylogenetic class was not only a dilution effect. Our results


90

further imply that an indigenous marine actinobacterial community exists, also in such near

shore environments with intense water exchange with estuarine and freshwater habitats. The

proportions of Actinobacteria we and Kirchman et al. (2005) found in the water column of the

Wadden Sea and the Delaware Bay are somewhat higher than proportions reported from the

upper 4 cm of Wadden Sea sediments, <1 to 3.6% (Llobet-Brossa et al., 1998) and also as

those of the genus Streptomyces detected in a shallow marine sediment by dot blot

hybridization with a genus-specific probe (2.0 to 5.1% of total extracted rRNA; Moran et al.,

1995). Studies based on PLFA indicate that Gram+ bacteria are significant components of the

bacterial communities in eutrophic bays in Japan (Rajendran et al., 1994; Rajendran and

Nagatomo, 1999). In open ocean environments the significance of Actinobacteria appears to

be variable. Whereas in the Sargasso Sea, Actinobacteria constitute <2% of total bacterial

numbers (Morris et al., 2005), in the Arabian Sea variable proportions of 2 and 13% of total

bacteria were found in oligotrophic waters and in the oxygen minimum zone, respectively

(Fuchs et al., 2005). Hence, these data indicate that Actinobacteria are a prominent

component of the bacterial community in shallow coastal ecosystems, in the water column as

well as in the sediment, even though its quantitative proportion is lower than that of other

important classes such as - and -Proteobacteria and Bacteroidetes (Kirchman et al., 2005;

Llobet-Brossa et al., 1998) and lower than that of Actinobacteria in freshwater ecosystems

(Glöckner et al., 2000; Allgaier and Grossart, 2006).

Our phylogenetic analysis of sequences from the isolates, the DGGE bands and clone

library shows a surprisingly high diversity of the Gram+ bacteria. The sequences from the

DGGE bands and the clone library affiliated to distinctly different groups than the isolates of

this phylogenetic lineage. The diversity within Actinobacteria we detected was greater than

that described in other studies using either culture-dependent (Jensen and Fenical, 1995;

Suzuki et al., 1997; Köpke et al., 2005; Maldonado et al., 2005; Du et al., 2006; Pathom-aree

et al., 2006) or culture–independent approaches (Fuhrman et al., 1993; Gray and Herwig,

1996; Suzuki et al., 1997; Rappé et al., 1999; Urakawa et al., 1999; Wilms et al., 2006). This

may be due to the various isolation procedures we applied, such as dilution series and

different substrates and to the various habitats we sampled, but may also reflect the specific

signature of the Wadden Sea ecosystem as a melting pot with marine as well as terrestrial

impacts.

The fact that we successfully enriched and isolated Actinobacteria from various

habitats of the Wadden Sea with various biopolymers shows that these strains are capable of

degrading a variety of polymeric substances. Some of these substances are typical for coastal


91

marine environments such as F. vesiculosus, cellulose, starch, chitin, and laminarin.

Actinobacteria are well known to be capable to degrade various polymeric substances such as

cellulose and lignin, but also rubber and polyester (Haider et al., 1978; Godden and

Penninckx, 1984; Jendrossek, 1997; Pranamuda et al., 1999; Linos et al., 2002). Their

hydrolytic potential appears comparable or even greater than that of the Sphingobacteria and

Flavobacteria group of the Bacteroidetes phylum (Reichenbach, 1992; Kirchman, 2002). It

may explain why Actinobacteria prosper in the bulk water and sediment of the Wadden Sea

and other coastal environments which are characterized by high concentrations of various

biopolymers (Harvey and Mannino, 2001). In fact, Piza et al. (2004) found a surprising

diversity of Actinobacteria in a Brazilian estuary subjected to high pollution during the last

fifty years, and in particular at its brackish end. This high diversity obviously reflects the high

potential of Actinobacteria to degrade complex organic substances including recalcitrant

compounds and pollutants.

In contrast to sequences affiliating to Actinobacteria, our sequences affiliating to

Firmicutes were much less divers. Sequences of all bacterial isolates of this group clustered

within the class Bacilli. The only clone obtained from the Firmicutes belongs to the class

Clostridia. Firmicutes appear to be more prominent members of the subsurface sediment,

possibly because of their ability to produce endospores. In a recent study a surprisingly high

diversity of Firmicutes was found in subsurface sediments of the German Wadden Sea

(Köpke et al., 2005). Since the early 1970s various Bacilli were isolated from marine habitats

(e.g., Denis, 1971; Bonde, 1976; Stackebrandt et al., 1997; Urakawa et al., 1999; Siefert et al.,

2000), but only very few marine isolates affiliated to the classes Lactobacillales, Clostridia,

and Mollicutes (Finne and Matches, 1974; Timmis et al., 1974; Marty, 1986; Gray and

Herwig, 1996). Hence, Bacilli seem to be the most abundant marine Firmicutes.

A further feature, explaining the high diversity of Gram+ bacteria in coastal

ecosystems may be their ability to grow at wide salinity ranges. From the 14 strains whose

growth adaptation to salinity we tested, all except one were able to grow at salinities >5‰

NaCl and exhibited growth optima ranging to 20‰ or higher (Table 1). Four of the 14 isolates

had as closest relative a strain also isolated from a marine habitat (GWS-BW-H301M, GWS-

AG-H268, GWS-SE-H117, GWS-TZ-H232, Table 3) and two of the latter isolates grew only

at salinities >5‰ NaCl. The fact that the next relatives of the other isolates were not of marine

origin is no indication that our isolates growing at higher salinity ranges were not truly marine

strains. Aeromicrobium marinum, the recently described marine species within the family

Nocardioidaceae, clusters with terrestrial isolates but exhibits a requirement for seawater


92

typical of marine bacteria (Bruns et al., 2003). Hence, these marine species clustering closely

together with other non-marine species may indicate that they were introduced into coastal

habitats from adjacent soil and freshwater habitats. This notion underlines the significance of

the close interactions of the land sea transition zone for evolutionary processes and may

further explain the great diversity of Gram+ bacteria occurring in estuarine and coastal

environments.

Four of the five phylotypes we obtained from the clone library and the DGGE-bands

of the marine samples clustered closely together with other phylotypes of exclusively marine

origin and only distantly related to phylotypes or isolates of freshwater or soil origin (Fig. 4).

Because none of the isolates appeared as a prominent DGGE band with the Actinobacteria-

specific PCR, the in situ dominant components of the Gram+ bacterial community

presumably constituted of these yet uncultured phylotypes to a great extent. Obviously the

community of Gram+ bacteria in coastal environments consists of two fractions, one

including the cultivated strains and one including only not-yet cultivated phylotypes, mainly

affiliated to the Marine Actinobacteria Clade (Rappé et al., 1999), and whose physiological

traits are basically unknown. Whereas the indigenous community appears to be most

important for the turnover of organic matter at ambient conditions, the cultivable fraction

appears as a valuable resource for isolates capable of interesting catalytic pathways and for

bioactive compounds (Grossart et al., 2004; Bull et al., 2005; Maldonado et al., 2005).

Our analysis of the community of Gram+ bacteria in the Wadden Sea shows that it

was surprisingly divers, mainly consisting of various groups of Actinobacteria, and to a much

lesser extent of Firmicutes. Phylotypes and isolates clustered to distinctly different groups.

The phylogenetic affiliation of the phylotypes and the broad salinity range of most of the

isolates indicates that the Gram+ bacterial community in the Wadden Sea is well adapted and

indigenous to the marine environment. On the basis of the CARD-FISH results we estimate

that Gram+ bacteria constitute around 5% of total bacteria in the water column of the Wadden

Sea. Hence, they appear to be prominent members of the bacterial communities and, because

of their high potential to degrade various biopolymers, are important in the turnover and

decomposition of organic matter in this ecosystem.


93

Experimental procedures

Sampling

Samples were collected on 27 May and 25 October 1999 in the East Frisian Wadden Sea,

Germany (station A, 53° 37´ N, 07° 08´ E; station B 53° 42´ N, 07° 43´ E; Fig 1). Water

samples were taken at high tide with pre-rinsed 10 l-plastic jugs. Sediment cores from an

intertidal mixed sand/mud flat were taken only in October with Plexiglas tubes (36 mm

diameter) at low tide. Samples were brought to the lab on ice in cooling boxes and processed

further within 2 h. Water samples for CARD-FISH and DGGE analysis were taken on 17

August 2005 at five sampling points along a salinity gradient from 0.3 to 31 psu close to

station B (Fig. 1) and processed further within 3 hours. For DGGE analysis, 100 ml of water

were filtered onto 5.0 µm polycarbonate filters (Nuclepore, Whatman) to obtain particle-

associated bacteria and subsequently onto 0.2 µm polycarbonate filters to obtain free-living

bacteria. Filters were stored at -20°C until nucleic acid extraction.

Isolation of bacteria

For the October samples we applied the MPN (most probable number) technique (Trolldenier,

1993) with different substrates and subsequent isolation of bacteria (Stevens et al., 2005b).

For the May samples dilution series were used for isolation of bacteria. Therefore, 1 ml of

bulk water was used as inoculum for 10-fold dilution series. Mineral media amended with

various substrates and MB 2216 (Difco, Germany) were prepared as described previously

(Stevens et al., 2005b). The following substrates were added (0.1%): agar, alginate, casein,

cellulose, chitin, laminarin, dried and pestiled Fucus vesiculosus (a brown algae growing

copiously along the coast line of the Wadden Sea), palmitate, starch and stearate. Growth was

checked by turbidity and microscopically. Bacteria were isolated from various dilution steps

on agar plates containing the same media as in the MPN assays amended with 1.5% agar. For

further cultivation Marine Agar 2216 (Difco, Germany) was used. Additionally, three isolates

were obtained from 1 l-rolling tanks filled with natural seawater and amended with 0.1% agar

and alginate, respectively, and incubated for 100 days at 15 °C in the dark. Single colonies

were transferred at least five times until considered as pure. The purity of the isolated strains

was examined by DGGE analysis according to Brinkhoff and Muyzer (1997). Isolates from

the same habitat and from assays with the same substrate with sequence similarities of ≥99%

[as determined by a similarity matrix calculated with ARB (Ludwig et al., 2004)] were


94

considered identical and only one sequence, either from the highest dilution step or from the

highest number of sequenced bp, was used in this study and submitted to GenBank.

Salinity dependent growth

In order to examine growth adaptation to the ambient salinity range 14 selected isolates from

the major families of Gram+ bacteria (Table 1) were grown on MB agar plates and transferred

to liquid MB medium of a salinity of 20‰ NaCl (w/v). These cultures were used to inoculate

5 ml liquid MB cultures in triplicate test tubes of salinities of 1, 5, 10, 20, 30 and 45‰ NaCl.

Test tubes were incubated in the dark at 20°C and growth was monitored over 4 to 10 days by

examining periodically the optical density at 660 nm spectrophotometrically directly in the

test tubes. Growth rates were determined as the slope of the exponential growth phase of the

log-plotted growth curves.

Fluorescence in situ hybridization (CARD-FISH)

CARD-FISH analyses were carried out basically following the protocol of Pernthaler et al.

(2004). Four replicates of one ml of each sample were filtered onto a 0.2 µm polycarbonate

filter (25 mm diameter, Nuclepore, Whatman) and fixed with 3 ml of paraformaldehyde (4%

w/v) for one hour. Subsequently, filters were embedded in low-gelling point agarose (0.2%

w/v, Metaphor) and incubated in lysozyme (10 mg ml-1) and achromopeptidase (2 U µl-1) as

described by Sekar et al. (2003). Cells were hybridized with the probes HRP-EUB338

(Amann et al., 1990), HRP-HGC69a (Roller et al., 1994) and HRP-NON338 (Wallner et al.,

1993) for 2 hours at 35°C. Amplification was performed at 37°C using FITC-labeled

Tyramide (fluorescein-5-isothiocyanate, Invitrogen) for 30 min in the dark. Samples were

incubated in 1x PBS (pH 7.3) amended with Triton X-100 (0.05%) for 30 min at room

temperature in the dark to remove residual dye. Cells were counterstained using Vectashield

with DAPI (4´,6´-diamidino-2-phenylindol, 1.5 µg ml-1; Vector Laboratories, Peterborough,

UK) and kept frozen at -20°C until further processing. Enumeration was performed by

epifluorescence microscopy (Axiolab, Zeiss, Germany) at 1,000 x magnification.

