Influence of Changing Plant Food Sources on the Gut Microbiota of Saltmarsh Detritivores

INVERTEBRATE MICROBIOLOGY

Influence of Changing Plant Food Sources on the GutMicrobiota of Saltmarsh Detritivores

Jessica Dittmer & Jérôme Lesobre & Roland Raimond &

Martin Zimmer & Didier Bouchon

Received: 13 February 2012 /Accepted: 10 April 2012 /Published online: 22 May 2012# Springer Science+Business Media, LLC 2012

Abstract Changes in agricultural land-use of saltmarshesalong the German North Sea coast have favoured the succes-sion of the marsh grass Elytrigia atherica over the long-established Spartina anglica. Consequently, E. atherica rep-resents a potential food source of increasing importance forplant-feeding soil detritivores. Considering the importance ofthis ecological guild for decomposition processes and nutrientcycling, we focussed on two sympatric saltmarsh soil macro-detritivores and their associated gut microbiota to investigatehow the digestive processes of these species may be affectedby changing plant food sources. Using genetic fingerprints ofpartial 16S rRNA gene sequences, we analysed compositionand diversity of the bacterial gut community in a diplopod andan amphipod crustacean in relation to different feedingregimes representing the natural vegetation changes. Effectsof syntopy on the host-specific gut microbiota were also takeninto account by feeding the two detritivore species eitherindependently or on the same plant sample. Bacterial commu-nity composition was influenced by both the host species andthe available plant food sources, but the latter had a stronger

effect on microbial community structure. Furthermore, bacte-rial diversity was highest after feeding on a mixture of bothplant species, regardless of the host species. The gut micro-biota of these two detritivores can thus be expected to changealong with the on-going succession at the plant communitylevel in this environment. Cloning and sequencing of bacterial16S rRNA gene fragments further indicated a host-relatedeffect since the two detritivores differed in terms ofpredominant bacterial taxa: diplopods harboured mainlyrepresentatives of the phyla Bacteroidetes and Gammap-roteobacteria. In contrast, the genus Vibrio was foundfor the amphipod host across all feeding conditions.

Introduction

The agricultural use of saltmarshes along the German Wad-den Sea coast for intensive sheep and cattle grazing has beenlessened or abandoned in most mainland saltmarshes since1991 [1, 2]. As a consequence, these ecosystems currentlyundergo a dynamic succession in the plant communities thathad hitherto been shaped by grazing: grazing-dependentsecondary vegetation mosaics are gradually replaced bycommunities dominated by grazing-sensitive, tall-growingspecies [2–5] such as the sea couch Elytrigia atherica (Link)Kerguélen ex Carreras Martinez (Poaceae). The ongoingexpansion of E. atherica in many abandoned saltmarshes[1–3] could eventually cause the retreat of other specieswhich had formerly been well established in the saltmarshplant communities, e.g. the cord grass Spartina anglicaHubbard (Poaceae), by competitive exclusion. Consequent-ly, E. atherica will have a growing share in the plant litter.

Saltmarsh food webs are detritus-based and soil macro-detrivores are of importance for ecosystem functioning sincethey make nutrients from the plant tissue available for other

J. Dittmer (*) : J. Lesobre : R. Raimond :D. BouchonUMR CNRS 7267, Ecologie et Biologie des Interactions,équipe Ecologie Evolution Symbiose, Université de Poitiers,40 Avenue du Recteur Pineau,86022 Poitiers, Francee-mail: [email protected]

M. ZimmerZoologisches Institut, Christian-Albrechts-Universität zu Kiel,Am Botanischen Garten 9,24118 Kiel, Germany

M. ZimmerFB Organismische Biologie, AG Ökologie, Biodiversität &Evolution der Tiere, Paris-Lodron-Universität Salzburg,Hellbrunner Str. 34,5020 Salzburg, Austria

