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R E S E A R C H A R T I C L E

Vertical distributionofmetabolicallyactive eukaryotes in thewatercolumnand sedimentsoftheBlackSeaMarco J.L. Coolen1 & Galina Shtereva2

1Marine Chemistry and Geochemistry Department, Woods Hole Oceanographic Institution, Woods Hole, MA, USA; and 2Bulgarian Academy of

Sciences, Institute of Oceanology (IO-BAS), Varna, Bulgaria

Correspondence: Marco J.L. Coolen, Marine

Chemistry and Geochemistry Department,

Woods Hole Oceanographic Institution, 360

Woods Hole Road, Woods Hole, MA 02543,

USA. Tel.: 11 508 289 2931; fax: 11 508 457

2164; e-mail: [email protected]

Received 8 January 2009; revised 28 July 2009;

accepted 29 July 2009.

Final version published online 1 September

2009.

DOI:10.1111/j.1574-6941.2009.00756.x

Editor: Riks Laanbroek

Keywords

Black Sea; suboxic; sulfidic; 18S rRNA gene

transcripts; DGGE; active eukaryotes.

Abstract

Recent DNA-based phylogenetic studies have reported high eukaryotal diversities

in a wide range of settings including samples obtained from anoxic environments.

However, parallel RNA-based surveys are required in order to verify whether the

species detected are in fact metabolically active in such extreme environments. The

Black Sea is the World’s largest anoxic basin but remains undersampled with

respect to molecular eukaryotic diversity studies. Here, we report the distribution

of active eukaryotes (18S rRNA-based survey) along a vertical nutrient and redox

gradient in the water column and surface sediments of the Black Sea. A wide

variety of eukaryotes were active in suboxic deep waters. Notably, certain species

were active but escaped detection during a parallel 18S rDNA survey. The 18S

rDNA survey from surface sediments yielded taxa of pelagic origin but none of

these were identified from the water column at the time of sampling. Our data

also indicate that gene transcripts do not always provide unequivocal proof

that active microorganisms are indigenous to a specific position in an environ-

mental gradient, because certain zoo- and phytoplankton species were still viable

with detectable 18S rRNA in up to 300-year-old sulfidic sediments that

underlie �830 m of sulfidic waters.

Introduction

Over the past several years, culture-independent molecular

surveys using small subunit (SSU) rDNA sequences as

phylogenetic markers have increasingly been used to study

the environmental diversity of microbial eukaryotes. Pio-

neering studies have revealed an unexpectedly high diversity

of microbial eukaryotes in the deep sea (Lopez-Garcia et al.,

2000) and in surface waters at different oceanic settings

(Dıez et al., 2001; Moon-van der Staay et al., 2001). Some of

the unique phylotypes from these studies represent novel

lineages within known eukaryotic groups, such as alveolates

and stramenopiles. In addition, 18S rDNA surveys have also

been used to study microbial eukaryotic diversity in extreme

environments, such as deep-sea hydrothermal vents (Edg-

comb et al., 2002; Lopez-Garcia et al., 2003), acidic and

iron-rich rivers (Zettler et al., 2002; Zettler et al., 2007), and

various suboxic or anoxic marine settings, including marine

and lacustrine surface sediments (Dawson & Pace, 2002),

salt marsh environments (Stoeck et al., 2003), submarine

caldera around fumaroles (Takishita et al., 2005), deep-sea

methane cold-seep sediments (Takishita et al., 2007), strati-

fied marine basins (Stoeck et al., 2003), and fjords (Behnke

et al., 2006; Zuendorf et al., 2006). The latter studies revealed

many novel phylotypes at high-taxonomic levels, although it

has been argued that some of these phylotypes could

represent artifacts such as chimeric sequences (e.g. Behnke

et al., 2006). Another issue is that such DNA-based commu-

nity analyses detect microorganisms irrespective of their

viability or metabolic activity (Stoeck et al., 2007). The

ability to identify living or active microbial eukaryotes is

particularly important for settings where chemical environ-

ments and microbial populations can change on a very small

spatial scale (Stoeck et al., 2007), such as along the redox

gradient in stratified, oxygen-depleted environments

(Stoeck et al., 2003; Behnke et al., 2006).

The identification of active microbial populations within

environmental samples can be addressed by targeting SSU

rRNA gene transcripts directly instead of DNA. This

approach has mainly been used to identify active

FEMS Microbiol Ecol 70 (2009) 525–539 c� 2009 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved

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prokaryotes in environmental samples (MacGregor et al.,

2001; Norris et al., 2002; Mills et al., 2004) and only recently

to detect active microbial eukaryotes (Coyne & Cary, 2005;

Stoeck et al., 2007; Alexander et al., 2009). rRNA is an

essential structural component of the ribosome, which is the

organelle responsible for protein synthesis in all prokaryotes

and eukaryotes. The number of cellular ribosomes, and thus

also the rRNA content, increases with growth rate and

decreases with starvation (Wagner, 1994; Muttray & Mohn,

1998; Buckley & Szmant, 2004; Chicharo & Chicharo, 2008;

Hawkins et al., 2008). Therefore, rRNA extracted from

environmental samples serves as a phylogenetic marker for

the identification and relative abundance of metabolically

active members within complex microbial communities.

Stoeck et al. (2007) constructed rRNA- and rDNA-based

clone libraries derived from an anoxic water sample from

the Framvaren Fjord (Norway) to compare the diversity and

relative abundance of active vs. inactive marine microeukar-

yotes. This study revealed 84 unique phylotypes of which

27% occurred in both libraries, 25% occurred exclusively in

the rRNA library and 48% occurred exclusively in the rDNA

library. Furthermore, phylotypes of phototrophic dinofla-

gellates, uncultured marine alveolates group I, and various

parasites were exclusively detected in the rDNA library and

represented nonindigenous members of the anoxic micro-

eukaryote community below the sulfidic chemocline

(Stoeck et al., 2007). Even though the analyzed clone library

was extensive (600 clones) and revealed many unique

phylotypes, only one depth was analyzed in the above study.

In addition, although the community structure below the

chemocline in stratified waters is likely to remain stable as

long as the stratification is not disturbed, pelagic samples

provide only a snapshot of photic zone-derived cells in

transit to the sediment (Coolen et al., 2007). Benthic rRNA

libraries in these settings would provide a more complete

picture of the capacity of organisms stemming from the

oxygenated part of the photic zone to remain active or viable

in suboxic or fully anoxic environments for substantial

periods of time. For example, another recent study has

shown that viable dinoflagellate cysts in anoxic Delaware’s

Inland Bays surface sediments could be identified using

PCR-based approaches from 18S rRNA libraries and even

from much more labile mRNA libraries (Coyne & Cary,

2005). This suggests that such gene transcripts could remain

detectable in resting stages of planktonic species not indi-

genous to anoxic water layers for substantial periods of time.