Nucleic acid extraction

Genomic DNA was extracted after bead-beating by phenol-chloroform as described earlier

(Stevens et al., 2005a). Precipitation of nucleic acids was done overnight at -20°C using

isopropanol. DNA pellets were resuspended in molecular grade water (Eppendorf, Hamburg,

Germany) and stored at -20°C until further processing.


95

PCR amplification of 16S rRNA gene fragments

PCR amplifications were performed with an Eppendorf Mastercycler (Eppendorf) as

described previously (Brinkhoff and Muyzer, 1997). 16S rRNA gene fragments of

Actinobacteria were amplified using the primers S-C-Act-235-a-S-20

(CGCGGCCTATCAGCTTGTTG, forward) and S-C-Act-878-a-A-19

(CCGTACTCCCCAGGCGGGG, reverse, Stach et al., 2003). A GC-clamp was added to the

forward-primer for subsequent DGGE analysis (Muyzer et al., 1998). Amplification was

performed according to Stach et al. (2003) with the following modifications: Denaturing and

annealing were extended from 45 to 60 s and elongation from 1 to 3 min. Touchdown from

72°C to 67°C in 0.5°C steps was done with two instead of one cycle per step and annealing at

68°C was extended from 15 to 20 cycles.

Four µl of the amplification products were analyzed by electrophoresis in 2% (w/v)

agarose gels and stained with ethidium bromide (1 µg ml-1) (Sambrook et al., 1989). For

subsequent sequencing analysis PCR products were purified by using the Qiaquick PCR

purification kit (Qiagen Inc.).

DGGE analysis of PCR products

DGGE was performed with the D-Code system (Bio-Rad Laboratories, Inc.). For the 16S

rRNA gene fragments of Actinobacteria, a gradient of 35 to 85% denaturant was used. After

electrophoresis, the gels were stained with SYBR Gold (Molecular Probes, Inc.) and

photographed using a BioDoc Analyze Transilluminator (Biometra, Göttingen, Germany).

Bands were excised with a scalpel sterilized with ethanol and transferred to sterile Eppendorf

caps. Fifty µl of water (molecular grade, Eppendorf, Germany) were added and the samples

were stored at –20°C.

Clone library construction

A water sample of 250 ml collected in October at station B was filtered onto a 0.2 µm

Nuclepore filter (47 mm diameter). The filter was immediately frozen at –80°C until DNA-

extraction. Bacterial genomic DNA of the sample was isolated after bead beating, phenol-

chloroform extraction, and isopropanol precipitation as described previously (Stahl et al.,

1988; MacGregor et al., 1997), but slightly modified. Lysozyme treatment was not applied,

precipitation done at –20°C and molecular grade water (Eppendorf, Hamburg, Germany)

was used for resuspension at 4°C over night. PCR amplification of almost complete 16S

rRNA gene fragments was performed as previously described (Brinkhoff and Muyzer, 1997)


96

with primers GM3F (8F) and GM4R (1492R) (Muyzer and Ramsing, 1995). Amplification

was done in triplicates and the products were pooled prior to cloning. For cloning the

pGEM®-T Vector System II (Promega, Madison, USA) was used according to the

manufacturer’s instructions. Sequencing of the clones was performed as described previously

(Stevens et al., 2005a). Clone sequences were checked for chimera formation with the

CHECK_CHIMERA software of the Ribosomal Database Project II (Maidak et al., 2001).

Sequencing and phylogenetic analysis

PCR amplification of 16S rRNA gene fragments of bacterial isolates and subsequent

sequencing was performed as described before (Brinkhoff and Muyzer, 1997). Sequences

were compared with similar sequences of reference organisms by BLAST search

(http://www.ncbi.nlm.nih.gov/blast (Altschul et al., 1998). Phylogenetic analysis was

performed with the ARB software package [http://www.arb-home.de (Ludwig et al., 2004)].

For tree calculation, only sequences with more than 1300 bp were considered using

maximum-likelihood analysis. Shorter sequences were added later to the final tree using the

maximum parsimony option of the ARB program. Alignment positions at which less than

50% of sequences of the entire set of data had the same residues were excluded from the

calculations to prevent uncertain alignments within highly variable positions of the 16S rRNA

gene fragments, which cause mistakes in tree topology (Ludwig et al., 2004). A phylogenetic

analysis of Gram+ bacteria of the Wadden Sea included sequences of the Gram+ isolates

obtained in May and October 1999 (this study), of a clone library and Actinobacteria-specific

DGGE bands (this study), and of previous studies of the water column of the Wadden Sea

using DGGE analysis or cultivation-based methods (Bruns et al., 2003; Brinkhoff et al., 2004;

Grossart et al., 2004; Selje et al., 2005; Stevens et al., 2005a). Sequences obtained from

isolates and clones in this study are available from GenBank under accession no. given in

Table 4.


97

ACKNOWLEDGEMENTS

We are grateful to Andrea Schlingloff for the sequencing, and to H.P. Grossart for the

introduction into the rolling tank incubation method. This work was supported by grants from

the Volkswagen Foundation within the Lower Saxonian priority Program Marine

Biotechnology and by the Deutsche Forschungsgemeinschaft within the research group

BioGeoChemistry of the Wadden Sea (FG-432, TP 5).


98

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Table 1: Strain ID, affiliation and salinity range of optimal growth of Gram positive bacteria tested. Growth was examined in a salinity range from 1 to 45‰ NaCl (w/v). For more details on the strains see Table 3 and Fig 3.

Strain Affiliation Salinity range (‰ NaCl)

GWS-BW-H60M Microbacteriaceae Gr I 1-30

GWS-BW-H301M Microbacteriaceae Gr II 5-30

GWS-AG-H268 Microbacteriaceae Gr II 5-30

GWS-BW-H45M Micrococcaceae 1-20

GWS-BW-H231 Micrococcaceae Gr V 1-30

GWS-BW-H260 Micrococcaceae Gr VI 10-20

GWS-BW-H82M Mycobacteriaceae 1-30

GWS-TZ-H135 Nocardiaceae 1-20

GWS-BW-H259 Nocardioidaceae 1-20

GWS-BW-H252 Nocardioidaceae 10-30

GWS-TZ-H118 Promicromonaspora citrea 1-45

GWS-BW-H222 Bacillus Gr XIII 1-5

GWS-SE-H117 Bacillaceae Gr XI 1-45

GWS-TZ-H232 Bacillus sp. 1-30


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Table 2: Cell numbers enumerated with the CARD-FISH probes EUB338 and HGC69a at five locations in the Wadden Sea near Neuharlingersiel. For exact locations see Fig. 1.

Station Salinity EUB338 HGC69a

(psu) (% DAPI cell numbers)

1 (fresh water) 0.3 56.3 + 9.3 8.0 + 1.3

2 29.1 67.7 + 7.5 4.2 + 0.9

3 31.0 48.9 + 7.4 4.7 + 1.4

4 30.8 59.1 + 10.3 5.1 + 1.4

5 30.5 62.3 + 7.7 6.5 + 2.1


105

Table 2: Clones and isolates obtained from the German Wadden Sea affiliating with Gram+ bacteria as well as their closest relatives determined by BLAST analysis (http://www.ncbi.nlm.nih.gov/blast). Sequences with a similarity ≥ 99% were grouped (sequence-groups I - XV), the given information pertains the longest obtained sequence of a sequence group. Given are phylogenetic affiliation, sequence / isolate ID and sequence group where applicable, summarized data on sequence or isolate, closest relative according to BLAST analysis and similarity of the 16S rRNA gene (%), and information concerning the closest relative. Remarks on the isolates give the MPN dilution steps (Dil. [10x]), habitats, substrates, and isolation dates. CAS= casein, CEL= cellulose; CHI= chitin, FUV= Fucus

vesiculosus; LAM= laminarin, PAL= palmitate; STA= starch, MB = Marine Broth 2216, MB* = Marine Broth 2216 prepared with natural sea water (Grossart et al., 2004). SW = autoclaved natural seawater amended with trace elements and vitamins (Selje et al., 2005), BW= bulk water, AG= aggregate, SE= oxic sediment; TZ= oxic/anoxic transition zone of the sediment; rt= rolling tank.

Phylum / family Sequence / isolate ID

(Representative of seq.- group)

Remarks on isolate / summary of sequence group

(Dil. [10x], habitat, substrate, date)

Closest relative (acc. number) [%] Habitat or environmental features of closest

relative

Actinobacteria

Microbacteriaceae GWS-SE-H242a -2, SE, PAL, Oct 99 Microbacterium sp. OS-6 (AJ296094)

99 coastal marsh (Galicia, Spain)

GWS-AG-H197 -3, AG, CHI, Oct 99 Microbacterium sp. V4.BP.11 (AJ244677)

98 marine bacterioplankton (Mediterranea)

GWS-TZ-H305 -2, TZ, FUV, Oct 99 Microbacterium esteraromaticum

(Y17231) 95 soil

GWS-TZ-H139 -1, ALG, TZ, Oct 99 Microbacterium testaceum SE034 (AF474327)

97 endophytic, agronomic crop

GWS-BW-H60M

(Sequence-group I)

-8, -7, -6, -1; BW, SE, TZ, AGA, CAS, CEL, STA; May, Oct 99

Microbacterium sp. VKM

Ac-2050 (AB042084)

99 plant nematode

GWS-SE-H300 -5, ALG, SE, Oct 99 Microbacterium sp. LB030 (AF474326)

99 endophytic, prairie plant

GWS-SE-H149 -2, CEL, SE, Oct 99 Gram+ bacterium strain 12-8 (AB008510)

99 copiotrophic, urban soil

GWS-BWrt-H97M

(Sequence-group II)

-1, rt; BW, AG, AGA, CAS, MB*, STA; May, Oct 99

Marine bacterium P_wp0234 (AY188942)

98 deep sea sediment/degrading PAH

GWS-SE-H243 -2, SE, PAL, Oct 99 Frigoribacterium faeni (Y18807) 98 psychrophilic, non-marine

Clone GWS- K13 From clone-library, BW, Oct 99

Actinobacterium MWH-Dar4 (AJ565416)

98 0.2 µm filtered freshwater

Sanguibacteraceae GWS-AG-H192 -3; AG, CHI, Oct 99 Cellulomonas fermentans (X79458) 94 municipal dumping site

Promicromonosporacea

e GWS-TZ-H118 -5, TZ, AGA, Oct 99 Cellulomonas sp. IFO16243

(AB023364) 96 no information available

Micrococcaceae GWS-BW-H45M

(Sequence-group III)

-5, -4, -1; BW, ALG, CEL, MB; May 99

Arthrobacter nicotianae SB42 (AJ315492)

97 starter culture (cheese)

GWS-SE-H161 -2, SE, CEL, Oct 99 Bacterium PS32 (AF200218) 99 psychrophilic, marine

GWS-BW-H126

(Sequence-group IV)

-4, -2; BW, SE; ALG, CEL, MB*; Oct 99

Bacterium isolate SI-12 (AJ252579)

99 agricultural soil

GWS-BW-H15M

(Sequence-group V)

-7, -2, -1; BW; CEL, FUV, MB; May, Oct 99

Micrococcus luteus strain Ballarat (AJ409096)

99 activated sludge

HP42 From aggregates Micrococcus sp. V4.MO.20 (AJ244665)


GWS-BWrt-H158

(Sequence-group VI)

-4, -2, rt; BW; CEL, STA; May, Oct 99

Kocuria rosea (Y11330) 99 soil and water.