Microb Ecol (2012) 64:814–825DOI 10.1007/s00248-012-0056-4

organisms via shredding of detrital particles and the releaseof nutrient-enriched faeces [6]. Previous studies indicate thatthere is no functional redundancy in saltmarsh communities.Instead, sympatric detritivore species feeding on similarfood sources have been shown to differ in terms of nutri-tional requirements and digestive capabilities [7, 8]. Withregard to ecosystem processes, this means that detritivore-mediated changes in litter nutrient contents depend on boththe detritivore species and the type of plant substrate [7, 8].Consequently, different detritivores most likely contributedifferently to the overall decomposition and nutrient cyclingand a higher diversity of detritivore species should enhanceecosystem processes and resilience. Important detritivores inWadden Sea saltmarshes are the semi-terrestrial beach fleaOrchestia gammarellus Pallas (Crustacea: Amphipoda: Tali-tridae) and the terrestrial diplopod Cylindroiulus latestriatusCurtis (Myriapoda: Diplopoda: Julidae). Orchestia gammar-ellus is often the dominant detritivore in supra-littoral hab-itats in northwestern Europe [9–12] and C. latestriatus,despite being characteristic for terrestrial habitats, common-ly occurs close to the sea shore [13].

Plant-feeding detritivores have to face the difficulties that(1) vascular plant substrate is low in nitrogen (C/N ratio of29 for E. atherica [14] and 34 for S. anglica [15]), essentialamino acids and vitamins and that (2) the nutrients presentin the plant material are not easily accessible due to thestructural complexity of lignocelluloses [16–18]. Therefore,most plant-feeding arthropods relying on this type of nutri-tion benefit from microbial consortia in the gut that providethe necessary enzymes, break down refractory fibrous foodor provide nutritional supplements that are lacking in thehost’s diet [19–24]. These microbial consortia may at leastin part be acquired from the environment [e.g. via necroph-agy, ingestion of faeces released by other individuals (co-prophagy) or ingestion together with the plant tissue] andthey have been observed to change along with the host’s dietin several invertebrates [25, 26]. However, studies carriedout in the context of land-use changes in alpine pasturesshowed that the gut microbiota of one detritivore (an earth-worm) was rather strongly influenced by different foodsources, while that of another (a diplopod) was more resil-ient to diet changes [26, 27]. This is in line with previousobservations suggesting that one particular plant foodsource cannot necessarily be replaced by another withoutaffecting detritivore nutrition and performance [8]. Consid-ering the refractory nature of vascular plant substrate and theimportance of the gut microbiota for digestive processes, itis not unlikely that these species-specific differences indigestive capabilities are at least partly mediated by thebacterial gut community.

In the light of the large-scale changes at the plant-community level in Wadden Sea saltmarshes, it is of interesthow these changes may affect ecologically relevant species

and, possibly, ecosystem functioning. Considering their im-portant role in decomposition processes, we chose to focuson two sympatric macrodetritivores and their associatedbacterial gut communities as a first approach to address thiscomplex question. In order to represent the dynamic plantsuccession as it occurs in nature, we analysed the intestinalbacterial communities of the detritivores O. gammarellusand C. latestriatus after feeding on S. anglica (well estab-lished in grazed saltmarshes), E. atherica (grazing-sensitivespecies, dominates in abandoned saltmarshes) or a mixtureof both plants (representing intermediate conditions). Bac-terial gut community composition and diversity were deter-mined based on genetic fingerprints of partial 16S rRNAgene sequences and additional phylogenetic informationconcerning the predominating bacterial taxa was obtainedby cloning and sequencing of selected PCR products.

Materials and Methods

Sampling and Feeding Experiment

Specimens of the detritivores O. gammarellus and C.latestriatus were collected in April 2009 from the salt-marshes at Westerhever (54° 23′ N, 8° 40′ E) and St.Peter-Ording (54° 18′ N, 8° 39′ E) on the GermanWadden Sea coast. The saltmarshes in these areas areof natural origin, while an important proportion of thesaltmarshes along the Wadden Sea coast result fromhuman land-use changes [2, 5]. Litter samples of thetwo saltmarsh grasses Elytrigia atherica and Spartinaanglica were collected from the saltmarshes at West-erhever at the same time. Green sprouts were removedby hand and the stems were cut into pieces of 5 cm anddried in a compartment drier at 50 °C for 48 h in orderto determine the exact mass (dry weight) of the plantsubstrate.