The Black Sea, the world’s largest permanently stratified

basin, is devoid of oxygen and contains abundant sulfide

from a depth of about 100 m to the seafloor at 2200 m. A

20–30-m-deep suboxic layer, depleted in both O2 and

sulfide, overlies the sulfide zone (Jørgensen et al., 1991).

This stable, vertically expanded redox gradient provides an

opportunity to study the distribution of microbial commu-

nities within different chemical environments, but thus far

remains undersampled with respect to eukaryotic diversity.

In the present study, we performed a high spatial resolution

analysis of the eukaryotic diversity (18S rDNA survey) and

sought to determine whether these organisms were metabo-

lically active (18S rRNA-based library) along a vertical

gradient of O2, H2S, and nutrients in the western Black Sea.

In addition, we performed a parallel survey on underlying

surface sediments to identify active benthic eukaryotes and,

more importantly, to establish whether eukaryotes derived

from the photic zones remain viable in this highly sulfidic

depositional setting.

Materials and methods

Water column chemistry and physical conditions

A Conductivity-Temperature-Depth profiler (CTD) was

deployed to measure conductivity, temperature, salinity,

fluorescence, dissolved oxygen (SBE-43 oxygen sensor; Sea-

Bird Electronics, Bellevue, WA) and to collect water to

measure N-species (nitrite, nitrate, ammonia), phosphate,

and sulfide. This effort revealed the exact depths of the

chlorophyll maximum, the upper and lower nitrite peaks,

the suboxic zone with o 0.2 mL L�1 of oxygen, as well as the

sulfidic chemocline. This information was used to select the

depths for the collection of particulate organic matter

(POM) for the molecular survey.

Nutrients (nitrite, nitrate, ammonia, and phosphate)

were measured spectrophotometrically according to Grassh-

off et al. (1983). Hydrogen sulfide was measured photome-

trically according to Cline (1969).

Sampling

Water samples were collected from 19 depths in the stratified

Black Sea (42146.56900N : 28140.64700E) aboard the R/VAkade-

mik (Institute of Oceanology, Bulgarian Academy of Sciences;

IO-BAS) using a SBE911 plus CTD (Sea-Bird Electronics)

equipped with twelve 5-L Niskin bottles. These depths in-

cluded the chlorophyll maximum at 22, 40, 60–115 m (5-m

intervals along the oxygenated mixolimnion), 120–135 m

(5-m intervals within the suboxic zone with o 0.2 mL L�1

oxygen), and within the sulfidic chemocline (150 and 160 m).

We took precautions to prevent contact of the oxygen-

depleted and sulfidic waters with the atmosphere by keeping

the water inside the Niskin bottle until cells in 3–5 L of water

were collected onto 142-mm, 0.2-mm pore size, polycarbonate

filters (Millipore, Billerica, MA). The water was directly

filtered from the Niskin bottles via a closed in-line tubing

system connected to the lower outlet of each Niskin

bottle under gentle pressure (o 50 mL min�1) provided by a

peristaltic pump (Masterflex, Cole and Palmer, Vernon Hills,

IL). The total water depth at this location was 971 m.

FEMS Microbiol Ecol 70 (2009) 525–539c� 2009 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved

526 M.J.L. Coolen & G. Shtereva

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Fifty-centimeter-long undisturbed cores of laminated

sediments (Fig. 1b) were obtained at the same location

using a MC800 multicorer (Ocean Instruments, San Diego,

CA). One subcore (Core MC06) was subsectioned in 2-cm

intervals aboard ship and sections from the upper 8 cm of

undisturbed laminated sediments, including the top fluffy

layer, were used for subsequent extraction of nucleic acids as

described below.

Simultaneous DNA and RNA extraction andcDNA synthesis

POM

Directly after filtration, the filters were placed in a 15-mL

sterile conical centrifuge tube with a mixture of 4-mL

denaturing buffer (Hurt et al., 2001), 50mL 2-mercapto-

ethanol, and 2 mL of nucleases-free low-binding zirconium

beads with a diameter of 200mm (OPS Diagnostics, Leba-

non, NJ). Cells on the filter were mechanically disrupted to

release the nucleic acids by bead beating for 5 min at

maximum speed using the Vortex Genie 2s Vortex and

adapter (MoBio, Carlsbad, CA). The presence of the dena-

turing buffer and 2-mercaptoethanol ensured instant inacti-

vation of the nucleases present. Then the mixtures were

immediately stored at � 80 1C or colder until further

analysis. At the Woods Hole Oceanographic Institution

(WHOI), all steps involved in the extraction and purifica-

tion of nucleic acids were performed inside a HEPA-filtered

fume hood, which was UV-sterilized and cleaned with

Eliminase (Decon Laboratories, King of Prussia, PA) to

destroy nucleases before use. Additional precautions, such

as using filter tips during pipetting, using only RNAse-free

reagents and disposable plastic ware were also taken. Vinyl

gloves were worn at all times and cleaned in between steps

with Eliminase. Then, each deeply frozen filter/buffer/bead

mixture was transferred to precombusted (225 1C for 8 h)

and deeply frozen (� 80 1C) mortars and ground with a cold

pestle upon the thawing of the mixture. Circular movements

during grinding were omitted to minimize shearing of the

nucleic acids. The mixture was transferred into 50-mL-

sterile conical centrifuge tubes and an equal volume

(�4 mL) of phenol : chloroform : isoamylalcohol (PCI),

25 : 24 : 1, pH 8 (Ambion, Austin, TX) was added. After

vigorously vortexing, the mixture was placed on ice for

15 min to ensure the complete disassociation of nucleopro-

tein complexes, and centrifuged at 4 1C for 15 min at

10 000 g. The aqueous phase was transferred to a new sterile

tube and the nucleic acids were precipitated with

0.1 volume of 5 M NaCL (Ambion) and 1 volume of

isopropanol. After centrifugation at 10 000 g for 20 min at

room temperature, the air-dried pellet was dissolved in RNA

Storage Solution (Ambion).