Mycobacteriaceae GWS-BW-H82M -1, BW, MB, May 99 Mycobacterium sp. IP20010961 (AY163341)

99 water supplies

GWS-BW-H50M -1, BW, STA, May 99 Mycobacterium sp. TH-2003 (AY266138)

98 associated with sepsis


106

Table 3 cont. Phylum / family Sequence / isolate ID


Remarks on isolate / summary of sequence group (Dil. [10x], habitat, substrate, date)

Closest relative (acc. number) [%] Habitat or environmental features of closest relative

Nocardiaceae GWS-BWrt-H95M rt, AGA, May 99 Rhodococcus sp. UFZ-B520 (AF235011)

98 aquifer / degrading chlorobenzene

GWS-TZ-H135

(Sequence-group VII)

-4, -1; BW, TZ; ALG; Oct 99

Rhodococcus fascians KM6 (AJ011329)

100 humus (spruce stands)

GWS-SE-H175 -1, SE, CHI, Oct 99 Rhodococcus sp. MBIC01430 (AB088667)

99 no infomation available

GWS-TZ-H309

(Sequence-group VIII)

-6, -5, BW, TZ, FUV, Oct 99

Rhodococcus tukisamuensis (AB067734)

98 depolymerizing, from soil

Pseudonocardiaceae Pseudonocardiaceae bacterium T4

-1, BW, MB, Oct 99 Pseudonocardia alni IMSNU 20049 (AJ252823)

99 root nodules of alders

GWS-BW-H127

(Sequence-group IX)

-2,-1; AG, BW, AGA, ALG, Oct 99

Pseudonocardia alni IMSNU 20049 (AJ252823)

99 root nodules of alders

Nocardioidaceae GWS-BW-H99

(Sequence-group X)

-6, -5, -4, -1; AG, BW, SE; AGA, CHI, FUV, PAL, STA; Oct 99

Uncult. actinobacterium (AB074621)

97 aposymbiotic pea aphids

GWS-BW-H259 -4, BW, STA, Oct 99 Uncult. Nocardioides sp. GCPF40 (AY129808)

98 nutrient-limited cave

GWS-BW-H311M -1, BW, LAM, May 99 Nocardioides sp. MWH-CaK6 (AJ565419)

99 0.2 µm filtered freshwater

GWS-AG-H266 -4, AG, STA, Oct 99 Nocardioides sp. V4.BE.17 (AJ244657)


GP-1 -4, estuary: mar., Aug 99 Nocardioides OS4 (U61298) 98 oil shale column (oxic zone)

Aeromicrobium marinum -8, BW, MB, Oct 99 Aeromicrobium fastidiosum (Z7820)

97 herbage

GWS-BW-H252 -2, BW, PAL, Oct 99 Nocardioides sp. NCFB3005 (X76178)

97 No information available

GWS-BW-H89M -4, BW, ALG, May 99 Nocardioides sp. 2.20 (AJ299233) 98 freshwater biofilm

GWS-BW-H84M -4 BW, STA, May 99 Nocardioides jensenii KCTC 9134 (AF005006)

97 soil

uncertain actinomycetes GP-5 -7, estuary: brack.,

Aug 99

Unident. bacterium strain rJ7 (AB021325)

97 activated sludge (0.5 g phenol)

GP-6 -6, estuary: brack., Aug 99 Unident. bacterium strain rJ7 (AB021325)


Clone GWS-K46 From clone library, BW, Oct 99

Unident. bacterium strain rJ7 (AB021325)



Uncultured bacterium AT425_EubY10 (AY053479)

90 Gulf of Mexico gas hydrates


Unidentified bacterium clone K2-30-12 (AY344421)

98 Hawaiian archipelago

GWS-FL-8 DGGE band May-Aug 99 Uncultured actinobacterium clone SAa03 (AY124414)

99 Marine sediment


Uncultured actinomycete clone BD2-10 (AB015539)

95 Deep sea sediment


Uncultured bacterium clone E17 (AJ966591

95 Deep sea sediment


Uncultured actinomycete OCS155 (AF001652)

98 Coastal NW Pacific

Clone GWS-DG1 DGGE band, Aug 05 Uncultured actinobacterium clone PRD18H10 (AY948072)

99 Temperate river

Clone GWS-DG2 DGGE band, Aug 05 uncultured bacterium, clone AV9-158 (AM181875)

98 Subtropical lake

Clone GWS-DG3 DGGE band, Aug 05 Uncultured bacterium clone NH10_01 (DQ372838)

98 Oregon coast


107

Table 3 cont.

Phylum / family Sequence / isolate ID


Remarks on isolate / summary of sequence group

(Dil. [10x], habitat, substrate, date)

Closest relative (acc. number) [%] Habitat or environmental features of closest

relative

Firmicutes

Bacillaceae GWS-SE-H117

(Sequence-group XI)

-7; AG, SE; AGA, MB*; Oct 99

"Bacillus baekryungensis" (AF541965)

99 seawater (Korea)

HP 8 From aggregates, MB* Bacillus sp. KMM3737 (AY228462)

99 seawater (Korea)

GWS-BW-H68M

(Sequence-group XII)

-1; BW; MB*, STA; May 99

Bacillus pumilus OM-F6 (AB020208)


HP 10 From aggregates, MB* Bacterium KA64 (AY345445) 95 Hawaiian archipelago

GWS-BW-H220M

(Sequence-group XIII)

-6, -1; BW, MB; May, Oct 99

Bacillus licheniformis Mo1 (AF372616)

99 GTN degrading

GWS-TZ-H114 -2, TZ, AGA, Oct 99 Bacillus sp. HT-1 (AF463535) 96 hamster feces

Planococcaceae GP14

(Sequence-group XIV)

-5, estuary: marine, August 99

“Planococcus psychrotolerantus” (AF324659)


GWS-TZ-H232 -1, TZ, MB, Oct 99 "Planococcus rifitiensis" M8 (AJ493659)

99 mineral water in Italy

GWS-SE-H236

(Sequence-group XV)

-8, -4; SE, TZ; ALG, MB; Oct 99

Bacillus sp. Fa25 (AY131220) 99 strawberry plants

Peptostreptococcaceae

(Clostridia)

Clone GWS-K48 from clone library, BW, Oct 99

Fusibacter paucivorans (AF050099)

92 oil-producing well

Table 4: Accession numbers in GenBank of isolates and clones of this study.

Accession-no. Accession-no. Accession-no. Accession-no.

AY332093-AY332098 AY332125 AY332163 AY332202

AY332101 AY332129-AY332131 AY332164 AY332211

AY332104 AY332134 AY332170 AY332214

AY332105 AY332140 AY332173 AY332220

AY332108 AY332144 AY332183 AY332221

AY332111-AY332113 AY332146 AY332185 AY370612-AY370633

AY332118 AY332148 AY332193 EF088451-EF088496

AY332121 AY332149 AY332197

AY332122 AY332152-AY332154 AY332200


108

Figure legends

Fig. 1:

Locations of sample collection for the isolates (A, B), the clone library (B), and the DGGE

and CARD-FISH analyses (1-5) in the Wadden Sea (lower panel). Station 1 is a fresh water

drainage channel and stations 2-5 are marine. Between stations 1 and 2 is a sluice.

Fig. 2:

Numbers of Gram+ strains obtained in May and October 1999 from various dilution steps of

dilution cultures of bulk water samples (May and October), of suspended aggregates

(October), the sediment surface (October), and the oxic-anoxic transition zone in the sediment

(October).

Fig. 3:

Banding patterns of an Actinobacteria-specific DGGE analysis of free-living (FL) and

particle-associated bacterial communities (PA) collected at stations 1 to 5 in the vicinity of

station B in the Wadden Sea. For location of station B see Fig. 1. Arrows indicate bands

excised for sequencing.

Fig. 4:

Maximum likelihood trees of all Gram+ isolates and clones obtained from the Wadden Sea

(bold) showing the affiliation within the Actinobacteria (A) and the Firmicutes (B).

Sequences <1300 bp were added with maximum parsimony. The scale bars indicate 10%

sequence divergence. The Marine Actinobacteria Clade was adopted from Rappé et al. (1999)

and the MAR 1 cluster from Mincer et al. ( 2002). Isolates from May are marked with an "M"

at the end of the name. If available, dilution step and substrate were added to the accession

number (CAS = casein, CEL = cellulose; CHI = chitin, FUV = Fucus vesiculosus; LAM =

laminarin, PAL = palmitate; STA = starch, MB = Marine Broth 2216, MB* = Marine Broth

2216 prepared with natural sea water (Grossart et al., 2004), SW = autoclaved natural

seawater amended with trace elements and vitamins (Selje et al., 2005). The sub-habitat can

be derived from the name of the May and October isolates (BW = bulk water, AG =

aggregates, SE = sediment surface, TZ = oxic /anoxic transition zone of the sediment).

Numbers on branches with pooled sequences indicate the number of sequences used to

calculate the cluster.


109

FIG. 1. Stevens et al.

River Ems

River Weser

North Sea

N

10 km

A

B

2

53

4

1

2

53

4

1

50 m

5

2

3

4

1


110



Number of isolates

0 5 10 15

Dilu

tio

n s

tep

(lo

g 1

0)

-9

-8

-7

-6

-5

-4

-3

-2

-1

0

42531

PA FL PA FL PA FL PA FL PA FL

1

2

3

42531

PA FL PA FL PA FL PA FL PA FL

1

2

3


111

FIG. 4A, Stevens et al.


112

FIG. 4A cont., Stevens et al.


113

FIG. 4B, Stevens et al.

114

V.

High regional variability of bacterial communities

in the German Bight, North Sea

Kapitel V Regional variability of bacterial communities in the German Bight

115

High regional variability of bacterial communities

in the German Bight, North Sea

Beate Rink, Thorsten Brinkhoff, Katja Ziegelmüller, Meinhard Simon*


University of Oldenburg, D-26111 Oldenburg, Germany

Running title: Bacteria in the North Sea

Key words: North Sea, free-living and attached bacteria, Roseobacter, DGGE, phytoplankton

___________________________________________________________________________

* Corresponding author. Institute for Chemistry and Biology of the Marine Environment

(ICBM), University of Oldenburg, PO Box 2503, D-26111 Oldenburg, Germany,

Phone: +49-441-798-5361. Fax: +49-441-798-3438. E-mail: [email protected]


116

ABSTRACT

The German Bight of the North Sea is characterized by near shore tidal flat regions with high

loads of suspended matter and estuarine inputs of organic matter and pelagic off shore

regions. Due to tidal and wind-induced currents its hydrography is highly dynamic. In order to

examine how these highly dynamic properties affect the regional distribution and composition

of the bacterioplankton we conducted two surveys in June in two consecutive years during

which we assessed the composition of the free-living (FL, 0.2-5.0 µm fraction) and particle-

associated (PA, >5.0 µm fraction) bacterial communities on the background of hydrographic

(salinity, temperature) and biogeochemical properties (suspended matter, particulate organic

carbon, chlorophyll, phytoplankton composition). The composition of the bacterial

communities was determined by denaturing gradient gel electrophoretic (DGGE) analysis of

16S rRNA gene fragments PCR-amplified by Bacteria-, α-Proteobacteria and Bacteroidetes-

specific primer sets and subsequent sequencing of excised bands.

The results showed that the FL-bacterial community was rather evenly distributed in the

German Bight irrespective of the regional hydrographic and biogeochemical differences.

Several prominent bands, identified as phylotypes affiliated to the Roseobacter clade of α-

Proteobacteria, persisted throughout all 10 stations visited. The PA bacterial community

exhibited distinct differences among the various stations. These differences were not simply

attributed to properties of the near shore tidal flat regions and to the more homogeneous

hydrographic situation of the off shore region. They were rather site-specific, obviously

reflecting local conditions of the phytoplankton present and its growth phase and the

resuspended particles in the tidal flat regions. The results of the PA bacterial community

showed that unspecific PCR-amplifications were obtained by the Bacteria– (chloroplasts) and

α-Proteobacteria–specific primer sets (δ-Proteobacteria), biasing the results to a certain

extent. Because one primer applied for amplifying α-proteobacterial 16S rRNA gene

fragments (ALF968) is frequently used as a probe in fluorescence in situ hybridization (FISH)

analyses, its application leads to overestimates of α-Proteobacteria in samples containing δ-

Proteobacteria.


117

INTRODUCTION

In aquatic environments, complex communities of free-living and particle-associated

heterotrophic bacteria are the main decomposers of dissolved (DOC) and particulate organic

carbon (POC) and play a key role in the global carbon cycle (Cotner & Biddanda 2002). It has

been shown that origin and composition of organic matter affect the composition and biomass

production of bacterial communities (Covert & Moran 2001, Crump et al. 2003, Lebaron et al.

1999) and that phylogenetic bacterioplankton groups exhibit distinct preferences for low and

high molecular weight carbon sources (Cottrell & Kirchman 2000). Marine ecosystems are

structured into the open sea and coastal environments including the polyhaline estuarine

regions, and the environmental conditions in these habitats comprise different physical and

biogeochemical properties which may affect the ambient microbial communities. In the

coastal regions of the southern North Sea, the German Bight, high loads of dissolved and

particulate inorganic and organic matter are introduced from the tidal flats and the rivers

Weser and Elbe, thus providing organic and inorganic nutrients as well as refractory organic

matter (Loewe et al. 2005). An easterly current follows the southern coastal line along the

East Frisian Islands and the Weser estuary and encounters the polyhaline plume of the Elbe

estuary. Both water masses circulate along the north Frisian coast, often building separate

layers of distinct salinities or, in the case of persisting strong winds, a salinity gradient.

