Polar Biol (2009) 32:105–112

DOI 10.1007/s00300-008-0510-4

ORIGINAL PAPER

Response of benthic microbial communities to chitin enrichment: an in situ study in the deep Arctic Ocean

Corinna Kanzog · Alban Ramette · Nadia V. Quéric · Michael Klages

Received: 27 January 2008 / Revised: 18 August 2008 / Accepted: 19 August 2008 / Published online: 3 September 2008© The Author(s) 2008. This article is published with open access at Springerlink.com

Abstract In situ enrichment experiments were carried outin the Arctic deep sea (Fram Strait region) to observe theresponse of benthic microbial communities to chitin supply.Chambers of a benthic lander were Wlled in July 2004 withdeep-sea sediments enriched with 1.3–7.0 g m¡2 of chitinand the eVects of chitin enrichment were assessed on themicrobial hydrolytic activity potential, cell number andcommunity structure after periods of 1 week and 1 year ofin situ deployment. The input of chitin had no eVect onmicrobial abundance and chitobiase activity after 7 days ofincubation, whereas community structure in enriched sedi-ments, determined by terminal restriction fragment lengthpolymorphism analysis of 16S rRNA genes, was diVerentfrom the controls. After 1 year, microbial numbers andactivity signiWcantly increased in sediments enriched withhigh chitin concentrations and bacterial community struc-ture was diVerent from that of the other treatments. Thepresent study suggests that microbial community structurein Arctic deep-sea sediments can react quickly to suddenlarge chitin inputs into the sediments and that this appearsto precondition subsequent enhanced growth and enzymaticactivity changes.

Keywords Chitin · Deep-sea sediments · Microbial communities

Introduction

Sinking particulate organic matter (POM) that reaches thedeep seaXoor mainly consists of less degradable materialincluding cellulose, chitin and various proteinaceous struc-tural compounds (Wakeham and Lee 1993). As a majorcomponent of cuticles and exoskeletons of worms, mol-luscs and arthropods, chitin is probably the most importantbiopolymer in marine environments (Gooday 1990).Although other organisms may be able to decompose chi-tin, the breakdown of such degradation-resistant organicmatter deposited in deep-sea sediment surfaces is mainlyaccomplished by bacteria which are able to extracellularlycleave structural polymeric compounds (Deming andBaross 1993).

A variety of time course experiments have been per-formed to test the functional responses of benthic deep-seabacteria to chitin input [e.g. North Atlantic (Deming 1985);Arctic continental slope (Boetius and Lochte 1996); Ara-bian Sea (Christiansen and Boetius 2000)]. These studiesindicated that chitin supplies induced the synthesis of spe-ciWc enzymes and that indigenous deep-sea bacteria wereable to decompose even high amounts. Previous experi-ments were conducted under laboratory conditions and didnot provide detailed information on the variation in micro-bial community over periods longer than weeks. In situexperiments focussing on microbial response to organicenrichments in the deep-sea are scarce (e.g. Moodley et al.2002; Bühring et al. 2006), especially in Polar regions.

Studies have reported mass sedimentation events ofcrustaceans in diVerent ocean regions (Sokolova 1994;

Electronic supplementary material The online version of this article (doi:10.1007/s00300-008-0510-4) contains supplementary material, which is available to authorized users.

C. Kanzog (&) · N. V. Quéric · M. KlagesAlfred Wegener Institute for Polar and Marine Research, Am Handelshafen 12, 27570 Bremerhaven, Germanye-mail: Corinna.Kanzog@awi.de

A. RametteMicrobial Habitat Group, Max Planck Institute for Marine Microbiology, Celsiusstr. 1, 28359 Bremen, Germany

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106 Polar Biol (2009) 32:105–112

Christiansen and Boetius 2000), which represent a highinput of chitinous material. The fact that high concentra-tions of the Arctic copepod Calanus hyperboreus wererepeatedly observed over the ocean Xoor of the GreenlandSea (Hirche et al. 2006) suggests that high amounts ofchitinous material would be naturally available when thislarge aggregation of organisms dies or moults. In addition,shifts in the distribution and abundance of key copepodspecies from the Arctic Ocean, possible caused by environ-mental changes associated with climate changes (Wdsiaw-ski and Legeryjska 1998; Hirche and Kosobokova 2007),may also contribute to enhanced chitin input to the deepseaXoor. Investigations are necessary to better understandthe eVects of chitin supply on deep-sea microbial assem-blages as main agents in the mineralisation of chitinousmaterial. Here, for the Wrst time, short-term (7 days) andlong-term (1 year) incubation experiments were carried outin situ to assess the eVects of varying concentrations ofchitin on microbial communities in Arctic deep-sea sedi-ments. We tested the hypothesis that a higher amount ofchitin introduced into the sediments leads to increasedmicrobial growth, activity and possibly to signiWcant changesin bacterial community structure.