Surface sediments

Total nucleic acids were extracted at 2-cm intervals from the

top 8 cm of sediment. Directly after sectioning, 1.5 mL of

sediment was transferred to 5-mL cyrovials and mixed with

equal volume of denaturing buffer (Hurt et al., 2001), 20 mL

of 2-mercaptoethanol, and 0.75 mL of zirconium beads.

Cells in the sediment were then immediately mechanically

disrupted to release the nucleic acids and to inactivate the

nucleases by bead beating as described above. The cyrovials

were stored in liquid nitrogen on board the R/V Akademik,

Fig. 1. (a) Sampling station in the western Black Sea off the coast of Varna, Bulgaria during cruise BS/AK06 (August 28–September 4, 2006) on the R/V

Akademik. All data presented in this report were obtained from filtered POM and multicore MC06 at Station 5 (Sta 5, 971 m). (b) MC06 with

undisturbed laminated anoxic top sediments.


527Viable and active eukaryotes in the Black Sea

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transferred on dry ice to WHOI, and stored at � 80 1C until

further analysis. At WHOI, the frozen mixture underwent

the same grinding procedure in sterile and deeply frozen

mortars as described above, but followed an additional

extraction step with 8 mL of extraction buffer as described

by Hurt et al. (2001). Following centrifugation (10 min,

10 000 g), total nucleic acids were extracted from the super-

natant using PCI and the additional steps described above.

The quality of the RNA and DNA extracts was verified by

agarose gel electrophoresis. The total nucleic acid extracts

from the POM and sediments were used for the 18S rDNA

survey.

Preparation of DNA-free RNA andcDNA synthesis

For the 18S rRNA survey, 0.1 volume of the extracted total

nucleic acids was treated for 30 min at 37 1C with 1 U DNAse

and 1� buffer using the DNA-freeTM kit (Ambion) followed

by inactivation of the DNAse using 0.1 volume of the

provided DNAse inactivation reagent. Before PCR, DNA-

free RNA samples were randomly transcribed into cDNA

using the iScript cDNA synthesis kit (Bio-Rad, Hercules,

CA). The 20 mL reaction mixtures contained RNAse-free

water, 1 U iScript reverse transcriptase, 1� iScript Reaction

Mix, 50 mg bovine serum albumin (BSA; Ambion), and

0.1 volume of DNA-free RNA sample. The reverse transcrip-

tase (RT) reactions were performed in a Mastercycler

(Eppendorf, Westbury, NY) and included incubation for

5 min at 25 1C, 30 min at 42 1C, and 5 min at 85 1C. The

latter cDNA served as template for PCR to study the diversity

of the metabolically active eukaryotes in the Black Sea.

PCR amplification of eukaryote 18S rDNA andreverse transcribed 18S rRNA

Each amplification reaction contained 0.25 mM of each

deoxynucleotide (dNTP), 4.5 mM of MgCl2, 0.5� (from

10 000� concentrated stock) SYBRsGreen I (Invitrogen,

Carlsbad, CA), 50 mg of BSA (Ambion), 5mL of

10� PicoMaxxTM reaction buffer (Stratagene, LaJolla, CA),

2.5 U of DNA polymerase (PicoMaxxTM high fidelity PCR

system; Stratagene), 0.2mM of primers (Thermo-Fisher,

Ulm, Germany), and 10 ng of DNA or cDNA template.

Partial (560-bp long) eukaryote 18S rDNA (or reverse

transcribed 18S rRNA) was PCR amplified using primer set

Euk1A and Euk516r-GC (Set A) as described previously

(Dıez et al., 2001) The reaction mixtures were adjusted to a

final volume of 50 mL with nucleic acids and nuclease-free

water (Ambion). Each PCR amplification series included a

reaction without a DNA template, which served as a control

for contaminations during the pipetting of the reaction

mixture components. For each cDNA sample, a parallel

reaction with 0.2 vol% of digested RNA was subjected to

PCR as a control for the presence of traces of incomplete

digested DNA in the various RNA templates used for cDNA

synthesis.

All reactions were performed in a Realplex quantitative

PCR cycler (Eppendorf) and involved initial denaturing

(4 min at 95 1C), followed by 38 cycles including denaturing

(30 s at 94 1C), primer annealing (40 s at 64 1C), primer

extension (60 s at 72 1C), and imaging of newly formed

fluorescent (SYBRsGreen I labeled) double-stranded

DNA (80 1C for 20 s). All DNA and cDNA samples

were then subjected again to real-time quantitative PCR

but this time all reactions were stopped at the end of the

exponential phase (i.e. after 25–32 cycles). This prevented

overamplification of individual 18S rDNA fragments

to allow semi-quantitative analysis by denaturing gradient

gel electrophoresis (DGGE; Schafer & Muyzer, 2001) as

outlined below.

Phylogenetic analysis of sequenced DGGE bands

For the majority of the samples, 100 ng of PCR-amplified

partial eukaryote 18S rRNA and RT-PCR amplified 18S

rRNA (600 bp including the 40-bp-long GC clamp) was

separated by DGGE (Schafer & Muyzer, 2001). The poly-

acrylamide gels (6%, w/v) contained a denaturing gradient

of 20–50% (with 100% denaturant equaling 7 M urea and

40% formamide). Gels were run for 15 h at 5 V cm�1 and

60 1C using a PhorU2 system (Ingeny, Leiden, the Nether-

lands). Afterwards, the gels were stained for 20 min by

covering the gels twice with 10 mL of 1�TAE buffer

(pH 8.3), containing 2 mL undiluted SYBRsGold (Invitro-

gen) followed by destaining for 60 min in 1�TAE buffer

(pH 8.3). In order to prevent DNA damage by UV, we used a

Dark Reader (Clare Chemicals Research Inc., Dolores, CO),

which uses visible light instead of UV in order to visualize

the SYBRsGold-stained DNA. Digital gel images were made

using a Foto/Analysts Express System (Fotodyne, Hartland,

WI) and IMAGEJ software. TOTALLAB TL100 v2006 1D-gel

analysis software (Nonlinear Dynamics, Durham, NC) was

used to determine the pixel density and vertical position of

each band. This information was used to determine the

relative abundance of each band within a given sample and

to identify the exact vertical position of each band in order

to characterize unique vs. identical bands between samples

(e.g. Dıez et al., 2001). DGGE bands were sliced from the gel

with a sterile scalpel and the DNA of each sliced gel fragment

was eluted in 75 mL sterile 10 mM Tris-HCl (pH 8.0) by

incubation for 48 h at 2 1C. One microliter of the eluted 18S

rDNAs (approximately 107 copies) was reamplified using

25 cycles and the primer combinations listed above, but this

time without the GC clamp. These amplicons served as DNA

for the subsequent cycle sequencing reactions. PCR reagents

were as described above but without the addition of



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SYBRsGreen I or extra MgCl2. All reamplification reactions

were performed in a Master cycler (Eppendorf) and in-

volved initial denaturing (4 min at 95 1C), followed by

25 cycles including denaturing (30 s at 94 1C), primer

annealing (40 s at 58 1C), and primer extension (12 s at

72 1C). A final extension step was set at 72 1C for 30 min.