Because of the shallow water depth (10 – 40 m), the variable river discharge and because of

often variable winds the extent and position of the different water masses may change

considerably even on a short-term scale.

Various aspects of the North Sea bacterioplankton have been the scope of quite a few

investigations, e.g. the culturability of pelagic bacteria (Eilers et al. 2001), seasonal and

interannual dynamics and abundance of specific phylogenetic groups (Eilers et al. 2000,

Gerdts et al. 2004) and the composition of bacterial communities as a function of bacterial

respiration and growth (Reinthaler et al. 2005). The spatial distribution of the bacterial

community composition, e.g. in and off shore gradients including the tidal flat areas and

pelagic regions has not been considered. It is not known how the diversity of the bacterial

communities varies within and among the various water bodies mentioned above, possibly as

a function of a patchy distribution of phytoplankton blooms and the strong tidal currents, or

whether the community composition remains unaffected. The strong currents within the

German Bight and the generally shallow water depth may also lead to a well mixed situation,

preventing the establishment of pronounced regional differences of the bacterial community


118

composition. Obtaining insight into such regional distributions of the composition of bacterial

communities is also important for designing sampling strategies for future investigations,

linking the community composition to hydrographical and biogeochemical processes, not

only in the German Bight, but also in other shallow coastal marine regions exhibiting strong

currents, and estuarine and terrestrial inputs of dissolved and particulate matter.

We investigated the bacterial communities at various near shore and off shore stations

in the German Bight together with properties to characterize the suspended particulate matter

(SPM) and phytoplankton. We used denaturing gradient gel electrophoresis (DGGE) of 16S

rRNA gene fragments and applied primer sets specific for Bacteria, α-Proteobacteria and the

Bacteroidetes phylum.

MATERIALS AND METHODS

Study area and sampling. Surface water samples were collected at various locations in

the German Bight from 11 to 13 June 2002 and from 24 to 27 June 2003 (Fig. 1, Table 1) on

board RV Heincke with a 10 L Niskin bottle. For analysis of suspended matter (SPM) dry

weight (DW), particulate organic carbon (POC), and chlorophyll a (Chl a) 500 to 1000 ml of

sample water were filtered in duplicates on precombusted and preweighed glass fiber filters

(GF/F, Whatman) and stored at –20°C in the dark until further processing in the lab within

four weeks. For enumeration of bacteria 100 ml of seawater were fixed with formaldehyde

(2% v/v) and stored at 4°C until further processing within four weeks. Phytoplankton cells

were fixed with Lugol’s solution as described elsewhere (Utermöhl 1958). For DGGE

analysis, 250 ml of sample water were fractionated by filtration on polycarbonate-filters

(Nuclepore) with pore sizes of 5.0 µm (particle-associated bacteria) and subsequently of 0.2

µm (free-living bacteria) and stored at –20°C until further processing within four months.

Temperature and salinity were recorded by a built-in probe of RV Heincke.

Enumeration of bacteria and algae. Bacteria were enumerated by epifluorescence

microscopy after staining with DAPI (4´-6-diamidino-2-phenylindole) on black 0.2 µm

Nuclepore filters at 1000x magnification (Porter & Feig 1980). We did not differentiate

between free-living (FL) and particle-associated (PA) cells, mainly because it was rather

difficult to reliably enumerate particle-associated cells in the near shore stations with high

concentrations of SPM. When numbers were assessed the reliable desorption technique of PA

bacteria by Lunau et al. (2005) was not yet available. Phytoplankton cells were counted by


119

inverted microscopy (Utermöhl 1958) and phytoplankton species were identified according to

Drebes (1974).

Phytoplankton pigments, SPM and POC. For chlorophyll analysis filters were

extracted at 75°C in 90% ethanol and concentrations of Chl a were determined by standard

procedures (Parsons et al. 1984). For phaeopigment determination, samples were acidified

with HCl (2N) prior to spectrophotometric analysis. For determination of DW, filters were

dried for 1 hour at 110°C and weighed on a micro-balance (Sartorius, Germany). In 2002,

DW was corrected for salt according to Lunau et al. (2006) and in 2003 filters were rinsed

with distilled H2O. POC was determined with a FlashEA 1112 CHN-analyzer (Thermo

Finnigan).

Nucleic acid extraction, primer sets and PCR amplification of 16S rRNA gene

fragments. Genomic DNA was extracted with phenol-chloroform as described in Rink et al.

(2006a) with slight modifications. DNA was precipitated at –20°C overnight using

isopropanol and resuspended in molecular grade water. Samples were stored at –20°C until

further processing. For the amplification of 16S rRNA gene fragments, primer sets were used

targeting eubacterial DNA (primer pair GC 341F, Muyzer et al. 1993; 907RM, Muyzer et al.

1998), the Bacteroidetes phylum (primer pair GC CF319aF, Jaspers et al. 2001, and 907RM)

and α-Proteobacteria (primer pair GC 341F and ALF 968R, Rink et al. 2006a). Specificity of

the primer sets and the applied PCR conditions are described by Rink et al. (2006a).

Amplification products were analyzed by electrophoresis in 1.5% (w/v) agarose gels and

stained with ethidium bromide (1 µg ml-1) (Sambrook et al. 1989). For subsequent sequence

analysis PCR products were purified by using the Qiaquick PCR purification kit (Qiagen Inc.,

Chatsworth, California).

DGGE analysis of PCR products and cluster analysis. DGGE was performed with an

INGENYphorU system (Ingeny International BV, Leiden, The Netherlands) following the

protocol of Rink et al. (2006a). After electrophoresis, the gels were stained with SYBR Gold

(Molecular Probes, Inc.) and documented using a BioDoc Analyze Transilluminator

(Biometra, Göttingen, Germany). Bands were excised, suspended in 50 µl of water (molecular

grade, Eppendorf, Germany) and centrifuged for 2 min. at 3,000 rpm. Samples were stored at

–20°C and 1 µl was used as template in subsequent PCR reactions. A cluster analysis of the

DGGE banding patterns was performed using the software GelCompare II, Version 2.5

(Applied Maths, St. Martens-Latem, Belgium). We applied 5 to 20% background subtraction

depending on the signal-to-noise ratio of the corresponding gel. Patterns were compared

curve-based using Pearson correlation as similarity coefficient and UPGMA (unpaired group


120

method of analysis) to generate the dendrogram. We used the position tolerance optimization

option of the software to fit the curves to the best possible matching.

Cloning. 16 DGGE bands were cloned using the pGEM®-T Vector System II (Promega,

Madison, USA) following the instruction manual. At least five clones per DGGE band with

inserts were picked after blue-white-screening and amplified with the specific DGGE primers.

Fragment length of the inserts was screened by agarose gel electrophoresis and positive

inserts were examined for their specific height using DGGE. Adequate clones were amplified

for sequencing using the primers pUC/M13f and pUC/M13r (Sambrook et al. 1989).

Sequencing and phylogenetic analysis. PCR products were sequenced with an

Automated DNA Sequencer (Model 4200, LI-COR Inc.) using the primers 341F and 907RM,

labeled with IRDyeTM800, and the DYEnamic Direct cycle sequencing kit (Amersham Life

Science Inc.). Clones were sequenced by Geneart (Regensburg, Germany) using the primer

M13f. At least 400 bp were determined for all sequences and the phylogenetic affiliation was

compared to those in GenBank using the BLAST function of the NCBI server

(http://www.ncbi.nlm.nih.gov). The phylogenetic trees were constructed using the ARB

software package (http://www.arb-home.de, Ludwig et al. 2004). The backbone tree was

calculated with the maximum likelihood method using sequences with a minimum of 1300 bp

length including type strains of the selected phylogenetic groups. To avoid uncertain

alignments, positions were excluded at which less than 50% of all sequences showed the same

residues. Sequences with less than 1300 bp were added to the backbone tree with the

maximum parsimony method using the same filter. 16S rRNA gene sequences of seven type

strains belonging to Cyanobacteria were used as outgroup.

Nucleotide sequence accession number. The sequences obtained in this study are

available from GenBank under accession no. DQ911759 to DQ911821.


121

RESULTS

Hydrography. In both years, the off shore stations with depths >17 m (sta 1, 8, 9, 10)

were characterized by lower water temperatures and higher salinities than the near shore

stations (sta 2-7, Fig. 2). At the latter stations water depths is only 8-11 m except at station 2

which exhibits more off shore than near shore properties (see below). In general, temperature

and salinity were higher in 2003 as compared to 2002, when the survey was carried out two

weeks earlier. In 2002, surface water temperatures ranged from 13.7°C (sta 1, 8) to 16.5°C

(sta 7) and in 2003 from 14.4°C (sta 10) to 17.6°C (sta 4). Salinity ranged from 29.6 (sta 7) to

33.2 psu (sta 9) in 2002 and from 31.8 (sta 6) to 35.5 psu (sta 9) in 2003.

Abundance and composition of SPM. In both years, SPM concentrations were higher at

the near shore than at the off shore stations (Fig. 3A, 3B). In 2002, SPM ranged from 4 to 5

mg DW l-1 at the off shore stations and station 2 to 7.6 to 16 mg DW l-1 at the near shore

stations with the highest value at station 5. In 2003, respective values at the off shore stations

(sta 1, 2, 8, 9, 10) were between 2 and 4 mg DW l-1 and at the near shore stations 6.5 to 15.4

mg DW l-1. POC varied from 0.10 mg l-1 at station 9 to 0.78 mg l-1 at station 6 with generally

higher values at the near shore than at the off shore stations (Fig. 3B). At stations 9, 10 and 2

POC constituted around 10% of DW but at the near shore stations generally <6%.

Phytoplankton. Chl a concentrations in 2002 were lower than in 2003 and varied from

0.8 to 4.5 µg l-1 (Fig. 3C, 3D). In 2002, concentrations at the near shore stations were higher

than at the offshore stations except at station 8. In 2003, highest concentrations occurred at

stations 3 and 8 with up to 6.3 µg Chl a l-1 without any clear-cut difference between the near

and off shore stations. Patterns of phaeopigment concentrations, which are only available for

2003, were quite different from that of Chl a (Fig. 3D), indicating that the growth phase and

thus the physiological status of the phytoplankton was quite variable among the stations.

Highest proportions of phaeopgments, close to or exceeding those of Chl a, occurred at

stations 6, 7 and 9. In 2002, Leptocylindricus danicus dominated the phytoplankton at the off

shore stations and Rhizosolenia imbricata at the near shore stations (data not shown). At the

North Frisian coast (sta 6, 7), Guinardia delicatula and Guinardia flaccida were also present

to substantial proportions. In 2003, the phytoplankton at all stations was highly dominated by

Rhizosolenia imbricata except at station 10 which showed a more diverse composition

including substantial proportions of Guinardia spp. (Fig. 3E). Also at station 8, Guinardia

spp. were the second most abundant taxa. Phytoplankton cell numbers generally reflected

concentrations of Chl a with highest values at stations 3 and 8. At station 6 and 7 the high


122

sediment load in the samples and low phytoplankton cell numbers prevented a reliable

enumeration.

Bacterial abundance. In 2002, bacterial abundance varied between 1.8 x 106cells ml-1 at

station 1 and 3.5 x 106cells ml-1 at station 7. Numbers did not covary with Chl a or SPM. In

2003, the off shore stations exhibited low bacterial numbers and highest numbers were

recorded at stations 3 and 6 together with high concentrations of Chl a and SPM.

DGGE and cluster analysis. Pronounced differences were detected between DGGE

banding patterns of FL and PA bacterial communities. These differences were substantiated

by a cluster analysis (Fig. 4). The application of the Bacteria-specific primer set yielded 7 to

13 DGGE bands of the FL bacterial community in 2002 and 15 to 24 bands in 2003. In the

former year, the lowest and highest number of bands occurred at stations 2 and 1 and in the

latter year at stations 1 and 6. Two prominent bands persisted throughout all stations in both

years, identified as clones GB02-e8-FL, GB02-e9-FL, GB03-e16-FL and GB03-e17-FL

(Table 2). Other bands occurred only at a few or individual stations such as at stations 1 and

10 (GB03-e15-FL), 6 and 7 (GB03-e24-FL, GB03-e25-FL), and 5 (GB03-e23-FL). There was

no band detected only at the off shore or near shore stations. In 2003, stations 6 and 7,

however, exhibited distinctly different patterns than the other stations.