Materials and methods

Experimental design

A free-falling benthic lander, containing deep-sea sedi-ments enriched with diVerent chitin concentrations, wasdeployed for 7 days and 1 year at 2,500 m water depth inthe long-term observatory HAUSGARTEN (79°5�N/4°2�E), in the eastern Fram Strait, west oV Svalbard (Soltw-edel et al. 2005). The lander was designed by the InstitutFrançais de Recherche pour l’Exploitation de la Mer (IFR-EMER) and consisted of four identical trays, each of whichwas equipped with four separate chambers (0.02 m2). Onlyone tray was used for our enrichment experiments.

Natural deep-sea sediments from the central HAUS-GARTEN station were collected with a multiple corer(MUC; Barnett et al. 1984) at 2,505 m water depth on 9July 2004 during the cruise ARK XX/1 with RV ‘Polarstern’.Sediments were composed of a mixture of 37% sand (particlesize 63–2,000 �m), 43% silt (4–63 �m) and 20% clay(<4 �m). Surface sediments (0–3 cm) of several sedimentcores were pooled and transferred into lander chambers.Following sediment freezing at ¡30°C for 48 h and subse-quent thawing, a ‘low’, ‘medium’ and ‘high’ chitin con-centration corresponding to 1.3, 3.0 and 7.0 gm¡2 chitin(puriWed chitin Xakes from crab shells, Sigma–Aldrich,Germany), respectively, were mixed with the upper layersof the sediments placed in the chambers. The additions

were equivalent to a substrate input of 0.5–3.0 g C m¡2,which are well above the background value of chitin indeep-sea sediments of 0.1 mg m¡2 (Poulicek and Jeuniaux1989). A control treatment consisted of one chamber Wlledwith deep-sea sediments without any addition of organicsubstrate. Before deployment, sediment samples were takenfor initial analysis of microbial parameters (0 day).

Sub-sampling and sample processing

During RV ‘Polarstern’ cruise ARK XX/1 in July 2004, thebenthic lander was deployed for 7 days. In order to preventdisturbance of the sediments inside the chambers, the trayswere closed during transport in the water column andopened again after landing at the seaXoor by means of aremotely controlled mechanism. After recovery and subse-quently sampling, trays were equipped with new chambers.These chambers were Wlled with sediments obtained fromMUC sampling for the short-term experiment which werestored frozen until the deployment. Amendment of sedi-ments with chitin was carried out in the same way asdescribed above. Trays for the long-term experiment weredeployed at the same station for 1 year. Sampling of the 1-year experiment took place during RV ‘Polarstern’ expedi-tion ARK XXI/1b in July 2005.

After each recovery of the trays, eight samples from theWrst centimetre of sediments in each chamber were takenusing plastic syringes with cut-oV ends (2.5 cm in diame-ter). Sub-sampling was carried out in a room at 4°C. Sedi-ments from each chamber were pooled and homogenisedbefore determining the abundance, enzymatic activity andchanges in bacterial community structure.

Microbial biomass

For total microbial cell numbers and biomass determina-tion, 2 ml of the sediment sub-sample (2.3 cm3) were takenby means of plastic syringes with cut-oV ends (1.2 cm indiameter) and stored at 4°C in 9 ml 2% formaldehyde solu-tion. After staining with acridine orange, total cell countswere determined by epiXuorescence microscopy accordingto Meyer-Reil (1983). Volumetric measurements of themicrobial cells were conducted with the New Porton grid,measuring randomly 50 cells per Wlter (Grossmann andReichardt 1991). Microbial biomass was estimated using aconversion factor of 3 £ 10¡13 g C �m¡3 (Børsheim et al.1990). For each sample 40 counted grids from 2 replicateWlters were analysed.