The quality of each sequencing template was checked by

agarose gel electrophoresis. The DNA concentration of each

amplicon was measured fluorometrically using picogreen

staining (e.g. Coolen et al., 2006). Thirty nanograms of

reamplified DGGE bands were submitted to Agencourt

(Beverly, MA) for subsequent bidirectional cycle sequencing

with primers EukA and Euk516r.

Sequences have been analyzed using the ARB software

package (version December 2007) (Ludwig et al., 2004) and

the corresponding SILVA SSURef 96 database (Pruesse et al.,

2007). After importing, all sequences were automatically

aligned according to the SILVA SSU reference alignment.

Manual refinement of the alignment was carried out taking

into account the secondary structure information of the

rRNA. Then, the phylogenetic bootstrap trees (1000 replica-

tions) were first reconstructed based on 1200-bp-long avail-

able sequences of the closest relatives using the neighbor-

joining method (ARB). The shorter aligned environmental

18S rDNA sequences from this study were inserted after-

wards using the parsimony interactive tool implemented in

the ARB software package without changing overall tree

topology. Sequences obtained in this study have been

deposited in the NCBI sequence database under accession

numbers GQ402464–GQ402489.

Results and discussion

Physical parameters and location of the suboxiczone, nutrients, and onset of sulfide

The upper 160 m of the Black Sea are brackish with salinities

between 17.2 psu at the surface and 20.6 psu toward the

sulfidic chemocline (Fig. 2). The surface water temperature

was 22.5 1C at the time of sampling, with a steep drop to

15.3 1C at the thermocline at 25 m, followed by relatively

constant temperatures (8.1� 0.6 1C) between 31 and 160 m

(Fig. 2). Fluorescence data show that a chlorophyll max-

imum was present just above the thermocline at 22 m,

whereas chlorophyll concentrations dropped close to zero

values o 80 m (Fig. 2). A small increase in fluorescence was

observed at the top of the suboxic zone at 120 m.

The suboxic zone with o 0.2 mL L�1 (i.e. o 10mM)

oxygen and undetectable sulfide concentrations (o 0.2mM)

(Murray et al., 1989; Codispoti et al., 1991) at Station 5 was

found between 120 m (st = 15.62) and 139 m (st = 15.90). At

140 m sulfide became detectable, showing 0.28mM, and

increased to 6mM at 150 m (st = 16.00) and 12.5mM at

160 m (st = 16.10) (Fig. 2).

Ammonia concentrations dropped from 8.5mM at

�160 m to undetectable levels (o 0.2mM) at the base of

Fig. 2. Water-depth profiles with physical parameters [temperature, salinity, st values, and fluorescence (chlorophyll a concentration)] as well as

chemical parameters (oxygen and sulfide) (left panel), nutrients (N-species and phosphate) (second panel from the left), and relative quantitative

distributions of 18S rDNA (closed circles) vs. 18S rRNA (open circles) of ‘total’ vs. active pelagic copepod, tunicate, haptophyte, dinoflagellate, ciliate,

and unclassified species. The suboxic zone with o 0.2 mL L�1 of dissolved oxygen and undetectable sulfide is marked in dark gray. The upper nitrite

maximum in the oxygenated waters and the lower nitrite maximum in the suboxic zone are marked in light gray.



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the suboxic zone. A small nitrite maximum (0.15mM) was

found in the suboxic zone at 130 m (Fig. 2). At depths

between �100 and 120 m where oxygen concentrations

reached up to 5 mL L�1, a second (upper) nitrite maximum

occurred, which agrees well with previous findings (Ward &

Kilpatrick, 1990; Murray & Yala-del-Rio, 2006). A nitrate

peak with up to 4mM of nitrate was observed between �80

and 120 m. Phosphate concentrations were highest at the

bottom of the suboxic zone and in the sulfidic waters down

to 160 m (�4 mM) (Fig. 2).

Distribution of metabolically active eukaryotesalong the environmental gradients

Metabolically active eukaryotes were identified within

19 POM samples collected along a vertical gradient (salinity,

oxygen, hydrogen sulfide, and nutrients), and within the

upper 8 cm of sulfidic sediments that underlie 830 m of

sulfidic water at the sampling location in the Black Sea. The

active population was studied from reverse transcribed and

PCR-amplified (two-step RT-PCR) 18S rRNA whereas com-

bined molecular signatures of both metabolically active and

inactive populations were studied from PCR-amplified

(genomic) 18S rDNA (i.e. 18S rRNA- vs. 18S rDNA-based

libraries). All amplicons were subjected to DGGE (Fig. 3a).

Unfortunately, we lost the DNA extract from 80 m during

the purification procedure and, as a result, no 18S rDNA

amplicons could be generated from that sample (Fig. 3a).

The corresponding DNAse-treated 18S rRNA extract from

the 80 m sample could subsequently be analyzed by RT-PCR

(Fig. 3a). Total DNA and RNA was successfully extracted

from 115, 75, 135, and 160 m, but the samples from these

depths contained fewer 18S rDNA (i.e. 115a) or 18S rRNA

(75b, 135b, 160b) template for PCR and yielded less

amplicons for subsequent DGGE analysis as compared with

other samples within the environmental gradient (Fig. 3a).

Fifty-six numbered DGGE bands were excised from the

DGGE gels and 53 of the excised bands were successfully

sequenced. These sequences represented 18 unique phylo-

types (based on a 99% sequence similarity cutoff) (Fig. 4).