In order to simulate the natural vegetation changes, aseries of feeding experiments were set up in the laboratory:groups of 10 individuals of either O. gammarellus or C.latestriatus as well as mixed groups of five individuals ofeach species were placed into plastic boxes (10×17×8 cm3)filled with a 2-cm layer of heat-sterilised sand to allowburrowing behaviour and 8.00±0.01 g (dry weight) of plantlitter from either one or both (4:4) plant species. Hence, ninedifferent plant–detritivore combinations were established(Table 1) and five replicate boxes were created for eachcombination. The boxes were then kept at a constant tem-perature of 15 °C and a photoperiod of 12/12 for 4 months.After that period, one individual per box (one of eachspecies in the cases where both species had been rearedtogether) was randomly picked, thus obtaining five repli-cates per single-species condition and 2×5 replicates (five

Gut Microbiota of Saltmarsh Detritivores 815

of each species) per mixed-species condition. The selectedindividuals were surface-sterilised using DNA Away™(Molecular BioProducts, Inc.) to ensure that there is nocross-contamination during dissection and that only internalbacterial communities are detected during subsequent anal-yses and were then stored in ethanol (≥99.8 %) until furtherexperimentation.

DNA Extraction

The gastrointestinal tracts were dissected and crushed in5 μl of proteinase K and 400 μl of extraction buffer[100 mM Tris (pH 8.0), 100 mM EDTA, 100 mM NaCl,0.1 % SDS, 50 mM DTT] and incubated at 37 °C for 4 h.DNAwas extracted twice with phenol–chloroform [1:1 (vol/vol)] and once with chloroform [28] and precipitated with3 M sodium acetate (pH 7) and isopropyl alcohol [1/10:1(vol/vol)] overnight at −20 °C. After centrifugation at15,000×g at 4 °C for 30 min, DNA pellets were resuspendedin sterile water and stored at 4 °C.

PCR Amplifications

For temperature gradient gel electrophoresis (TGGE), a 196-bp fragment spanning the variable V3 region of the bacterial16S rRNA gene was amplified from all DNA extracts usingthe universal primers 338F-GC and 520R [29, 30]. Toensure a stable melting behaviour during TGGE, a 42-nucleotide GC clamp was added to primer 338F, preventingthe complete denaturation of the two DNA strands duringmigration [29, 30]. PCR reaction mixtures of 50 μlcontained 1 μl (80 ng) of template DNA, 1× GoTaq reactionbuffer (Promega), 250 μM of each dNTP, 0.5 μM of eachprimer and 0.5 U of GoTaq DNA polymerase (Promega).PCR cycles consisted of an initial denaturation at 94 °C for2 min, followed by seven cycles of 94 °C for 30 s, 55 °C for30 s and 72 °C for 1 min; another 21 cycles of 92 °C for30 s, 50 °C for 30 s, 72 °C for 1 min and 21 s and a finalextension at 72 °C for 10 min [30].

For cloning and sequencing, primers 27F and 973R [31,32] were used to amplify a 965-bp fragment of the bacterial16S rRNA gene including the variable V3 region. The 25-μlreaction mixtures contained 0.5 μl of template DNA in 1×GoTaq reaction buffer (Promega), 215 μM of each deoxy-nucleoside triphosphate (dNTP), 1 μM of each primer and0.6 U of GoTaq DNA polymerase (Promega). PCR condi-tions were as follows: 95 °C for 3 min; 35 cycles of 95 °Cfor 1 min, 50 °C for 1 min and 72 °C for 1 min 20 s; and 72 °C for 10 min.

All PCR amplifications were performed in a MyCyclerthermocycler (Bio-Rad) and confirmed by electrophoresis in1.5 % agarose gels followed by staining with ethidiumbromide.