Enzyme activity

To assess the chitinolytic activity of microbial organisms,we used the assay for chitobiase activity, providing a

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simple and sensitive determination of extracellular enzy-matic hydrolysis rates after relatively short incubation timeand adaptable at low temperature (Hoppe 1983; Poremba1995). The extracellular enzymatic activity of chitobiasewas measured Xuorometrically using the methylumbellifer-one (MUF) labelled substrate N-acetyl-�-glucosaminide(Sigma–Aldrich; Boetius and Lochte 1994). Previous trialsto determine the substrate saturation level gave a Wnalconcentration of 100 �mol MUF-substrate. Relative unitsof Xuorescence were calibrated with seven MUF standardconcentrations between 10 and 300 nmol. The enzymeactivity was calculated per volume of sediment and time(�mol cm¡3 h¡1) by linear regression. The regressioncoeYcient was always better than 0.95. For each samplethree measurements per incubation were performed.

Terminal-restriction fragment length polymorphism (T-RFLP) analysis

Sediment samples for microbial community analysis werestored at ¡30°C until analysis. Total DNA was extractedand puriWed using a FastDNA®SPIN Kit for soil (Qbiogene,Heidelberg, Germany) following manufacturer’s instruc-tions. DNA concentrations ranged from 50 to 80 ng �l¡1.

Analysis of community structure focussed exclusivelyon bacterial groups since they are one of the most importantagents in the mineralisation of chitin (Cottrell and Kirch-man 2000). The primers used for ampliWcation of eubacte-rial 16S rDNA were 27F and 1492R (both synthesised byInteractiva; Ulm, Germany) (Rappé and Giovannoni 2003).The bacterial speciWc forward primer 27F was end-labelledwith phosphoramidite Xuorochrome 5-carboxyXuorescein(5�6-FAM). The PCR mixture contained 2 �l of templateDNA, 0.25 �mol (each) primer, 1.5 mmol KCl, 10 mmolTris–HCl pH 9, 1.5 mmol MgCl2, 250 �mol dNTP, and1.25 U Taq-Polymerase (Amersham Pharmacia Biotech,Freiburg, Germany) in a Wnal volume of 50 �l. DNA ampli-Wcation was performed by using the following cycling con-ditions: a 3-min hot start at 95°C, followed by 29 cyclesconsisting of denaturation (1 min at 95°C), annealing(1 min at 55°C), and extension (1 min at 72°C). A Wnalextension at 72°C was then done for 7 min. Fluorescentlylabelled PCR products were run onto 1% agarose gel andpuriWed using QIAquick Gel Extraction kit (Qiagen, Hil-den, Germany) following the manufacturer’s instructions.Approximately 300 ng of the PCR product was digestedwith 10 U of endonucleases HhaI and MspI (AmershamPharmacia Biotech). The digestions were performed sepa-rately in a total volume of 50 �l at 37°C for 5 h. Restrictiondigestions were desalted (isopropanol 100%) and driedprior to storage at ¡20°C. Fragment separation wasperformed by Gene Analysis Service GmbH (Berlin,Germany) using an ABI Prism 310 capillary analyser (PE

Applied Biosystems, CA, USA). Peak size determination ofend-labelled fragments, considered here as diVerent Opera-tional Taxonomic Units (OTUs), was done by comparisonwith an internal size standard (GS2500 TAMRA, PEApplied Biosystems). Peaks between 50 and 1,000 basepairs long were analysed using Genescan analytic software2.02 (ABI). An additional check for artefacts was manuallyperformed and peaks whose areas were smaller than 1% ofthe total peak area were excluded from data analysis.Due to the limited amount of available sediment samples,T-RFLP analyses were done once for each treatment.

Data analysis

Microbial cell number and chitobiase activity were log10-transformed prior to performing statistical analyses in orderto normalise their distribution and homogenise treatmentsvariances (Crochan test of homogeneity, Sokal and Rohlf1995). Analysis of variance (ANOVA) was applied to eachdata set to test the eVects of chitin input and incubationtime, and of their interactions. Post-hoc comparisonsbetween treatment means were done using Tukey’s hon-estly signiWcant diVerence (HSD) test at P < 0.05.

T-RFLP Wngerprints obtained from the two diVerentrestriction digests were combined to generate a binarymatrix according to the presence or absence of bacterialOTUs. The unweighted pair group with arithmetic mean(UPGMA) method was applied with the Dice similarityindex (Dice 1945). Nodal support in the resulting dendro-gram was determined by performing 100 bootstrap repli-cates (Hammer et al. 2001). In addition, non-metricmultidimensional scaling was used to obtain an ordinationof the samples based on a matrix of Dice dissimilarities ofT-RFLP proWles (Ramette 2007). A canonical redundancyanalysis (RDA) was used to determine how well diVerentfactors could explain the variation in community structure,and in order to assess the respective importance of thediVerent factors and of their covariation on shaping com-munity structure, a variation partitioning procedure(Ramette and Tiedje 2007) was subsequently applied to thecommunity data. Statistical analyses were performed withthe statistical software JMP IN version 5.1 (SAS Institute,Cary, NC, USA) and with the R package (R version 2.4.0;The R foundation for Statistical Computing).