Fig. 3. DGGE with PCR-amplified 18S rDNA and reverse transcribed and PCR-amplified 18S rRNA (from cDNA pool) from (a) water column POM

samples and (b) sulfidic sediments from MC06. Fifty-six bands were excised from the gel and 53 were successfully sequenced and revealed 18 unique

phylotypes, denoted 1–18 and with different colors for each taxonomic group. Bands from different samples that melted at identical heights in the gel

and appeared to represent the same phylotype carry the same number (1–18). Unsuccessfully sequenced bands are indicated with ‘X’ and chimeric

sequences with ‘C’. Sampling depths (in meter for POM samples and in centimeter for the sulfidic sediments) are located above panels (a) and (b),

respectively. Lanes that contained 18S rRNA amplicons are denoted with ‘a’ and lanes that contain reverse-transcribed and PCR-amplified 18S rRNA are

denoted with ‘b’. PM, position marker in the gel, not a sample; gDNA, genomic DNA that contains the 18S rRNA genes; cDNA, cDNA from total pool of

reverse transcribed RNA. The total DNA from 80 m was lost during extraction and, for that reason, did not result in 18S rDNA amplicons (indicated with

red ‘x’ in 80 m lane ‘a’).



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Fig. 4. Phylogenetic tree showing the

relationship of eukaryote 18S rDNA and

18S rRNA sequences retrieved from Black

Sea water column and surface sediments

to reference sequences obtained from the

NCBI database (http://www.ncbi.nlm.nih.

gov). The eukaryote sequences from POM

and MC06 samples were determined

from the DGGE represented in Fig. 3.

Phylotypes are indicated as, for example,

‘3_ BSA6S5Euk’, which stands for DGGE

band representing the unique eukaryotic

phylotype # 3 obtained from the Black Sea

with the RV Akademik in 2006, Station 5.

Bar indicates 0.1 fixed point mutations per

nucleotide. Numbers at nodes indicate

bootstrap values out of 1000 replications

for phylogenetic trees calculated by

Felsenstein correction and Parsimony

methods.



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The DGGE bands that represented unique phylotypes are

numbered 1–18 in the DGGE (Fig. 3a). These phylotypes

comprised copepods (four phylotypes; 12_BSA6S5Euk,

16–18), tunicates (phylotype 11), haptophytes (phylotype

1), dinoflagellates (eight phylotypes; 3, 6, 7, 8, 9, 10, 13, 14),

and ciliates (phylotypes; 2 and 4) as well as a sequence

(phylotype 15), which did not fall into established taxo-

nomic groups but which showed up to 96% sequence

similarity to 18S rDNA sequences recovered from the

sulfidic waters of Framvaren Fjord (Fig. 4, Table 1) (Behnke

et al., 2006).

Copepods

A copepod (phylotype 12) with 100% sequence similarity to

partial 18S rDNA of Pseudocalanus elongatus (Fig. 4, Table 1)

was identified from both 18S rDNA/rRNA libraries in the

oxygenated part of the mixolimnion (between 40 and 95 m)

(Fig. 3a) and represented up to 40% of the total reverse-

transcribed and PCR-amplified 18S rRNA pool (Fig. 2).

Pseudocalanus elongatus is known to be highly abundant

throughout the Black Sea (Niermann & Greve, 1997) and

makes up �40% of the zooplankton in autumn within the

offshore regions (Unal et al., 2006). It is a cold-water species

with an upper temperature limit of occurrence of 13 1C,

which would explain its absence in the chlorophyll max-

imum (Figs 2 and 3a) that was 22.2 1C at the time of

sampling.

The sequence of phylotype 16 was identical to that of both

Calanus pacificus as well as Calanus helgolandicus (Fig. 4,

Table 1) and was present at depth intervals throughout

the oxygenated part of the water column (Figs 2 and 3a).

Most likely, phylotype 16 represents the endemic Calanus

euxinus (Hulsemann) (Besiktepe et al., 1998) instead, for

which no 18S rDNA sequence is available. Both C. helgolan-

dicus and C. euxinus are morphologically similar and show

o 0.5% sequence differentiation in the faster evolving

mitochondrial cytochrome oxidase subunit I gene (mtCOI).

This recently raised the question of whether C. euxinus

is a different species (Unal et al., 2006). Calanus helgolandi-

cus is an epipelagic copepod and typically inhabits the saline

waters of world’s oceans (Fleminger & Hulsemann, 1977)

with salinity values of 32–39, whereas C. euxinus is a key

component of the Black Sea pelagic ecosystem (Besiktepe

et al., 1998) and has adapted to the lower salinity of the

Black Sea (�18 psu). Furthermore, the predominance of

phylotype 16 in both 18S rDNA/rRNA libraries within the

deeper waters, notably the suboxic zone, corroborates with

the lifestyle of C. euxinus in the Black Sea. Previous studies

have shown that C. euxinus descends during the day to the

cold suboxic zone thereby significantly decreasing its total

metabolism as well as enabling it to utilize the energy of

consumed food for growth and lipid accumulation (Sve-

tlichny et al., 2000; Svetlichny et al., 2006).

The copepod represented by phylotype 17 was metaboli-

cally active at most depths between 40 and 80 m, whereas the

closely related phylotype 18 was identified only from the

narrow interval within the oxygenated upper nitrite max-

imum between 105 and 110 m (Figs 2, 3a, and 4, Table 1).

The closest relative of both phylotypes was an unassigned

Fig. 4b. Continued.



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Table 1. General information about the eukaryotic phylotypes recovered from the Black Sea

Phyl. no. Tax. grp Closest (named) relative(s)

Sequence

similarity (%) General observations

12 Cop Pseudocalanus elongatus (AY 446903) 100 Present and active between 40 and 95 m

16 Cop Calanus helgolandicus (AY446908) 100 Present throughout most analyzed depths but mainly

active between 105 and 150 m

Calanus pacificus (L81939) 100

17 Cop Clone of Baffin Bay (EU182744) 100 Present and active at most depths between 40 and

80 m

18 Cop Clone of Baffin Bay (EU182744) 98.5 Active at 105 m, just present at 110 m

11 Tun Oikopleura dioica (AY116613) 99 Active in the chlorophyll maximum and in the upper

nitrite maximum (between 110 and 115 m)

1 Hap Cruciplacolithus neohelis (AB058348) 96 Active at most analyzed depths between 40 and

115 m

9 Din Gyrodinium rubrum (AB120003) and clones

from the Sargasso Sea deep chlorophyll

maximum (AY664955/72)

96–97 Present and active between 40 and 95 m, absent

from chlorophyll-depleted deeper waters

6 Din Clone SL163B11 (AY664961) Ross Sea 99 Found only at 60 m (active)

3 Din Gyrodinium fusiforme (AB120002) 98.5 Present between 90 and 100 m (80 m DNA sample

missing); active between 80 and 95 m

7 Din Gyrodinium fusiforme (AB120002) 99 Present at 100 m; activity not confirmed

14 Din Clone BB01_39 (AY885030) Mid-Atlantic

Estuary (Barnegat Bay, New Jersey)