In the PA bacterial community in both years, the number of bands exceeded that of the

FL bacterial community. In 2002, between 10 and 17 bands were detected with lowest and

highest numbers at stations 10 and 1, respectively. In 2003, band numbers ranged between 12

and 28 with lowest and highest numbers at stations 1 and 6. The variability of the PA bacterial

community among the different stations was more pronounced than that of the FL bacterial

community. No single band was detected at all stations. The banding patterns of the PA

bacterial community were highly biased by chloroplast-derived 16S rRNA gene fragments.

Two of the 7 bands of the 2002 samples and 8 of the 14 bands of the 2003 samples sequenced

turned out as chloroplast-like phylotypes.

The cluster analysis substantiated the different banding patterns of the FL- and PA

bacterial communities detected by the Bacteria-specific primer set (Fig. 4). The banding

patterns of the PA bacterial communities exhibited a lower similarity (>55% Pearson

correlation) than those of the FL bacterial communities (>76% Pearson correlation). Even

though micro-clusters occurred, near shore and off shore stations did not exhibit distinct

clusters. Stations 6 and 7, however, formed a separate cluster in both bacterial communities in

2003.


123

The DGGE analysis of the FL- and PA associated bacterial communities applying the

Bacteroidetes-specific primer set was only done in 2003. The results also revealed

pronounced differences between both communities with a higher diversity in the PA bacterial

community. In the FL bacterial community between 7 (sta 8) and 13 bands (sta 1) were

detected, and in the PA bacterial community between 9 (sta 7) and 17 bands (sta 5). There

was no band which was detected at all stations, neither in the FL- nor in the PA bacterial

community. However, several bands occurred at distinct stations, such as at stations 1-5 and 8

(GB03-c5-PA, Table 3), at stations 2-5 (GB03-c8-FL, Table 2), and at stations 1, 8 and 10

(GB03-c12-FL). In the PA bacterial community, stations 2-5 and 10 clustered together, as did

stations 6 and 7 (Fig. 4). The off shore stations 1, 8 and 9 branched deeply separated. In the

FL fraction, no specific sub-clusters were detected.

The α-Proteobacteria–specific DGGE banding patterns revealed lowest band numbers of

all target groups with 5 to 7 bands in the FL bacterial community and 7 to 16 bands in the PA

bacterial community. In the former community, two conspicuous bands (GB-a14-FL, GB-

a15-FL, Table 2) dominated the banding patterns at all stations and the similarity between the

banding patterns was very high, as confirmed by the cluster analysis (Fig. 4).The banding

patterns of the PA bacterial community were much more diverse. Only stations 6 and 7

clustered together.

Phylogenetic affiliation. Sequencing of the excised DGGE bands obtained from the

Bacteria-specific amplicons showed that 12 bands of a total of 38 contained chloroplast-

derived 16S rRNA gene fragments. Ten of them were detected in the PA fraction. In addition,

13 sequences affiliated to α-, 1 to β, 2 to γ-Proteobacteria and 10 to the Bacteroidetes-

phylum.

All sequences of α-Proteobacteria obtained from the Bacteria-specific DGGE gels

affiliated to the Roseobacter clade except GB02-e3-PA (station 4), which was related most

closely to Acidiphilium aminolytica (Tables 2, 3, Fig. 5A). The phylotypes occurring at all

stations in both years, GB02-e8-FL, GB03-e16-FL, GB02-e9-FL and GB03-e17-FL, affiliated

to the NAC11-7 cluster detected in the North Atlantic (Gonzalez et al. 2000) and the WM11-

36 cluster identified in the polyhaline section of the Weser estuary (Selje & Simon 2003).

The two clones which affiliated to γ-Proteobacteria (GB03-e7-PA, GB03-e10-PA) were

detected in 2003 in the PA bacterial community of station 6 and were closely related to clones

from the East Frisian Wadden Sea (GWS-AG-6, GWS-SE-4, Fig. 5C).

Sequences of DGGE bands obtained with the Bacteroidetes specific primer set revealed 2

unspecific amplifications. DGGE band GB03-c1-PA (sta 1, PA fraction) was related to a


124

Firmicutes species (Table 3) and band GB03-c6-PA to chloroplast-derived 16S rRNA genes.

The sequences of two DGGE bands obtained from the PA fraction of station 6 (North Frisian

coast, GB03-c3-PA and GB03-c4-PA) were closely related to 16S rRNA gene fragments

detected as a diatom-associated bacterium and in coastal bacterioplankton (GWS-AG-8,

GWS-c2-FL, Fig. 5D), respectively. Clone GB03-c2-PA (sta 6) was closely related to strain

T15, isolated from a high dilution step of a dilution culture from the East Frisian Wadden Sea

(Brinkhoff et al. 2004).

Sequencing of 16S rRNA gene fragments obtained from the α-Proteobacteria specific

DGGE gel showed unspecific amplification in the PA bacterial community. Nine of the 13

sequences obtained were identified as δ-Proteobacteria predominantly originating from

sediments and only 4 as α-Proteobacteria (Fig. 5A and B, Table 3). Two of them affiliated to

the Roseobacter clade, to the RCA cluster (GB03-a5-PA) and to Sulfitobacter pontiacus (GB-

a4-PA), and the other two to Rhodobacterales (GB-a7-PA) and to the genus Sphingomonas

(GB03-a1-PA), respectively. The 4 sequenced bands of the FL bacterial community of the α-

Proteobacteria specific DGGE gel affiliated to the clusters RCA (GB03-a14-FL) and WM11-

36 (GB03-a15-FL, GB03-a16-FL) of the Roseobacter clade and one closely to a phylotype

retrieved from the German Wadden Sea and related to Acidiphilum aminolytica. (GB03-a17-

FL).

DISCUSSION

We found a surprisingly high variability of the DGGE banding patterns both of FL and

PA bacterial communities of either primer set applied in the German Bight of the North Sea

which, however, only partially reflected the clear differences of salinity and SPM

concentrations between the near shore and off shore stations. Similarly, banding patterns did

not reflect patterns in the distribution of Chl a or phytoplankton composition. We did find

distinct differences between banding patterns of PA and FL-bacterial communities,

substantiated by the cluster analysis. A number of phylotypes in both communities only

occurred at certain stations near shore or off shore, indicating that hydrographic and

biogeochemical differences did affect the composition of the bacterial communities to a

certain extent. The only stations which exhibited clearly different banding patterns and

formed a distinct subcluster in 2003 were stations 6 and 7 close to the North Frisian coast.

When samples were collected at these shallow stations rather strong wind (Beaufort scale 6-7)

prevailed leading to high sediment resuspension. The unspecific detection of sediment-


125

associated δ-Proteobacteria in these samples is a further indication of the sediment

resuspension. Several prominent DGGE bands of the FL bacterial communities amplified

with the Bacteria- and α-Proteobacteria-specific primer sets, however, were present at all

stations, indicating that several populations of this community persisted in the German Bight,

irrespective of the given hydrographic and biogeochemical conditions. The variability within

the PA bacterial community among the various stations was greater than that of the FL

bacterial community, indicating that site-specific properties affected more the former than the

FL bacterial community. However, unspecific amplification of chloroplast-derived 16S rRNA

gene fragments by the Bacteria-specific primer set and of δ-proteobacterial 16S rRNA gene

fragments by the α-Proteobacteria-specific primer set contributed to this variability and

biased the banding patterns and cluster analysis of the PA bacterial community.

Our investigation was carried out during several days in June of two consecutive years,

thus covering only a short period of the annual development of the German Bight. There are

consistent reports of general annual patterns of the hydrography and biological development

at individual stations such as at Helgoland Roads, at Norderney (East Frisian Wadden Sea)

and Büsum and Sylt (North Frisian Wadden Sea) (BSH 2002 and 2003, Loewe et al. 2005).

Even though they exhibit a general seasonal trend, the patterns of each station do vary.

Further, the individual stations show short-term deviations from the seasonal trends, also for

June 2002 and 2003, presumably because of wind- and current-induced movements of water

masses with different physico-chemical and biological properties. Our surveys in both years

encountered two different biological situations, as shown by the different Chl a

concentrations and composition of the phytoplankton, and also reflected by the generally

higher number of DGGE bands detected both in the FL and the PA bacterial community in

2003. The physiological state of the phytoplankton among the near shore and off shore station

varied, as indicated by the variable proportions of phaeopigments relative to total chlorophyll.

Hence it seems not surprising that we did not find consistent patterns of the composition of

the bacterial communities in the near and off shore regions in a rather dynamic regional

coastal sea.

As mentioned, our DGGE and cluster analysis was biased by unspecific PCR

amplifications. Unspecific amplifications by the Bacteria-specific primer set applied has been

reported previously (e.g. Selje & Simon 2003, Stevens et al. 2005a) and amplification of

chloroplast-derived 16S rRNA gene fragments in the PA bacterial community with this

primer set appears a general problem for samples containing phytoplankton cells. Unspecific

amplification with primer ALF968r (α-Proteobacteria) has also been reported by Rink et al.


126

(2006a) for samples from the East Frisian Wadden Sea and by Overmann et al. (2005). The

latter authors found amplification of Actinobacteria in freshwater samples and possible

detection of few γ-Proteobacteria. Unspecific amplification of δ-Proteobacteria by the

ALF968r primer predominantly occurred in the PA bacterial community at the shallow near

shore stations (3-7) with high concentrations of resuspended SPM. Marine sediments contain

high numbers of sulfate reducing bacteria affiliated to δ-Proteobacteria (Llobet-Brossa et al.

1998, Musat et al. 2006) and also myxobacteria which affiliate to this subclass of

Proteobacteria as well (Stevens et al. 2005a). Hence, when these bacteria are present in the

samples to detectable amounts, their 16S rRNA genes are amplified by this primer. In the FL

bacterial community, no unspecific amplification occurred, indicating that this community did

not include δ-Proteobacteria in proportions high enough to be amplified. In conclusion, a

reliable assessment of the composition of PA α-Proteobacteria by the subclass-specific

primer set was not possible. Our finding of unspecific amplification of the ALF968r primer

has important implications for the interpretation of data obtained by fluorescence in situ

hybridization (FISH) applying this oligonucleotide as a probe (ALF968). In habitats with high

proportions of δ-Proteobacteria they may be included in the detection of α-Proteobacteria

and thus lead to overestimating this subclass.

Also the Bacteroidetes–specific primer set (Jaspers et al. 2001) resulted in unspecific

amplification in three of 12 cases. Even though this is also a bias of the DGGE results, it

appears not as critical as that with the α-Proteobacteria–specific primer set, but emphasizes

the importance for sequencing of prominent bands applying group-specific primer sets in

DGGE analyses.

These biases of unspecific amplification predominantly affected the DGGE banding

pattern of the PA bacterial community obtained by the Bacteria–specific (Fig. 4B) and α-

Proteobacteria–specific primer sets in 2003 (Fig. 4D). The other banding patterns were only

marginally affected. Hence our general findings of greater differences of the composition of

the PA bacterial community among the various stations and a greater diversity as compared to

the FL bacterial community is unaffected by these biases. Our findings are consistent with

previous reports from single stations of the same region (Gerdts et al. 2004, Stevens et al.

2005a). However, similarly as these two reports, we did not enumerate the PA bacteria and

thus can not directly assess their significance relative to that of the FL bacteria. Particle-

associated bacteria have been enumerated in the SPM-rich East Frisian Wadden Sea by Lunau

et al. (2006). In June and July of various years, numbers of PA bacteria vary from 23 to 32%

of total bacteria, and we assume that these numbers are representative for the near shore


127

stations in general. Comparable numbers are not available for the off shore region of the

German Bight. This region is characterized by low numbers of aggregates (Riebesell 1991),

and PA bacteria in such neritic seas usually constitute <10% of total bacterial biomass and

activity (Simon et al. 2002). Therefore, we assume that PA bacteria are of much lower

significance in the off shore than in the near shore region of the German Bight.

The phylogenetic lineages we identified have previously been shown to constitute the

bacterial communities in the German Bight to a great extent. Applying FISH, Eilers et al.