Results and discussion

EVects of chitin addition on microbial abundance and activity

Changes in microbial biomass were mainly explained byvariation in microbial cell numbers and less by changes in

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cell volume (data not shown), thus we mainly focussed ouranalysis on variations in microbial cell numbers. Each fac-tor taken alone (chitin input and incubation time) as well asthe interaction between factors had highly signiWcanteVects on microbial abundance and chitobiase activity(Table 1). In sediments incubated for 1 week, regardless ofthe chitin amounts added, no signiWcant changes in micro-bial abundance were observed and initial chitobiase activityalso remained constantly low in all treatments (Fig. 1a, b).These results indicate that within 1 week, a pulse input ofchitin to the seaXoor has no eVect on microbial cell num-bers and chitobiase activity. These observations are corrob-orated by other enrichment studies with deep-sea sedimentswhich have shown time lags for the microbial response tonutrient pulses of up to 2 weeks (e.g. Witte et al. 2003).More speciWcally, Boetius and Lochte (1996) detected an

increase in chitobiase activity 10 days after chitin addition.Investigations of Deming (1985) with deep-sea sedimentsamended with chitin showed that bacterial doubling timewere in the order of weeks or months. Thus, bacterialgrowth is suYciently slow to preclude detection of signiW-cant increases in cell number after 7 days of incubation.

After 1-year incubation substantial increases of micro-bial cell number associated with increasing chitin concen-trations were observed (Fig. 1a). Total cell counts in thehighest (7.0 g m¡2) and medium (3.0 g m¡2) chitin amend-ments were, respectively, about nine and Wve times higherthan prior to incubation, whereas in low chitin amendment(1.3 g m¡2) only a marginal increase was found. Similartrends were recorded for microbial chitobiase activity(Fig. 1b). The activity levels in control samples andsediments enriched with low chitin amounts were not

Table 1 Analysis of variance of microbial abundance and activity as a function of chitin input and incubation time

a The total cell number was log10 transformed prior to performing statistical analyses to normalise its distributionb Degrees of freedomc The F ratios were all highly signiWcant with P < 0.001 (***)

Source Sum of squares df b Mean square F ratioc

Microbial cell numbera

Chitin input 0.51 3 0.17 153.404***

Incubation time 1.313 2 0.656 592.487***

Chitin input £ Incubation time 0.409 6 0.068 61.537***

Chitobiase activity

Chitin input 21.252 3 7.084 201.375***

Incubation time 21.617 2 10.808 307.248***

Chitin input £ Incubation time 42.598 6 7.1 201.822***

Fig. 1 Microbial cell number (a) and chitobiase activity (b) in control and chitin enriched sediments after 7-day and 1-year incubations. Bars represent mean values § SD calculated from replicate samples. DiVerent letters (A, B, C) above the bars indicate signiWcant diVerences between mean treatments as determined by post hoc Tukey’s HSD tests at P < 0.05

2

4

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7 days 1 year

Control 3.0 Control 1.3 3.0 7.00

2

4

6

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b)

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robi

al c

ell n

umbe

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ase

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ity

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ol c

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Chitin concentration [g m ]-2

C C C C C C C C C C

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A

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Polar Biol (2009) 32:105–112 109

signiWcantly diVerent from the short-term incubation,whereas fourfold and 30-fold activity increases were foundin medium and high treatments, respectively (Fig. 1b).