98 Active at the oxic/suboxic zone transition (120 m);

present but inactive at 125 m

10 Din Clone SCM37C45 (AY664929) 99 Present at the oxic/suboxic zone transition (120 m);

activity not confirmed

Sargasso Sea deep chlorophyll maximum

13 Din Clones from sulfidic waters Mariager Fjord

(DQ103845/46/60/71)

98–99.8 Present in suboxic zone at 125 m; activity not

confirmed

8 Din Clone SCM28C60 (AY664961) Sargasso Sea

deep chlorophyll maximum

98.5 Present and active in suboxic zone (120–130 m) and

sulfidic chemocline at 150 m in the presence of 6 mM

of sulfide

4 Cil Clones from sulfidic waters of Framvaren Fjord

(DQ310304/14)

99.9 Active in fully oxygenated part of the photic zone at

40 m; not detected from sulfidic Black Sea waters

2 Cil Strombidium styliferum (DQ631805) 98 Active at the oxic/suboxic zone transition (120 m);

present but inactive at 125 m

15 Unc Clone FV23_2E8 (DQ310250) 83 Present at 150 and 160 m; active at 150 m in the

presence of 6 mM of sulfide

27 Cop Tortanus sp. New Caledonia (AY626995) 90 Present and active in up to 200-year-old (upper 4 cm)

sulfidic sediments

24 Rot Mytilina mucronata (DQ297708) 98 Low 18S rDNA content but activity confirmed in up to

300-year-old (upper 6 cm) sulfidic sediments

22 Hap Emiliania huxleyi (AB183618) 98 Low 18S rDNA content but activity confirmed in

upper 2 cm of sulfidic sediments

23 Din Scrippsiella sp. MBIC11164 (AB183674) 100 Present in upper 8 cm; active in upper 6 cm

21 Din Pentapharsodinium sp. CCMP771 (AF274270) 99.8 Present between 2 and 6 cm; activity not confirmed

25 Cil Clone M3_18E07 (DQ103803) 93 Present between 2 and 4 cm; activity not confirmed

Sulfidic waters Mariager Fjord

20 Cil Clone BS_DGGE_Euk-6 (DQ234286); Black Sea

sulfidic Unit I sediment

100 Present between 2 and 8 cm; activity not confirmed

26 Cil Clone M3_18E07 (DQ103803) 96 Present between 4 and 8 cm; activity not confirmed

Sulfidic waters Mariager Fjord

19 Cil Clone BS_DGGE_Euk-1 (DQ234281); Black Sea

sulfidic sapropel Unit II

97 Present between 6 and 8 cm; activity not confirmed

Phyl. no., phylotype number depicted from Fig. 4; Tax. grp, taxonomic group; Cop, copepods; Tun, tunicates; Hap, haptophytes; Din, dinoflagellates;

Cil, ciliates; Unc, unclassified; Rot, rotifers. Light- and dark-gray shaded part of the table represent phylotypes recovered from, respectively, the water

column and the sulfidic sediments of the Black Sea.



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copepod found at a depth of 180 m in the Arctic Baffin Bay

(Fig. 4, Table 1) with low in situ temperatures (1 1C) and low

chlorophyll concentrations (Hamilton et al., 2008). A paral-

lel identification of these unassigned copepod species based

on morphological characteristics was no longer possible

because the filters were completely used up by the nucleic

acids extraction.

Tunicata

Phylotype 11 represented Oikopleura dioica, the only re-

corded gelatinous tunicate (appendicularian) in the Black

Sea (Shiganova, 2005) (Fig. 4, Table 1). Its 18S rRNA (DGGE

band 11, Fig. 3a) was mainly found in the chlorophyll

maximum at 22 m but a faint DGGE band 11 was also

present in the lower part of the upper nitrite maximum

(between 110 and 115 m) (Figs 2 and 3a). The predomi-

nance of its 18S rDNA at 22 m is in accordance with the

knowledge that most individuals of this euryhaline and

eurythermic tunicate are present in the thermocline,

whereas fewer individuals appear to be present in the deeper

oxygenated waters of the Black Sea (Shiganova, 2005 and

references therein). The coinciding presence of Oikopleura

18S rRNA implies that viable Oikopleura individuals are

present in the chlorophyll maximum as well as in the deeper

oxygenated waters of the Black Sea (Figs 2 and 3a).

Oikopleura dioica populations grow exceptionally fast (Seo

et al., 2001) and this species ingests phytoplankton (5%) but

its main food is bacteria and detritus (95%) (Shiganova,

2005). The specialized filter apparatus of O. dioica retains

particles no larger than 15 mm and this feature allows feeding

in deep, detritus-based ecosystems. This utilization of small

size classes decreases competition with other zooplankton

species (Tiselius et al., 2003). The presence of viable

O. dioica in the nitrite maximum just above the suboxic

zone with low chlorophyll levels (Figs 2 and 3a) suggests that

O. dioica could be actively involved in filtering mainly

bacteria and/or detritus at this depth.

Haptophytes

Our molecular survey revealed a previously overlooked

haptophyte (i.e. phylotype 1), which is affiliated with

calcifying haptophytes and Cruciplacolithus neohelis as its

closest named relative (Fig. 4, Table 1). Whereas 18S rRNA

of this active haptophyte represented up to 30% of the total

RT-PCR-amplified 18S rRNA pool (Fig. 2), the correspond-

ing genomic 18S rDNA of this species was often below

detection on DGGE (Fig. 3a) and most likely also escaped

prior identification from an 18S rDNA library constructed

for additional locations in the Black Sea (Coolen et al.,

2006). This discrepancy can be caused by the fact that DGGE

is insensitive to less dominant species (e.g. Coolen et al.,

2007), or those with low genomic rRNA gene copy numbers.

The presence of active cells of this haptophyte down to

115 m could point to a capacity for heterotrophic growth,

given the extremely low light intensity and narrow light

spectrum supporting photosynthetic activity at this depth

(Overmann et al., 1992). Many haptophytes from early

diverging, noncalcifying lineages (Prymnesiales, Phaeocys-

tales) have retained heterotrophic behavior such as preda-

tion on bacteria and small algae (i.e. phagotrophy) (Kawachi

et al., 1991; Tillmann, 1998). Phagotrophy is restricted to

motile haptophyte cells with an emerging prey-catching

haptonema (Kawachi et al., 1991; Jones et al., 1994;

Tillmann, 1998). Also the motile haploid flagellate phase of

certain calcifying haptophytes were found to be phago-

trophic (Houdan et al., 2006). A phagotrophic life style for

the haptophyte related to calcifying species is thus plausible

and could be verified as soon as a culture from the Black

Sea’s deep suboxic zone would become available.