(2000 and 2001) found that at Helgoland Roads Bacteroidetes and α- and γ-Proteobacteria

constitute 18–30, 15–25 and 6–9% of DAPI-stainable bacteria, respectively, at various

seasonal situations including June. These groups also constitute the bacterial communities in

the Wadden Sea to a great extent, with rather equal proportions in the FL (each group 10–

20% of DAPI stainable bacteria) and PA fractions (each group 15–40%) (Rink et al. 2006b).

Our results, in agreement with previous studies (Eilers et al. 2000, Selje et al. 2004, Stevens et

al. 2005a, Zubkov et al. 2002), show that members of the Roseobacter clade and in particular

of distinct subclusters (RCA, WM11-36, NAC11-7) are important components of the FL

bacterial community (Fig. 5A). As indicated by the prominent DGGE bands persisting at all

stations (GB02-e8-FL, GB02-e9-FL, GB-a14-FL, GB-a15-FL, GB03-e16-FL, GB03-e17-FL)

and the high similarity of the banding patterns, the FL α-Proteobacteria sub-community was

most evenly distributed in the German Bight and least affected by hydrographic and

biogeochemical differences of the various regions. The occurrence of closely related

phylotypes in other neritic and oceanic regions (Gonzalez et al. 2000, Selje et al. 2004)

emphasizes that bacteria of this lineage are important components of the marine

bacterioplankton globally.

The FL sub–community of the Bacteroidetes phylum exhibited a greater variability and

diversity than that of α-Proteobacteria, as shown by the group-specific DGGE banding

patterns and cluster analysis and the occurrence of more phylotypes at distinct stations and of

none at all stations. The phylotypes were retrieved from the bands amplified by both the

Bacteria-specific and Bacteroidetes–specific primer sets. This notion indicates the

significance of site-specific factors for controlling the composition of this sub-community.

The phylotypes affiliated to rather different clusters of the Bacteroidetes phylum but in most

cases were closely related to other phylotypes previously detected in the Wadden Sea, the

German Bight or North Sea during phytoplankton blooms (Table 2, Fig. 5D, Rink et al.

2006a, Stevens et al. 2005a and 2005b, Zubkov et al. 2002, J. Pernthaler et al. unpubl.). Close

relationships between phylotypes and isolates of the Bacteroidetes phylum and distinct


128

phytoplankton and algal populations have been reported (Schäfer et al. 2002, Grossart et al.

2005, Rooney-Varga et al. 2005). Further, bacteria of this phylum are known to degrade

complex organic polymers (Cottrell & Kirchman 2000, Kirchman 2002). Therefore, it appears

that the phylotypes affiliated to Bacteroidetes reflect more the specific substrate conditions

related to the phytoplankton community of different growth stages and composition and to

other substrate sources at the various stations.

Particles and suspended aggregates provide much more diverse micro-habitats than the

surrounding water. Depending on the given conditions and the water depth, they may include

phytoplankton-derived organic matter as well as organic and inorganic matter resuspended

from the sediment. Therefore, it appears not surprising that the PA bacterial community was

more diverse than the FL bacterial community and contained more site-specific phylotypes

reflecting the local environmental conditions. The PA bacterial community contained quite a

few phylotypes affiliated to various clusters of the Bacteroidetes phylum (Fig. 5D, Table 3),

but also various phylotypes affiliated to α- and γ-Proteobacteria and distinct from those in the

FL bacterial community. The PA bacterial phylotypes affiliated to α-Proteobacteria were

related mainly to various clusters of the Roseobacter clade which differ from those containing

the FL bacterial phylotypes. Others affiliated to Sphingomonas and Acidiphilum aminolyticum

(Fig. 5A, Table 2). The two γ-Proteobacteria phylotypes were detected in 2003 at stations 6

and 7 in the North Frisian Wadden Sea and closely related to phylotypes retrieved from the

East Frisian Wadden Sea (Fig. 5B, Stevens et al. 2005b). One phylotype fell into the

SAMMIC-cluster (Surface Attached Marine MICrobes, Stevens et al. 2005b). Members of

this cluster are uncultured, globally distributed and always associated to suspended aggregates

in coastal systems or to sediments. The unspecific amplification of the tentatively α-

Proteobacteria–specific primer set also retrieved exclusively phylotypes of sediment-

associated δ-Proteobacteria (Fig. 5B, Table 3).

In conclusion, our results show a variable diversity and distribution patterns of the

bacterial community in the German Bight of the North Sea. The FL bacterial community is

rather similar in this coastal sea and harbors several widely distributed members, affiliated to

the Roseobacter clade of α-Proteobacteria and obviously little affected by site-specific

environmental conditions. However, other members occurred only at specific locations,

obviously as the result of site-specific environmental conditions of the tidal flat areas and the

phytoplankton communities. The PA bacterial community was more divers and reflected

mainly local environmental features, also related to the specific environmental conditions of


129

the tidal flat areas with high SPM concentrations and intense resuspension and of the

phytoplankton communities.

ACKNOWLEDGEMENTS

We appreciate the hospitality and cooperation of the captain and crew of RV Heincke. We

thank B. Kuerzel for dry weight and chlorophyll analyses and A. Luek for phytoplankton and

bacterial cell counts. This work was supported by the Deutsche Forschungsgemeinschaft

(DFG) within the research group “BioGeoChemistry of the Wadden Sea” (FG 432 TP5).


130

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Table 1: Location of sampling stations, water depth and days of sampling.

Station-no.

Latitude (°N)

Longitude (°E)

Water depth (m)

Days of sampling June ‘02 June ‘03

1 54° 07.98’ 7° 04.64’ 32 13 26

2 53° 49.69’ 7° 15.31’ 18 12 26

3 53° 48.33’ 7° 38.45’ 8 12 27

4 53° 52.95’ 8° 05.24’ 8 13 24

5 53° 59.58’ 8° 03.52’ 8 13 24

6 54° 13.91’ 8° 20.66’ 11 11 25

7 54° 32.08’ 8° 10.98’ 9 11 25

8 54° 36.88’ 7° 42.26’ 17 11 25

9 54° 38.44’ 6° 56.41’ 36 11 26

10 54° 28.15’ 7° 15.05’ 29 11 26


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Table 2: Phylogenetic affiliation, band-identification (ID), accession number and origin (station) of the free-living bacterial 16S rRNA gene phylotypes retrieved in this study, and their closest related phylotypes, similarity to them and origin.

Phylogen. class Band ID Acc. no. Station Closest relative (acc. no.) Similarity (%)

Origin

α-Proteobacteria GB02-e8-FL DQ911764 1 Uncultured alpha proteobacterium clone CONW88 (AY828363)

96 environmental sample, Loch Fyne, Scotland

GB02-e9-FL DQ911765 1 Uncultured Roseobacter sp. clone EF100-65C12 (AY627371)

98 environmental sample, Monterey Bay, California

GB03-e16-FL DQ911810 1 Uncultured Roseobacter NAC11-7 (AF245635)

98 associated with a DMSP-producing North Atlantic algal bloom



GB03-e18-FL DQ911812 2 Uncultured alpha proteobacterium clone RAN-63 (AY499446)

94 bacterioplankton of Ria de Aveiro, Portuguese estuary

GB03-e19-FL DQ911813 2 Uncultured alpha proteobacterium clone CONW83 (AY828403)




GB03-e26-FL DQ911820 9 Uncultured Roseobacter NAC11-7 (AF245635)

99 associated with a DMSP-producing North Atlantic algal bloom

GB03-e27-FL DQ911821 10 Uncultured alpha proteobacterium clone CONW83 (AY828403)


GB03-c10-FL DQ911777 9 Uncultured Rhodobacteraceae bacterium clone ESP450-K6III-60 (DQ810729)

97 Oxygen minimum zone, Chile

GB03-c11-FL DQ911778 9 Uncultured alpha proteobacterium isolate DGGE band GWS-TC-e9-FL (DQ911830)

99 German Wadden Sea

GB03-a14-FL DQ911793 1 Uncultured alpha proteobacterium isolate DGGE band GWS-FL-2 (AY274228)

100 German Wadden Sea, June 2000

GB03-a15-FL DQ911794 1 Uncultured Roseobacter sp. clone EF100-65C12 (AY627371)


GB03-a16-FL DQ911795 4 uncultured alpha proteobacterium CHAB-I-5 (UAL240910)

99 marine bacterial assemblage during confinement

GB03-a17-FL DQ911796 6 uncultured alpha proteobacterium CHAB-I-5 (UAL240910)

99 marine bacterial assemblage during confinement

β-Proteobacteria GB03-e25-FL DQ911819 7 Uncultured marine bacterium clone

SPOTSAPR01_5m110 (DQ009366) 99 marine bacterioplankton, San

Pedro Ocean Time Series Bacteroidetes

GB02-e10-FL DQ911766 5 Uncultured Flavobacteria bacterium 16S rRNA gene, clone NorSea33 (AM279185)

97 Helgoland Roads; coastal North Sea

GB03-e15-FL DQ911809 1 Uncultured Bacteroidetes bacterium isolate DGGE band GWS-c3-FL (DQ080946)

96 German Wadden Sea



GB03-e23-FL DQ911817 5 Uncultured bacterium isolate DGGE band D37 (AF466926)

95 environmental sample, associated with dinoflagellate A. tamarense



GB03-c7-FL DQ911774 3 Uncultured Flavobacteria bacterium 16S rRNA gene, clone NorSea76 (AM279177)


GB03-c8-FL DQ911775 4 Uncultured Flavobacteria bacterium 16S rRNA gene, clone NorSea58 (AM279203)


GB03-c9-FL DQ911776 5 Uncultured Sphingobacteria bacterium 16S rRNA gene, clone NorSea49 (AM279196)


GB03-c12-FL DQ911779 10 Uncultured bacterium clone CTD5B (AF469385)

97 environmental sample, subseafloor, after deep-sea volcanic eruption


135

Table 3: Phylogenetic affiliation, band-identification (ID), accession number and origin (station) of the particle-associated bacterial 16S rRNA gene phylotypes retrieved in this study, and their closest related phylotypes, similarity to them and origin.

Phylogen. class Band ID Acc. no. Station Closest relative (acc. no.) Similarity (%)

Origin

α-Proteobacteria GB02-e3-PA DQ911759 4 Uncultured proteobacterium OCS126 (AF001638)

99 bacterioplankton, continental shelf off Oregon, USA; SAR116 cluster

GB02-e4-PA DQ911760 5 Sulfitobacter sp. DG1020 (AY258095) 98 strain associated with the dinoflagellate Gymnodinium

catenatum GB02-e6-PA DQ911762 10 Roseobacter sp. MED008 (AY136104) 98 environmental sample, eastern

Mediterranean Sea GB02-e7-PA DQ911763 10 Roseobacter sp. RED1 (AY136122) 97 environmental sample, Gulf of

Eilat, Red Sea GB03-a1-PA DQ911780 1 Sphingomonas sp. Pd-S-(s)-m-D-1(2)

(AB242948) 99 Endophytic at Rice Plants (Oryza

sativa) GB03-a4-PA DQ911783 4 Uncultured marine bacterium clone

AntCL2C1 (DQ906745) 98 Antarctica: near Anvers Island

GB03-a5-PA DQ911784 5 Uncultured Rhodobacteraceae bacterium clone RCA-H28 (DQ489286)

99 10e-5 dilution step, marine sample, Weser Estuary, Germany

GB03-a7-PA DQ911786 6 Uncultured alpha proteobacterium, clone T63ANG236 (AJ633963)

96 epibiotic bacteria in the accessory nidamentalglands of squids

δ-Proteobacteria GB03-a2-PA DQ911781 3 Uncultured delta proteobacterium clone

YS-UMF1_C112 (DQ901575) 98 environmental sample, intertidal

sediment, Korea GB03-a3-PA DQ911782 3 Uncultured delta proteobacterium, isolate

DGGE band 160NF32 (AM072605) 99 German Wadden Sea, sediment

GB03-a6-PA DQ911785 5 Uncultured delta proteobacterium clone Belgica2005/10-130-17 (DQ351762)

100 Sediment surface associated, North Sea

GB03-a8-PA DQ911787 7 Uncultured delta proteobacterium clone YS-UMF1_C112 (DQ901575)

93 environmental sample, intertidal sediment, Korea

GB03-a9-PA DQ911788 7 Unidentified proteobacterium OM27 (U70713)

99 marine coastal picoplankton, continental shelf, off Cape Hatteras, North Carolina