To estimate changes in activity potential that were notrelated to variation in microbial biomass, chitobiase activitywas normalised by the total cell number. The highest chito-biase activity per cell was measured in sediments withhighest chitin input after 1 year (7.5 £ 10¡18 molcell¡1 h¡1), which was approximately three times higherthan in the other incubations at the same time. Previousinvestigations on microbial activity in continental slopesediments have shown positive correlations between chito-biase and organic matter supply to the benthos (Vetter andDeming 1994; Boetius and Lochte 1996). Our resultsshowed that chitin amendment of 7.0 g m¡2 can induceincreasing speciWc chitobiase activity similar to in situ mea-surements weeks after sedimentation of a large numbers ofcrab carcasses by Christiansen and Boetius (2000). In gen-eral, values for chitin production in oceanic ecosystems arealmost inexistent (Cauchie 2002), particularly for Polarregions. According to estimates by Cauchie (2002), totalchitin production by crustaceans in marine environmentswould amount to about 109 t chitin year¡1. In particular, theauthor used a mean value of 40 mg chitin m¡2 year¡1 as anestimation of the combined chitin production by benthicdeep-sea macrofauna such as copepods, decapods and iso-pod. Therefore, a chitin amendment of 7.0 gm¡2 is largerthan the chitin input that may result from the sedimentationof one crab to a square metre of sediment (about 1.0 g;Christiansen and Boetius 2000), and much more than thebackground chitin value in deep-sea sediments of0.1 mgm¡2 (Poulicek and Jeuniaux 1989). Hence, we sug-gest that, if a large amount of chitinous material naturallybecomes available in deep-sea areas, benthic microbialcommunities may react by a signiWcant increase in their cellnumbers and chitinolytic enzyme production. Such a highchitin input would be produced by mass sedimentation ofArctic copepods for example (Hirche et al. 2006), whichhave a mean chitin content of about 7% of the whole bodydry weight (Cartes and Maynou 1998).

To assess the substrate conversion by microbial organ-isms, we divided the biomass yield after 1 year by theorganic C amount introduced by the chitin concentrations(Boetius and Lochte 1996). Results indicated that of thelow, medium and high chitin-C amounts added to theenrichments, 6, 26 and 31% were converted into microbialbiomass after 1 year, respectively. Despite the high chitinconcentration, its degradation rate has been rarely estimatedin marine environment directly (Herwig et al. 1988; Pou-licek and Jeuniaux 1991; Kirchner 1995). It is commonlyassumed that chitin must be degraded at the same rate as itis produced because chitin does not accumulate to a largeextent in most marine sediments (Poulicek and Jeuniaux

1991). Indeed, Poulicek and Jeuniaux (1991) demonstratedthat about 90% of chitin produced in the water column inmarine environments was digested within 150 h. In sedi-ments, chitin digestion took longer than in the open water,taking at least 1 year. We assume that in contrast to themedium and high chitin treatments the lowest chitinamount of 1.3 gm¡2 was already consumed after 1 year.Furthermore, in sediments with medium and high chitinamounts the selection of chitinoclastic communities mighthave increased over time as a result of higher carbonamounts entering into the system, possibly explaining vari-ations in substrate conversion. In addition, potential lossesdue to meiofaunal and microeukaryotic grazers could notbe completely avoided since the trays were covered by5 mm mesh size grids. It is also likely that additionalorganic materials may have been introduced from the watercolumn to the sediment trays during 1 year. This eVect,however, contributed only marginally to changes in micro-bial biomass, as suggested by the only small variations inmicrobial abundance detected in control sediments(Fig. 1a).

Shifts in bacterial community structure as a response to chitin inputs

Several studies have already shown that microbial commu-nities are able to respond within short time to variations inthe quality and quantity of organic matter supply bychanges in their composition (Riemann et al. 2000; Eileret al. 2003). These short-term shifts were generally accom-panied by a functional response of bacterial communities.Our results from T-RFLP analysis (see Electronic Supple-mentary Material) also showed changes in communitystructure during incubation for 7 days. Interestingly here,rapid community changes apparently occurred without con-comitant changes in microbial growth and activity (Fig. 1).Cluster analysis of T-RFLP data generally indicated theexistence of three well-deWned clusters (Fig. 2), which wereconsistent with diVerences in incubation times and sedi-ment treatment. Cluster composed of control samples atdiVerent incubation times indicated that communities insediments without additional chitin remained similar intheir composition during the duration of the whole experi-ment. Unfortunately, we do not have any information aboutbacterial communities in natural deep-sea sediments thatwere not treated by freezing which may lead to communityalteration (Stocum and Plante 2006). Our results indicated,however, that there were only minor structural variationsbetween communities in initial control samples and thoseincubated for 1 year in which the community structure waspresumably representative for those naturally present indeep-sea sediments at the experimental site due to gradualcolonisation by indigenous populations. Additionally,