The widespread marine bloom-forming calcareous hap-

tophyte Emiliania huxleyi, which is also common in the

Black Sea (Eker-Develi & Kideys, 2003), was not identified in

our molecular surveys. A test with parallel tag-encoded FLX

amplicon pyrosequencing (e.g. Sogin et al., 2006) of a

130-bp-long fragment of the V9 region revealed that 18S

rDNA of haptophytes of the order Isochrysidales to which E.

huxleyi belongs, represented only 2% of the total eukaryotic

18S rDNA in the chlorophyll maximum at 22 m (M.J.L.

Coolen & C. Davis, unpublished data). As outlined earlier,

DGGE is biased to identify only phylotypes that represent

4�2% of the total pool of PCR-amplified DNA (e.g. Coolen

et al., 2007), and would explain why E. huxleyi was missed in

our general eukaryotic 18S rDNA PCR/DGGE approach.

Dinoflagellates

Our sampling set revealed eight unique dinoflagellate phy-

lotypes, which were found to exhibit high vertical stratifica-

tion, and their presence often coincided with the presence of

specific nitrogen species or suboxic/anoxic conditions. For

example, the dinoflagellates represented by phylotypes 9 and

3 were present and active at specific depths between 40 and

95 m (Figs 2 and 3a), and were closely affiliated with

uncultivated dinoflagellates from the deep chlorophyll max-

imum in the Sargasso Sea and/or phagotrophic Gyrodinium

spp. (Fig. 4, Table 1).

In addition, nucleic acids of some unassigned dinoflagel-

lates were found only in the suboxic or anoxic waters of the

Black Sea. The dinoflagellates represented by phylotypes 10

and 13 (Fig. 4, Table 1) were rare in the water column and

only their 18S rDNA was identified from the upper part of

the suboxic zone (Figs 2 and 3a). Therefore, their activity or

their indigeneity to suboxic waters could not be confirmed.



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On the other hand, phylotype 14 (Fig. 4, Table 1) was clearly

active at the oxic/suboxic transition at 120 in the presence of

3 mM of oxygen (Figs 2 and 3a). The DNA of this dino-

flagellate was most abundant at 125 m, whereas its RNA was

below the detection limit at this depth, underlying the value

of comparative analysis of gene transcripts for verification of

metabolic activity of microbiota within environmental gra-

dients. Phylotype 8, related to a clone from the Sargasso

Sea deep chlorophyll maximum, represented the only dino-

flagellate to be active in the deeper part of the suboxic zone

with undetectable levels of oxygen, as well as at 150 m in the

presence of 6mM of hydrogen sulfide (Fig. 2).

The sulfidic chemocline furthermore contained tran-

scripts of the 18S rRNA gene of an unclassified eukaryote

(i.e. phylotype 15) with up to 96% sequence similarity to

18S rDNA sequences recovered from sulfidic waters of

Framvaren Fjord (Fig. 4, Table 1) (Behnke et al., 2006). This

phylotype was not detected from the oxygenated waters

above, but a more sensitive PCR approach using species-

specific primers would be required to verify whether the

presence of this eukaryote was restricted to the slightly

sulfidic part of the stratified water column of the Black Sea.

Ciliates

Information about the species composition of ciliates in the

Black Sea is sparse and mostly relates to coastal areas

(Kovaleva & Golemanski, 1979; Bouvier et al., 1998). Our

survey revealed only two unique phylotypes of ciliates

throughout the upper 160 m of the water column. Phylotype

4 (Fig. 4, Table 1) is closely related to ciliates of the class

Oligohymenophorea and was found in the RNA-based (me-

tabolically active) library from fully oxygenated waters at a

depth of 40 m (Figs 2 and 3a). Environmental sequences

from DNA-based libraries of highly sulfidic Framvaren

Fjord waters (Behnke et al., 2006) were the closest relatives

of phylotype 4 (Fig. 4).

A ciliate with Strombidium styliferum as its closest relative

(phylotype 2) (Fig. 4, Table 1) was only found to be

metabolically active at the oxic/suboxic zone transition at

120 m and just present at 125 m. However, more precise

genotyping would be required to identify this ciliate because

a total of 63 morphotypes have been described from this

genus (Agatha, 2004) and the estimates of genetic diversity is

much greater (Song & Packroff, 1997).

Eukaryotes in the Black Sea’s sulfidic sediments

Nine out of 17 successfully sequenced DGGE bands recov-

ered from the upper 8 cm of sulfidic sediments that underlie

830 m of sulfidic waters represented unique phylotypes and

grouped with copepods (phylotype 27), rotifers (phylotype

24), haptophytes (phylotype 22), dinoflagellates (phylotypes

21 and 23), and ciliates (four phylotypes; 19, 20, 25, 26)

(Fig. 4, Table 1). Figure 3b displays the position of the

sequenced DGGE bands corresponding to these phylotypes.

The phylogenetic affiliation with other eukaryotes, their

relative abundance, and whether they represent species

indigenous to the sediments or allochthonous species de-

rived from the water column above, will be discussed next.

Copepods

The upper 4 cm of sediment contained both DNA and RNA

from a copepod distantly related to Tortanus sp. New

Caledonia (phylotype 27; Fig. 4, Table 1). In the surface

Fig. 5. The relative distributions (%) of 18S

rDNA (closed circles) vs. 18S rRNA (open circles)

of ‘total’ vs. viable eukaryotes found in the

upper 8 cm of sulfidic Black Sea sediments.



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sediment (0–2 cm), 18S rRNA of this copepod represented

40% of the total RT-PCR-amplified 18S rRNA pool (Fig. 5).