GB03-a10-PA DQ911789 7 Uncultured bacterium DGGE gel band FD 20 (DQ385045)

96 Faroe Deep of the central Baltic Sea (2 m)

GB03-a11-PA DQ911790 7 Uncultured delta proteobacterium clone Belgica2005/10-130-15 (DQ351760)


GB03-a12-PA DQ911791 7 Uncultured delta proteobacterium clone Belgica2005/10-ZG-2 (DQ351798)


GB03-a13-PA DQ911792 8 Uncultured delta proteobacterium clone AKYH967 (AY922176)

93 farm soil adjacent to a silage storage bunker

γ-Proteobacteria GB03-e7-PA DQ911802 6 Uncultured gamma proteobacterium isolate DGGE band GWS-TC-e4-PA (DQ911825)

98 aggregate-associated, German Wadden Sea

GB03-e10-PA DQ911804 7 Uncultured delta Proteobacterium, isolate DGGE band IIIA3 (AJ889159)

96 coastal subsurface sediment German Wadden Sea

Bacteroidetes GB02-e5-PA DQ911761 9 Marine Eubacterial sp. (aggregate agg32)

(L10944) 99 aggregate-attached, marine

bacterial assemblages GB03-e4-PA DQ911799 6 Uncultured bacteroidetes bacterium, isolate

DGGE band 100G15 (AJ880446) 98 Tidal flat sediment (1 m) German

Wadden Sea GB03-e5-PA DQ911800 6 Uncultured bacterium SB-42-DB

(AJ319829) 97 satellite bacterium of Dytilum

brightwellii GB03-e14-PA DQ911808 9 Marine Eubacterial sp. (aggregate agg32)

(L10944) 98 aggregate-attached, marine

bacterial assemblages GB03-c2-PA DQ911769 6 Flavobacteriaceae bacterium T15

(AY177723) 98 strain, 10e-5 dilution step, German

Wadden Sea GB03-c3-PA DQ911770 6 Uncultured Bacteroidetes bacterium clone

PI_4q10f (AY580698) 96 coastal bacterioplankton sample of

Plum Island Sound Estuary GB03-c4-PA DQ911771 6 Uncultured bacterium SB-42-DB

(AJ319829) 98 satellite bacterium of Dytilum

brightwellii GB03-c5-PA DQ911772 8 Flavobacteriaceae bacterium T15

(AY177723) 98 strain, 10e-5 dilution step, German

Wadden Sea Firmicutes GB03-c1-PA DQ911768 1 Uncultured bacterium clone Napoli-4B-79

(AY592793) 96 marine sediment, Napoli mud

volcano, Eastern Mediterranean


136

Figure Legends

Fig. 1.

Study area and locations of the sampling stations in the German Bight, Southern North Sea.

Fig. 2.

Scatter plot of temperature versus salinity of the sampling stations in the German Bight in

June 2002 and 2003. For exact locations and dates see Table 1.

Fig. 3.

Suspended matter dry weight (SPM) in 2002 (A) and 2003 (B), POC in 2003 (B), Chlorophyll

a in 2002 (C) and 2003 (D), phaeopigments in 2003 (D), phytoplankton cell numbers in 2003

(E) bacterial numbers in 2002 (F) and 2003 (G) at various stations in the German Bight.

Samples were collected in June of both years. Stations 8-10 and 1 are off shore, stations 2-7

near shore. For exact locations and dates see Table 1. Missing bars: data not available.

Fig. 4.

Cluster analyses of the DGGE banding patterns of particle-attached and free-living Bacteria

(EUB02, A; EUB03, B), Bacteroidetes (CFB03, C) and α-Proteobacteria (ALF03, D)

retrieved from samples at various stations in the German Bight in June 2002 (02) and 2003

(03). PA: Particle associated bacterial community.

Fig. 5.

Phylogenetic trees of α-Proteobacteria (A), δ-Proteobacteria (B), β- and γ-Proteobacteria

(C) and the Bacteroidetes phylum (D) calculated with Maximum-Likelihood based on 16S

rRNA gene fragments. Sequences obtained in this study are highlighted in bold.


137

Fig. 1. Rink et al.

Fig. 2. Rink et al.

Salin

ity

(psu)

28

30

32

34

36

Temperature (°C)

13 14 15 16 17 18

28

30

32

34

36

2002

2003

7

54

2

6

9

10

1

8

3

8

7

19

102

34

5

6


138

Fig. 3. Rink et al.

SP

M d

ry w

eig

ht

(mg

L-1

)

0

4

8

12

16

PO

C (

mg L

-1)

0,0

0,4

0,8

1,2

1,6

Ch

loro

ph

yll

a (

µg

L-1

)

0

2

4

6

SPM

POC

Ph

ae

o (

µg L

-1)

0

2

4

6Chl

Phaeo

2002 2003

Ph

yto

pla

nkto

n (

ce

lls m

l-1)

0

200

400

600

800

1000

1200

8 9 10 1 2 3 4 5 6 78 9 10 1 2 3 4 5 6 7

Stations

A B

C D

E

F G

Stations

Ba

cte

ria

(10

6 c

ells

mL

-1)

0

1

2

3

4

5

6

Rhizosolenia imbricata

Rhizosolenia spp


Guinardia flaccida



Leptocylindrus danicus

F


139

Fig. 4. Rink et al.

St. 6 St. 7

St. 3 St. 8

St. 6

100

75

50

25

100

10095

8094

87

84

83

10095

100

10067

68

75

100

76

94

100

96

St. 7 St. 5 St. 3 St. 2 St. 10 St. 1 St. 8 St. 9 St. 10 St. 1

St. 5 St. 2 St. 4 St. 9

Std. rightStd. left

Particle-attached

Free-living

St. 6 St. 7 St. 4 St. 5 St. 10 St. 2 St. 3 St. 8 St. 1 St. 9 St. 6 St. 5 St. 2 St. 3St. 7 St. 4St. 1St. 9St. 8St. 10


St. 7St. 6 St. 8St. 10St. 2St. 3St. 5St. 9St. 4St. 7St. 6St. 1St. 2St. 4St. 5St. 3St. 9 St. 8 St. 10

Particle-attached

Free-living

D

Free-living

Particle-attached

St. 4St. 5St. 1St. 10St. 6St. 8St. 3St. 9St. 7St. 2St. 10 PASt. 5St. 1St. 8St. 7St. 9St. 4St. 2St. 6St. 3Std. leftStd. right

EUB 02

A

EUB 03

CFB 03

ALF 03

B

CParticle-attached

Free-living

St. 6 St. 7

St. 3 St. 8

St. 6

100

75

50

25

100

10095

8094

87

84

83

10095

100

10067

68

75

100

76

94

100

96

St. 7 St. 5 St. 3 St. 2 St. 10 St. 1 St. 8 St. 9 St. 10 St. 1

St. 5 St. 2 St. 4 St. 9

Std. rightStd. left

Particle-attached

Free-living

St. 6 St. 7 St. 4 St. 5 St. 10 St. 2 St. 3 St. 8 St. 1 St. 9 St. 6 St. 5 St. 2 St. 3St. 7 St. 4St. 1St. 9St. 8St. 10


St. 7St. 6 St. 8St. 10St. 2St. 3St. 5St. 9St. 4St. 7St. 6St. 1St. 2St. 4St. 5St. 3St. 9 St. 8 St. 10

Particle-attached

Free-living

D

Free-living

Particle-attached

St. 4St. 5St. 1St. 10St. 6St. 8St. 3St. 9St. 7St. 2St. 10 PASt. 5St. 1St. 8St. 7St. 9St. 4St. 2St. 6St. 3Std. leftStd. right

EUB 02

A

EUB 03

CFB 03

ALF 03

B

CParticle-attached

Free-living


140


A

Roseobacterclade

RCA

WM11-36

NAC11-7

A

Roseobacterclade

RCA

WM11-36

NAC11-7


141

Fig. 5B, C. Rink et al.

B

C

gamma

SAMMIC

beta

B

C

gamma

SAMMIC

beta


142

D

Fig. 5D. Rink et al.

D

Fig. 5D. Rink et al.


143

Supplementary Figure:

DGGE banding patterns which are the basis for the cluster analysis. Numbers and arrows

indicate excised and sequenced bands and asterisks chloroplast-derived 16S rRNA gene

phylotypes.

144

VI.

Schlussbetrachtung und Ausblick

Kapitel VI Schlussbetrachtung und Ausblick

145

Die vorliegende Arbeit umfasst detaillierte Untersuchungen zur Struktur und Abundanz des

Bakterioplankton im ostfriesischen Wattenmeer sowie der Deutschen Bucht. Besondere

Berücksichtigung fand die Entwicklung und Anwendung spezifischer Nachweise für

Bakteriengruppen, die durch methodische Einschränkungen in der Vergangenheit als bisher

unterrepräsentiert angesehen werden mußten. Die Ergebnisse dieser Arbeit wurden im

Rahmen der DFG geförderten Forschergruppe „BioGeoChemie des Watts“ gewonnen und

stellen eine wesentliche Grundlage für zukünftige Untersuchungen bezüglich des

Stoffumsatzes abundanter Bakteriengruppen dar.

Das Bakterioplankton im Wattenmeer kann in die Kompartimente freilebend, Aggregat- und

Sediment- assoziiert unterteilt werden (vgl. Stevens et al. , 2005,). Die DGGE-Auswertung

von Probenahmen in monatlichen Abständen ergab eine relative Stabilität der

Bakteriengemeinschaften im Wattenmeer (Stevens et al., 2005). Diese veränderten sich nur

geringfügig zu extremen Ereignissen, z. B. während und nach Phytoplanktonblüten und bei

Sturm. Da diese Ereignisse fester Bestandteil der Jahreszeiten sind und regelmäßig

wiederkehren, wurden sie als stabile Merkmale des Ökosystems Wattenmeer gewertet und die

Schlussfolgerung war, dass das Bakterioplankton praktisch keinen Änderungen unterliegt. Die

Aussagen von H. Stevens sind, basierend auf den damals erhobenen Daten, durchaus

zutreffend, bedürfen allerdings durch die Ergebnisse der vorliegenden Arbeit nun der

Ergänzung.

In Kapitel II wurde deutlich, dass die große Stabilität der Bakteriengemeinschaft bei

genauerer Betrachtung über den Zeitraum einer Phytoplanktonblüte moduliert wird. Man kann

annehmen, dass durch Ausscheidungen der Diatomeen und auch deren Absterben bestimmte

Bakteriengruppen in einem kurzen Zeitintervall stark beeinflusst werden. Dies betrifft vor

allem Bakterien des Phylums Bacteroidetes und der Roseobacter-Gruppe innerhalb der alpha-

Proteobakterien, die mit Änderungen der Phytoplanktonzusammensetzung korreliert waren.

Es zeigte sich auch, dass die Bakteriengemeinschaften der unterschiedlichen Kompartimente

(freilebend und Aggregat-assoziiert) auf unterschiedliche Weise beeinflusst werden. Eine

überraschend hohe Artenvielfalt von Bakterien der Roseobacter-Gruppe war auf Aggregaten

zu beobachten, die annehmen lässt, dass diese Gruppe für die Zersetzung partikulären

organischen Materials (POM) bisher unterschätzt wurde. Ebenso deutlich wurde, dass frei

lebende Bacteroidetes sehr schnell von Änderungen in der Umgebung beeinflusst werden und

ihre Rolle im Abbau von gelöstem organischem Material (DOM) wahrscheinlich als

wesentlich größer eingeschätzt werden kann als bisher angenommen. Die Bedeutung der


146

vorliegenden Arbeit liegt daher in der Beschreibung des Potentials einzelner

Bakteriengruppen, die im Ökosystem Wattenmeer extremen Einflüssen unterliegen und

sowohl Stabilität als auch Flexibilität besitzen. Die Auswirkung der einzelnen

Bakteriengruppen auf den gesamten Stoffumsatz ist daher komplex zu betrachten und sollte in

weiteren Untersuchungen besonderen Stellenwert einnehmen.

Hinweise über Veränderungen bakterieller Aktivität wurden sogar während eines Tidenzyklus

beschrieben (Grossart et al. 2004). Diese Beobachtung ließ die Frage entstehen, auf welcher

Zeitskala Reaktionen der Bakteriengemeinschaft nachweisbar sind. In Kapitel II war

ersichtlich, dass Bakteriengemeinschaften in wöchentlichen Zeitabständen starken

Änderungen unterliegen können. Die Abhängigkeit der Änderungen in der

Bakteriengemeinschaft von saisonalen Variablen, z. B. Sturmereignisse im Herbst und die

Phytoplanktonblüte im Frühjahr und Sommer, war bereits beschrieben worden (Stevens et al.