TGGE Fingerprints

The Dcode Universal Mutation Detection System™ (Bio-Rad) was used to obtain 16S rRNA gene fingerprints byTGGE [29]. 15 μl of the V3 PCR products were run on 8 %polyacrylamide gels [8 M urea, 20 % formamide, 10 %ammonium persulfate, tetramethylethylenediamine(TEMED) and 50× TAE buffer (40 mM Tris, 20 mM aceticacid, 1 mM EDTA)] in 1.25× TAE running buffer. Electro-phoresis was performed at 75 V across a temperature gradi-ent from 36 °C to 52 °C (constant ramping of 0.9 °C/h).After electrophoresis, gels were stained with ethidium bro-mide and photographed under UV. In each run, a set of ninesamples (one from each feeding condition, Table 1) wasadded to allow for the standardisation of gels from differentTGGE runs.

TGGE profiles were analysed using the Bio-1D software(Vilber Lourmat Inc.). The position of each band was deter-mined via the standardised retention factor (Rf) values, result-ing in a presence/absence matrix of each band per sample.Differences in microbial community composition dependingon host species and different feeding conditions were analysedby principal component analysis (PCA) and between-groupanalysis (BGA) [33]. The statistical significance of the BGA

Table 1 Detritivore–plant feeding experiments

Plant Detritivore

C. latestriatus(diplopod)

O. gammarellus(amphipod)

C. latestriatus +O. gammarellus

E. atherica DE AE DAE + ADE

S. anglica DS AS DAS + ADS

E. atherica + S. anglica DSE ASE DASE + ADSE

Note that combinations involving both detritivores represent two different conditions in order to distinguish between individuals of either species

A amphipod, D diplopod,E Elytrigia atherica, S Spartina anglica

816 J. Dittmer et al.

was verified by a Monte Carlo randomisation test on thebetween-groups inertia percentage (10,000 permutations). Sec-ondly, mean species richness as well as the Shannon Indices ofdiversity and evenness [34] were calculated for each conditionbased on the position (Rf) and relative abundance (i.e. intensi-ty) of the bands. The Kruskal–Wallis rank sum test followed bya multiple comparison test was used to identify significantdifferences in mean species richness and diversity indicesbetween different experimental conditions. All statistical anal-yses were done using the R software (R Project 2.14.0).

Cloning and Sequencing

PCR amplicons (obtained using the primer pair 27F and973R) from two replicate individuals per condition involv-ing only one host species, and one replicate from eachspecies per condition combining both hosts, were chosenfor subsequent cloning and sequencing. The PCR productswere purified using the QIAquick Spin PCR Purification kit(Qiagen) and cloned into competent Escherichia coli JM109vector cells using the pGEM-T Easy Vector Systems kit(Promega) according to the manufacturer’s instructions. Re-combinant clones were amplified with the sequencing pri-mers M13F and M13R (Promega). The resulting PCRproducts were purified using the ExoSAP-IT purificationkit (USB) and the sequencing reaction was done using theBigDye Terminator kit (Applied Biosystems) according tothe manufacturer’s instructions. Sequencing was performedin an ABI 310 sequencer (Applied Biosystems).

The sequences were analysed using the BLASTN searchtool (http://blast.ncbi.nlm.nih.gov) and the RDP Classifier(Ribosomal Database Project, http://rdp.cme.msu.edu). Bothtools yielded congruent results. The Blast output was thenused to create phylogenetic trees using MEGAN 4.0 (MEta-Genome ANalyzer, [35]) based on an LCA-assignment algo-rithm (LCA 0 lowest common ancestor) applied to the top10 % of the Blast output. Rarefaction curves were generatedbased on the expected species richness calculated using Esti-mateS 8.2.0 (Colwell 2009, http://purl.oclc.org/estimates) inorder to confirm that our sequencing effort adequately repre-sents the species richness present in both detritivores. Allsequences obtained in this study were deposited in the EMBLNucleotide Sequence Database (http://www.ebi.ac.uk/embl/)under accession numbers FR848093 to FR848154.