Most likely, the 18S rRNA of this Tortanus-related copepod

was extracted from resting eggs derived from the overlying

oxygenated water column. Assuming an absence of a post-

depositional redistribution of the exquisitely laminated

sulfidic sediments (Fig. 1b) and an average sedimentation

rate of 0.2 mm year�1 (M.J.L. Coolen, unpublished data), the

sediment interval at 4-cm depth with detectable Tortanus

18S rRNA gene transcripts would correspond to a deposi-

tional age of 200 years before present. This finding indicates

that copepod eggs (depending on the species) could remain

viable with detectable levels of 18S rRNA gene transcripts in

highly sulfidic environments for up to 200 years after

sedimentation. This is significantly longer than the previous

reported recovery of viable copepod eggs from up to

40-year-old sulfidic sediments from the stratified Petta-

quamscutt River Basin (RI) in the presence of 2 mM of

hydrogen sulfide (Marcus et al., 1994), which is approxi-

mately seven times the concentration of sulfide in the Black

Sea surface sediments (Zopfi et al., 2004). Because the

primary targets of sulfide are aerobic respiration enzymes,

for example cytochrome c oxidase, it was suggested that

copepod eggs may gain protection from sulfide by under-

going anaerobic respiration (Marcus et al., 1997).

Rotifers

The rotifer Mytilina mucronata was the closest relative of

phylotype 24 (Fig. 4). Its 18S rRNA, probably extracted from

viable resting eggs, comprised up to 20% of the total 18S

rRNA pool within up to 300-year-old (i.e. the upper 6 cm)

sulfidic sediments (Figs 3b and 5). 18S rDNA of the rotifer

was also identified but resulted only in a faint DGGE band

(band 24; Fig. 3b) at 2–4 cm, which was also the sample with

the most intense corresponding band with RT-PCR ampli-

fied 18S rRNA. Our data suggest that rotifer eggs in the

Black Sea sediments remained longer viable than copepod

eggs because the copepod 18S rRNA rapidly declined and

reached undetectable levels o 4 cm (i.e. �200-year-old

sediments; Fig. 5). This is in agreement with the results from

Marcus et al. (1994) who found that a higher percentage of

eggs from a rotifer of the genus Brachionus could be hatched

from up to 40-year-old sulfidic Pettaquamscutt River Basin

sediments as compared with copepod eggs.

Haptophytes

Interestingly, a unique haptophyte (phylotype 22; Figs 3b

and 4, Table 1) with E. huxleyi as its closest relative was

found to be active in the top 2 cm of sediment, whereas its

DNA was too faint to be detected by DGGE. This haptophyte

was not detected from the water column at the time of

sampling. These results imply that this haptophyte has

developed a strategy to remain metabolically active for

substantial periods of time (at least weeks and up to

decades) in the presence of �175 mM of sulfide. Previously,

it was shown that haptophytes were among various protists

that were found to proliferate from incubated Swedish

coastal sediments (Persson, 2002) but it is unclear to what

extent those protists were exposed to hydrogen sulfide.

Dinoflagellates

Nucleic acids of two dinoflagellate phylotypes were recov-

ered from the sediments. Phylotype 23 is related to dino-

flagellates of the cyst-forming genus Scrippsiella with an

identical sequence to that of Scrippsiella sp. MBIC11164

(Fig. 4, Table 1). Phylotype 23 comprised up to 40% of the

total 18S rRNA pool in the upper 6 cm of sulfidic sediment

(Fig. 5), which suggests that cysts of this species remain

viable for up to 300 years.

On the other hand, phylotype 21 with 99.8% sequence

similarity to species of the genus Pentapharsodinium (Fig. 4,

Table 1) was only recovered from the sedimentary 18S rDNA

pool. Its DNA was most likely protected inside organic-

walled cysts of Pentapharsodinium, which have been

described from recent Black Sea sediments (Marret et al.,

2004). However, the absence of Pentapharsodinium RNA

suggests that cysts of Pentapharsodinium were no longer

viable in the sulfidic sediments.

Ciliates

Phylotypes 19, 20, 25, and 26 were recovered from up to

8 cm of sulfidic Black Sea sediments (Figs 3b and 5) and

were closely related to 18S rRNA clones from unnamed

ciliates previously retrieved from the sulfidic waters (i.e.

�50 mM of sulfide) of the Danish Mariager Fjord (Fig. 4,

Table 1) (Zuendorf et al., 2006). The absence of 18S rRNA

from the four ciliates implied that they were not viable in the

subsurface Black Sea sediments, which exhibit a sulfide

concentration three times higher than in the Mariager Fjord

waters. Whether the sedimentary ciliate DNA was derived

from cells stemming from the less sulfidic chemocline could

not be confirmed because the same phylotypes were not

detected in the two analyzed sulfidic waters at 150 and

160 m.

Summary and conclusions

We have shown that analysis of gene transcripts has enabled

the identification of species that were active but escaped

identification in a parallel 18S rDNA survey. In addition, the

above examples also indicate that gene transcripts do not

always provide unequivocal proof that metabolically active



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microorganisms are indigenous to a certain position in an

environmental gradient, because certain species stemming

from the oxygenated part of the photic zone can remain

viable in the presence of high sulfide concentrations. This is

especially true when environmental sequences represent

novel taxonomic clades with unknown phylogenetic affilia-

tion. The 18S rRNA gene survey from surface sediments

yielded taxa of pelagic origin, but none of these were

identified in the water column at the time of sampling. Vice

versa, none of the species, which were thriving in the water

column at the time of sampling, could be detected in the

fossil DNA fraction. There are various possible explanations

for this discrepancy: (1) the filtered POM only represents a

snapshot of the total annual plankton community, (2) not

all cellular material is equally well transported to the

sediment, or (3) there are species- or cell-specific variations

in the level of (post)depositional degradation of intracellular

DNA. Although the DGGE method represents a relatively

fast approach in screening a large number of samples along

the environmental gradient for major shifts in the eukaryote

community structure, it is insensitive to less dominant

species, or those with low genomic rDNA copy numbers.

Future research will require application of other molecular

biological approaches, notably tag-encoded FLX amplicon

pyrosequencing, in order to determine the diversity and

relative abundance of total vs. active eukaryotes.

Acknowledgements

We would especially like to thank Ognyana Hristova and

Tatyana Nikolova, IOBAS, for the analysis of the water

column chemistry and Dr Cornelia Wuchter, Dr Angela

Dickens, Alan Gagnon, Daniel Montlucon, Chris Ward

(WHOI), and the R/V Akademik staff, in particular Delcho

Solakov, for their extensive organizational and participatory

help with the cruise. We thank Dr Timothy Eglinton

(WHOI) and two anonymous reviewers for suggestions to

improve the manuscript. We are grateful for the financial

support from the US National Science Foundation grant

OCE 0602423, as well as funding from WHOI’s Access to the

Sea program, and a grant from the Andrew W. Mellon

Foundation Endowed Fund for Innovative Research.

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