2005).

Daher wurde in Kapitel III anhand von Probenahmen zu den Kenterpunkten bei Hoch- und

Niedrigwasser sowie zum Strömungsmaximum und hoch sensitiver Nachweismethoden der

Einfluss der Tide auf die Zusammensetzung der Bakteriengemeinschaften zu

unterschiedlichen Jahreszeiten untersucht. Die Ergebnisse zeigen, dass wenige Änderungen

innerhalb der Bakteriengemeinschaften sogar im Gezeitenwechsel zu den Kenterpunkten

stattfinden. Einige Phylotypen waren stets nachweisbar und wurden schon beschrieben, z. B.

das RCA-Cluster und das WAC I-Cluster (Selje et al., 2004, Stevens et al, 2005), andere

Phylotypen jedoch erschienen nur kurzzeitig zu bestimmten Ereignissen. Diese Änderungen

waren im Wesentlichen durch den kombinierten Einsatz von RNA-basierter PCR und

spezifischen Primern für Bacteroidetes, alpha-Proteobakterien und Roseobacter detektierbar,

wodurch die Bedeutung dieser Bakterien für den gesamten Stoffumsatz als eher gering

einzuschätzen ist. Die Frage nach ihrer Funktion bleibt dennoch interessant, da es sich

innerhalb der Bakteriengemeinschaft um funktionell wichtige Prozesse handeln könnte, die zu

bestimmten Zeitpunkten relevant sind. Darüber hinaus war auffällig, dass die Bandenmuster

der aktiven Bakterien (RNA-basiert) stark abwichen von den Bandenmustern der als häufig

anzunehmenden Bakterien (DNA-basiert). Dies lässt den Rückschluss zu, dass die

Anwendung RNA-basierter Methoden bei weiteren Untersuchungen im Wattenmeer dringend

notwendig ist, um Korrelationen zwischen Aktivität und Phylogenie herstellen zu können.

Große Bedeutung hat die Anwendung der FISH- und CARD-FISH Methode, durch die

erstmalig die Abundanz der verschiedenen Bakteriengruppen in der Wassersäule des


147

Wattenmeeres und damit auch der jeweilige potentielle Anteil der Gruppen am Stoffumsatz

gezeigt werden konnte. Die Verknüpfung zu spezifischen tidalen Ereignissen war trotz der

hochauflösenden CARD-FISH nicht erkennbar, jedoch zeigte sich, dass in den hoch

produktiven Frühjahrs- und Sommermonaten wesentlich höhere Varianz innerhalb der

Bakteriengruppen auftrat, was ebenfalls auf erhöhte Aktivität der Organismen schließen

hinweist. Dies bedarf ebenfalls weiterer Untersuchungen, z. B. in Kombination mit der Micro-

FISH Methode, um diese Aussagen stützen zu können.

Kapitel IV zeigte, dass die Actinobakterien im Wattenmeer unerwartet hohe Abundanz und

Artenvielfalt aufweisen. Die Frage nach der ökologischen Bedeutung dieser Organismen für

Küstengebiete rückt daher immer mehr in den Vordergrund und stellt eine interessante

Perspektive für weitere Forschungsarbeiten dar. Auch für diese Gruppe zeigte sich, dass die

Einteilung in die Kompartimente frei lebend und aggregat-assoziiert essentiell ist, um weitere

Rückschlüsse ziehen zu können. Beide Fraktionen zeigten Unterschiede in der

Zusammensetzung der Bakteriengemeinschaft und sind daher auch gesondert zu betrachten.

Besonders interessant ist die klare Unterscheidung von Actinobakterien, die im Süßwasser

leben, und den halotoleranten marinen Vertretern im Wattenmeer. Actinobakterien wurden

bisher durch Produktion von Sekundärstoffen in biotechnologischen Fragestellungen

fokussiert, aber auch im ökologischen Sinne ist diese Eigenschaft untersuchenswert.

Besonders auf Aggregaten können diese als Sekundärstoffe bezeichneten Botenstoffe

besondere Bedeutung für Kommunikation oder Abwehr innerhalb der

Bakteriengemeinschaften einnehmen. Als Hauptursache für die Artenvielfalt und Abundanz

der Actinobakterien in diesem schwebstoffreichen Grenzbereich zwischen Land und Meer

kann jedoch der Abbau komplexer organischer Verbindungen, z. B. Huminstoffe,

angenommen werden.

Der Austausch von Wasserkörpern zwischen dem Wattenmeer und der offenen Nordsee und

damit verbundene Stoffflüsse sind lange Zeit Mittelpunkt sedimentologischer und weiterer

geochemischer Untersuchungen gewesen. Inwiefern Aggregat-assoziierte oder auch frei

lebende Bakterien durch diese Prozesse beeinflusst sind und welchen Einfluss

standortgebundene Bedingungen auf die Bakteriengemeinschaften haben, zeigen die

Ergebnisse aus Kapitel V.


148

Trotz standortspezifischer Unterschiede, die durch die Bandenmuster der partikel-assoziierten

Bakteriengemeinschaften reflektiert wurden, war eine homogene Verteilung frei lebender

Bakteriengemeinschaften zu beobachten. Die phylogenetischen Stammbäume ergaben große

Ähnlichkeit der partikulären Bakteriengemeinschaften des ostfriesischen und nordfriesischen

Wattenmeeres. Es konnten Indikatororganismen ausgemacht werden, die ausschließlich

Aggregat-assoziiert leben und somit den Verlauf partikulären Materials aus den

Wattenmeeren und von resuspendiertem Sediment in der Wassersäule aufzeigen.

Überraschend waren auch hier die eindeutige Trennung der Kompartimente, und die große

Stabilität innerhalb der alpha-Proteobakterien bzw. der Roseobacter-Gruppe. Nicht

veröffentlichte Daten, die im Rahmen dieser Arbeit erhoben wurden, ergaben für die frei

lebenden Roseobacter nahezu identische DGGE-Bandenmuster, unabhängig von der

Jahreszeit und vom Standort. Die ökologische Bedeutung dieser Gruppe für den Umsatz von

gelöstem organischen Kohlenstoff (DOC) ist daher als essentiell anzunehmen und bedarf

dringend einer weiteren Erforschung.

Zusammenfassend ergibt sich daher aus dieser Arbeit ein komplexes Bild mit vertieften

Erkenntnissen über die Zusammensetzung der Bakteriengemeinschaften im ostfriesischen

Wattenmeer und der Deutschen Bucht. Die sensitiven Untersuchungsmethoden konnten

zeigen, welche Einflussfaktoren in diesem als persistent bezeichneten Ökosystem eine

wesentliche Rolle einnehmen und bilden daher die Grundlage für weitere Forschungsarbeiten.

Aktivitätsmessungen und Erhebung des genetischen Potentials für Stoffwechselwege stellen

daher eine geeignete Ergänzung dar, die durch weitere Arbeiten im Rahmen des

Forschergruppenverbundes erhoben werden können. Intensive Datenerhebung von

bakterieller Biomasseproduktion findet derzeit in stündlichen bis saisonalen Zeitskalen statt,

die gleichzeitige Etablierung der Micro-CARD-FISH stellt eine weitere, höchst effektive

Möglichkeit zur Aktivitätsmessung im Wattenmeer dar. Die Überprüfung der Ergebnisse über

experimentelle Ansätze ist derzeit ebenfalls in der Ausführung, und die Verknüpfung zum

natürlichen System wird über mathematische Modelle gelenkt. Die Metagenomik bleibt

vorerst ein zukunftsträchtiger Ausblick, wird aber im Zuge der Fortschritte in der molekularen

Ökologie ein logischer Schritt zur weiteren Erforschung der Bakteriengemeinschaften in der

Wassersäule des Wattenmeeres und der Deutschen Bucht darstellen.

Erklärung

Hiermit bestätige ich, dass ich die vorliegende Dissertation selbständig verfasst und nur die

angegebenen Quellen und Hilfsmittel verwendet habe.

Weiterhin erkläre ich, dass diese Dissertation weder in ihrer Gesamtheit noch in Teilen einer

anderen wissenschaftlichen Hochschule zur Begutachtung in einem Promotionsverfahren

vorliegt oder vorgelegen hat.

Oldenburg, den

Danksagung

Mein besonderer Dank gilt Prof. Meinhard Simon für das Vertrauen, die Unterstützung und

die große Flexibilität, vor allem zum Ende der Fertigstellung dieser Arbeit. Diese

Eigenschaften sind keineswegs selbstverständlich und ich bin dankbar, dass ich diese positive

Erfahrung machen durfte! Vielen Dank an dieser Stelle auch an Prof. Heribert Cypionka, der

unkompliziert die Aufgabe des Gutachters übernommen hat. Für ihn habe ich mich besonders

um Prägnanz bemüht - ich hoffe, es ist gelungen.

Für Zielstrebigkeit, fachliche Diskussion und einfach immer wohltuenden freundlichen Rat

möchte ich Dr. Thorsten Brinkhoff danken. Er hat mich mehr als einmal vor dem Verzetteln

bewahrt und ist mir ein großes Vorbild im Umgang mit Kollegen. Vielen Dank auch für die

große Flexibilität zum Ende meiner Arbeit!

Der gesamten Arbeitsgruppe, besonders Andrea Schlingloff und Birgit Kürzel, möchte ich

danken für die Geduld, Offenheit und Hilfsbereitschaft. Heike Stevens bin ich mehr als

dankbar für 2500000000 motivierende Gespräche und ihre erfrischende Art, das Leben positiv

zu sehen. Und für die Fahrt im Schäbi Pop durch Yucatan! Katja Walther und Beate Köpke

danke ich für Fachgespräche zu jeder Tages- und Nachtzeit und für die Freundschaft, die eine

sehr wichtige Begleitung für mich war und ist.

Die größte Unterstützung fand ich bei meiner Familie, die mir jederzeit zur Seite stand und

mir gezeigt hat, wie wichtig dieser Halt für mich ist. Ihr habt alle einen großen Teil zu dieser

Arbeit beigetragen! Das werde ich nie vergessen und daher widme ich Euch meine Arbeit aus

tiefstem Dank.

Meinem Freundeskreis, für den das Wort „Doktorarbeit“ mittlerweile fest mit Begrüßungs-

und Abschiedsformeln verknüpft ist, danke ich für unendliche Ausdauer und Geduld, für

seelischen Ausgleich, für tolle Gespräche, und fürs Dasein. Es ist geschafft!!! Ihr könnt

wieder ans Telefon gehen!

Lebenslauf

Beate Rink, geboren am 23.01.1974 in Bremerhaven

Schulausbildung

1980-1984 Grundschule

Friedrich-Ebert-Schule, Bremerhaven

1984-1986 Orientierungsstufe

Wilhelm-Raabe-Schule, Bremerhaven

1986-1990 Gymnasium

Wilhelm-Raabe-Schule, Bremerhaven

1990-1993 Gymnasiale Oberstufe

Geschwister-Scholl-Schule, Bremerhaven

Leistungskurse: Latein, Kunst

Abschluß: Abitur

Berufsausbildung

1993-1995 Ausbildung zur Industriekauffrau

Fa. Schottke GmbH & Co. KG, Bremerhaven

Abschluß: Industriekauffrau

1995-1997 Biologiestudium

Universität Regensburg

1997-2001 Biologiestudium

Carl-von-Ossietzky-Universität, Oldenburg

Hauptfächer: Mikrobiologie, Genetik, Biochemie

Abschluß: Diplom

Thema Diplomarbeit: „Besiedelung und Abbau von Fucus-detritus durch

heterotrophe marine Bakterien“

2001-2006 Wissenschaftliche Angestellte am Institut für Chemie und Biologie des Meeres

Carl-von-Ossietzky-Universität, Oldenburg

Arbeitsgruppe „Biologie geologischer Prozesse“, Prof. Meinhard Simon

Documents

Struktur und Dynamik heterotropher Bakteriengemeinschaften ...oops.uni-oldenburg.de/42/1/rinstr07.pdf · bacteria affiliated to the Bacteroidetes phylum, gamma- and delta-Proteobacteria