Results

Bacterial Community Composition Based on TGGEFingerprints

Regardless of the host species, TGGE banding profilesdiffered between feeding conditions in that a higher number

of bands (indicating a higher number of bacterial strains)were found when the animal(s) had been fed on both plants.This is mainly due to the fact that two bands, characterisedby a distinctively lower Rf, occurred only in those condi-tions and were absent from all conditions where the detri-tivores had been fed on one plant only. This was furthersupported by a principal component analysis (PCA) usingthe first two principal axes, which indicated that the bandsB1 and B2 are correlated and hence likely to co-occur in thesame individual (data not shown).

A between-group analysis (BGA) based on the differentplant food sources showed that the differences in bandingpatterns (i.e. bacterial community structure) due to differentfeeding conditions (i.e. Spartina, Elytrigia and Spartina +Elytrigia) explained 32.5 % of the total variation. This resultwas statistically significant (p<0.0001) based on a MonteCarlo test with 10,000 permutations and resulted in a dis-tinctive graphical repartition of the bands depending on thethree feeding conditions (Fig. 1a). Such a clear separationwas not observed when the different host species were usedas grouping factors for the BGA (Fig. 1b). However, the

a

b

Figure 1 Between-group analysis (BGA) depending on a differentfeeding conditions and b different host species. Labels indicate thegravity centres of the groups


http://blast.ncbi.nlm.nih.gov

http://rdp.cme.msu.edu

http://purl.oclc.org/estimates

http://www.ebi.ac.uk/embl/

gravity centres of the different rearing conditions (i.e.single-species groups and mixed detritivore associations)were still clearly repartitioned along the two axes: the con-ditions were vertically subdivided depending on the hostspecies, while single-species groups were horizontally sub-divided from detritivore associations (Fig. 1b). Statistically,the differences in banding patterns due to different hostspecies still explained 8.8 % of the total variation (p00.04after 10,000 permutations). These results indicate that, al-though the composition of the gut microbiota is influencedby both the host species and the plant food source, the latterhas a stronger effect on microbial community structure.

Microbial Diversity

Mean species richness, diversity (Shannon H) and evennesswere calculated for all conditions based on the number andintensity of TGGE bands. Evenness was not significantlydifferent across experimental conditions (mean±SE00.87±0.04), indicating that the microbial communities did notcontain any dominating taxa. Both species richness(Fig. 2a) and diversity (Fig. 2b) were generally higher whenthe detritivores had been fed on both plant species, withmean species richness ranging from 3.0±0.3 to 4.6±0.24and mean diversity ranging from 1.05±0.1 to 1.4±0.03. Incontrast, mean species richness remained between 2.0±0.3and 3.0±0 and Shannon H values between 0.6±0.1 and 0.84±0.04 throughout all conditions that involved only one plantspecies. The highest overall species richness (4.6±0.24) aswell as the highest diversity (1.43±0.03) were found fordiplopods fed on both plants. These values were significant-ly higher (species richness—Kruskal–Wallis chi-squared034.1, df011, p<0.001; Shannon H—Kruskal–Wallis chi-squared032.56, df011, p<0.001) than those obtained forthe conditions representing the lowest mean species richnessand/or the lowest diversity (both detritivore species rearedtogether and fed on E. atherica only, amphipods rearedtogether with diplopods and fed on S. anglica; Fig. 2a, b).

Predominant Bacterial Taxa

The entire sequence dataset was assigned to bacterial taxa andis represented in Table 2. Despite the relatively low number ofsequences, rarefaction curves representing the expected spe-cies richness present in both host species reach a plateauphase, indicating that our sequencing effort was sufficient tocapture the most common taxa (Fig. 3). Although the majorityof the sequences were assigned to the phyla Proteobacteria(53 % of all sequences, n033) and Bacteroidetes (27 %, n017) based on a conservative LCA algorithm, both detritivorespecies harboured different bacterial assemblages throughoutall conditions (Fig. 4). For the amphipod O. gammarellus, wefound numerous representatives of the Gammaproteobacteria

across all feeding conditions in single-species groups. Thesequences that could be further identified were mostlyassigned to the genus Vibrio (52 % of all sequences from theamphipod host, n012). In addition, the genus Granulosicoc-cus was detected in individuals that had been feeding on S.anglica and the genus Flavobacterium (phylum Bacteroi-detes) was found after feeding on both plants.