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initial community data from sediments amended with thelowest chitin concentration revealed only a marginal eVectof microbes possibly introduced by the non-sterilised chitinon community structure identiWed in deep-sea sediments.T-RFLP proWles from these initial chitin-amended sedi-ments were remarkably similar to those from control sedi-ments without chitin (Fig. 2). In general, bacterialcommunity structure in chitin-amended sediments wasfound to be gradually more dissimilar over the course of theexperiment, suggesting that chitin had a time-dependenteVect on the resident bacterial communities. Indeed,sample points were found to be more spread in the two-

dimensional nMDS plot with increasing incubation time,indicating more variability in community structure (Fig. 3).Despite the diYculty of obtaining enough materials to per-form T-RFLP analyses when setting up the experiment at2,500 m water depth, the eVects of both incubation time andchitin input on community structure were conWrmed byredundancy analysis (P = 0.031 based on 1,000 MonteCarlo permutations). Further variation partitioning of thebiological variation into the respective eVects of the twofactors indicated that chitin input was the most importantfactor (12% of biological variation was explained by thefactor alone when taking duration of the experiment as acovariable), closely followed by the duration of incubation(9%; data not shown). The total amount of unexplained bio-logical variation was 79%, which yet suggests thatunmeasured factors would also need to be added in thefuture to better understand community dynamics in deep-seasediments.

Few studies have demonstrated that metabolic changefollowing alterations of environmental conditions (e.g. sub-strate amendments) may occur without simultaneouschanges in bacterial community structure (Riemann andMiddelboe 2002; Findlay and Sinsabaugh 2003). In con-trast, mesocosm experiments conducted with manipulateddissolved organic matter (DOM) concentrations in fresh-water environments (Judd et al. 2006) demonstrated thateVects of changing DOM supply on bacterial productivitywere mediated by initial shifts in community structure.Similarly, our results suggest that for deep-sea sediments, afunctional response of microbial communities following alarge chitin input may be triggered by an initial change incommunity structure before eYcient utilisation of chitincompounds can be made. It may be also envisaged thatthe selection of speciWc populations (e.g. eYcient chitindegraders) may accelerate over time as a function of theamount of carbon entering into the system.

Our results revealed chitin-dependent microbialresponses and that POM supply can shape benthic microbial

Fig. 2 Cluster analysis of T-RFLP data. The UPGMA dendrogramwas constructed based on OTU presence/absence data obtained byT-RFLP analysis of the diVerent sediment samples. Nodal support wasassessed by 100 bootstrap replicates

100

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0.3 0.4 0.5 0.6 0.7 0.8 0.9 1

Similarity

71

80

49

98

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65

1.3 g, 1 year

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7.0 g, 1 year

Control, 0 days

Control, 7 days

7.0 g, 7 days

3.0 g, 7

1.3 g, 7 days

Control, 1 year

1.3 g, 0 days

Fig. 3 Non-metric multidimen-sional scaling (nMDS) ordination of T-RFLP data based on Dice dissimilarities amongst samples. Full and empty symbols corre-spond to chitin-amended and not-amended controls. The size of full symbols increases with increasing chitin concentration

Stress: 0.05

Control, 0d

1.3 g, 0d

Control, 7d

1.3 g, 7d

3.0 g, 7d

7.0 g, 7d

Control, 1 yr

1.3 g, 1 yr

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Polar Biol (2009) 32:105–112 111

communities in the deep Arctic Ocean. Unfortunately, thelack of replication limited the analysis of signiWcance ofour results since the variation within each treatment wasunknown. Therefore, further experiments in other deep-searegions and greater replication are needed to establish thegenerality of these Wndings. To our knowledge, this is theWrst in situ study that describes responses of microbialabundance, activity potential and community structure overa long-term period. Future investigations will be dedicatedto the assessment of the fate and rate of chitin degradationby using radiolabeled substrates. Furthermore, by usingphylogenetic analysis of the microbial communities con-jointly enriched under high chitin inputs and novelapproaches, like MARFISH and stable isotope probing, wewill be able to identify the microbes involved in recyclingthis highly insoluble organic substrate.

Acknowledgments We gratefully like to thank the ship oYcers andcrew of the German research-icebreaker ‘Polarstern’ for their help dur-ing the expeditions to HAUSGARTEN in summers 2004 and 2005.Experiments with our free-falling benthic lander would not have beenpossible without the support of J. Wegner and I. Schewe. We are alsoindebted to I. Kolar for assistance with the sample preparation, bio-chemical analyses. We wish to thank A. Boetius, Gustavo Fonseca andFabiane Gallucci for useful comments and discussions.

Open Access This article is distributed under the terms of the Cre-ative Commons Attribution Noncommercial License which permitsany noncommercial use, distribution, and reproduction in any medium,provided the original author(s) and source are credited.

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