The results for the diplopod C. latestriatus showed a higherdegree of variability: members of the Gammaproteobacteria(that could not be further identified) appeared in individualsfrom single-species groups across all feeding conditions (27 %of all sequences from the diplopod host, n06). The phylumBacteroidetes (in part further identified as belonging to the classBacteroidia) was equally represented, but it was not observed indiplopods that had been fed on E. atherica alone. Sequencesassigned to the Alphaproteobacteria (Acetobacteraceae) were

a

b

Figure 2 Species richness (a) and Shannon’s diversity index (b) forthe different feeding conditions. A amphipod, D diplopod


Tab

le2

Phy

logenetic

affiliatio

nof

16SrRNA

gene

sequ

encesob

tained

inthisstud

ybasedon

both

Megan

andBlastresults

Host

Nutritio

nClone

name

Size

(bp)

Accession

no.

Megan

Blast

LCA

Closestrelatedorganism

Accession

no.

Identity

score(%

)Sou

rce

Amph

ipod

S.an

glica

11A-1

703

FR84

8100

Vibrio

sp.

Vibrio

sp.OM02

AB03

8029.1

679/710(96%)

Marineenvironm

entalsample

11A-2

692

FR84

8101

Granulosicoccus

sp.

Granu

losicoccus

sp.Z27

1FJ535

355.1

680/691(98%)

Leavesof

seagrass

Zostera

marina

11A-3

724

FR848102

Gam

maproteobacteria

Granu

losicoccus

sp.ZS4-22

FJ889674.1

663/734(92%)

Atlantic

oceansediment

11A-4

743

FR84

8103

Granulosicoccus

sp.

Granu

losicoccus

sp.Z27

1FJ535

355.1

722/736(98%)

Leavesof

seagrass

Zostera

marina

15A-1

729

FR84

8104

Vibrio

sp.

Vibrio

sp.GWS-TZ-H

188

AY332204.1

704/733(96%)

German

WaddenSea

sediment

15A-2

734

FR84

8105

Vibrio

sp.

Vibrio

litoralisstrain

MANO22P

DQ09

7524

.172

0/736(98%)

Tidal

flat

15A-3

733

FR84

8106

Vibrio

sp.

Vibrio

sp.GWS-TZ-H

188

AY332204.1

702/734(96%)

German

WaddenSea

sediment

15A-4

726

FR84

8107

Vibrionaceae

Vibrio

sp.T-S2-8

FJ176467.1

691/736(94%)

Saltm

arsh

E.atherica

31A-1

730

FR84

8124

Vibrio

sp.

Vibrio

sp.GWS-TZ-H

188

AY332204.1

708/733(97%)

German

WaddenSea

sediment

31A-2

602

FR84

8125

Vibrio

sp.

Vibrio

sp.MM12

JN79

1370

.158

7/605(97%)

Marinesediment

31A-3

592

FR84

8126

Vibrio

sp.

Vibrio

sp.MM12

JN79

1370

.157

7/595(97%)

Marinesediment

34A-2

390

FR84

8127

Vibrio

sp.

Vibrio

sp.UDC49

9HM0320

18.1

380/390(98%)

Sea

water

34A-3

555

FR84

8128

Bacteria

Unculturedbacterium

clon

eMF-Jan-63

HQ225136.1

531/557(97%)

Environmentalsample

34A-5

439

FR848129

Gam

maproteobacteria

Vibrio

sp.OM02

AB03

8029.1

417/438(95%)

Marineenvironm

entalsample

34A-6

383

FR848130

Gam

maproteobacteria

Unculturedproteobacterium

clone24

7AJ310689.1

370/390(95%)

Environmentalsample

S.an

glica+E.

atherica

51A-2

671

FR84

8142

Flavoba

cterium

sp.

Flavoba

cterium

sp.MTFA

JN820147.1

634/673(94%)

Environmentalsample

51A-5

644

FR84

8143

Vibrio

sp.

Vibrio

sp.OM02

AB03

8029.1

620/651(96%)

Marineenvironm

entalsample

51A-6

673

FR84

8144

Flavoba

cterium

sp.

Flavoba

cterium

sp.MTFA

JN820147.1

645/676(95%)

Environmentalsample

51A-7

661

FR84

8145

Flavoba

cterium

sp.

Flavoba

cterium

sp.HME61

20HM7769

81.1

646/663(97%)

Pon

d

53A-2

523

FR84

8146

Vibrio

sp.

Vibrio

sp.UDC49

9HM0320

18.1

502/514(98%)

Sea

water

53A-4

442

FR84

8147

Vibrio

sp.

Vibrio

sp.UDC49

9HM0320

18.1

423/442(96%)

Sea

water

53A-7

716

FR84

8148

Flavoba

cterium

sp.

Flavoba

cterium

sp.HME61

20HM7769

81.1

698/718(97%)

Pon

d

53A-8

463

FR84

8149

Vibrio

sp.

Vibrio

sp.GWS-TZ-H

188

AY332204.1

428/463(92%)

German

WaddenSea

sediment

Amph

ipod

[+diplop

od]

S.an

glica

16A-1

749

FR84

8108

Bacteria

Pseud

oalteromon

assp.BSs20068

EU365491.1

745/749(99%)

Arctic

Ocean

marinesediments

16A-2

761

FR848109

Bacteroidetes

Bacteroidetes

bacterium

CECT7477

FN554868.1

706/734(97%)

MediterraneanSea

water

16A-3

751

FR84

8110

Bacteroidetes

Marinebacterium

HP28

AY24

1563

.168

1/748(91%)

Organic

aggregates

from

German

WaddenSea

16A-4

724

FR848111

Bacteroidetes

Flavoba

cteria

bacterium

Yb0

09AB49

6666.1

703/726(97%)

Sea

water

E.atherica

36A-2

719

FR84

8131

Bacteria

Unculturedbacterium

clon

eM3m

1-12

3JN

092226

.166

8/729(92%)

Gut

ofNephrop

sno

rvegicus


Tab

le2

(con

tinued)

Host

Nutritio

nClone

name

Size

(bp)

Accession

no.

Megan

Blast

LCA


Accession

no.

Identity

score(%

)Sou

rce

36A-8

738

FR84

8132

Bacteria

Unculturedbacterium

cloneC13

DQ34

0200

.168

5/72

0(95%)

Gut

oflong

-jaw

edmud

sucker

Gillichthysmirabilisfrom

Californian

saltm

arshes

S.an

glica+E.

atherica

56A-1

690

FR84

8153

Vibrio

sp.

Vibrio

sp.cloneA7

GU211017.1

680/690(99%)

Hepatopancreasof

intertidal

isopod

Ligia

palla

sii

56A-15

686

FR84

8154

Vibrio

sp.

Vibrio

sp.SM05

09EU5480

74.1

670/69

2(97%)

Decayed

brow

nalgae

Diplopod

6D-1

756

FR848093

Bacteroidetes

Flavoba

cterium

sp.PB00

2GQ33

9900

.174

5/75

7(98%)

Stream

sediment

6D-2

729

FR848094

Acetobacteraceae

UnculturedAcetoba

cteraceae

clon

eElev_16

S_1

212

EF0198

54.1

692/72

8(96%)

Soil

S.an

glica

6D-4

778

FR848095

Bacteria

Unculturedalphaproteobacterium

clon

eNLS3.24

HQ39

7111.1

752/77

8(97%)

Soil

6D-5

712

FR848096

Acetobacteraceae

UnculturedAcetoba

cteraceaeclone

MWM2-34

HQ67

4805

.167

2/71

7(94%)

Env

iron

mentalsample

9D-1

728

FR848097

Gam

maproteobacteria

Gam

maproteobacterium

TUD-JY37

GU206550.1

676/733(92%)

Environmentalsample

9D-3

678

FR84

8098

Gam

maproteob

acteria

Unculturedbacterium

clone55

-P2

AB53

4002.1

609/68

3(89%)

Env

iron

mentalsample

9D-4

717

FR84

8099

Bacteroidetes

Unculturedbacterium

cloneIC70

FJ798

899.1

682/72

1(95%)

Gut

ofCylindroiulus

fulviceps

(Diplopoda)

E.atherica

26D-1

745

FR848116

Gam

maproteobacteria

Unculturedbacterium

clone26B3-C1

FN814085.1

725/743(98%)

Lactuca

sativaph

yllosphere

26D-5

610

FR848117

Firmicutes

UnculturedVeillonellaceae

clon

eM1-34

JN65

3052

.160

3/61

0(99%)

Gut

ofmillipedeTachypod

oiulus

niger

26D-6

694

FR84

8118

Proteob

acteria

UnculturedDesulfovibrionaceae

clon

eM1-28

JN65

3048

.167

6/69

7(97%)

Gut

ofmillipedeTachypod

oiulus

niger

26D-7

590

FR848119

Firmicutes

UnculturedFirmicutes

cloneA27

AM713389.1

516/598(87%)

Lakesediment

29D-2

383

FR848120

Clostridiales

Unculturedbacterium

clone

BHSD_aaa04

h07

EU4744

82.1

357/37

9(94%)

Sheep

faeces

29D-3

371

FR848121

Gam

maproteobacteria

Unculturedbacterium

clone

O23

9306

C11

FN8669

21.1

369/37

1(99%)

River

water

29D-4

351

FR84

8122

Bacteria

Unculturedbacterium

clone16

5-12

HM99

6804.1

279/34

1(82%)

Ant

Ecitonbu

rchellii

29D-9

436

FR848123

Clostridiales

Unculturedbacterium

clone

LSW-L1-53

AJF745391.1

407/443(92%)

PitcherplantSa

rracenia

purpurea

S.an

glica+E.

atherica

46D-1

674

FR84

8135

Bacteroidetes

Unculturedbacterium

cloneIC17

FJ798

846.1

665/68

1(98%)

Gut

ofCylindroiulus

fulviceps

(Diplopoda)

46D-2

675

FR84

8136

Bacteroidia

Unculturedbacterium

cloneIC2

FJ798

831.1

655/68

5(96%)

Gut

ofCylindroiulus

fulviceps

(Diplopoda)

46D-3

679

FR848137

Gam

maproteobacteria

Unculturedbacterium

clone

2B16

sC08

AB53

3876.1

611/68

5(89%)

Env

iron

mentalsample


Tab

le2

(con

tinued)

Host

Nutritio

nClone

name

Size

(bp)

Accession

no.

Megan

Blast

LCA


Accession

no.

Identity

score(%

)Sou

rce

46D-4

678

FR848138

Gam

maproteobacteria

Unculturedbacterium

clone

2B16

sC08

AB53

3876.1

610/684(89%)

Env

iron

mentalsample

49D-2

437

FR848139

Bacteroidia

Unculturedbacterium

cloneIC2

FJ798831.1

415/443(94%)

Gut

ofCylindroiulus

fulviceps

(Diplopoda)

49D-3

462

FR84

8140

Bacteria

Unculturedbacterium

clone

KNR2007

11-126

EU8812

09.1

391/468(84%)

Soil

49D-4

362

FR848141

Bacteroidetes

Unculturedbacterium

cloneIC2

FJ798831.1

353/361(98%)

Gut

ofCylindroiulus

fulviceps

(Diplopoda)

Diplopod

[+am

phipod]

S.an

glica

16D-1

746

FR848112

Bacteroidetes

Unculturedbacterium

cloneIC2

FJ798831.1

710/749(95%)

Gut

ofCylindroiulus

fulviceps

(Diplopoda)

16D-2

736

FR848113

Bacteroidia

Unculturedbacterium

cloneIC2

FJ798831.1

713/740(96%)

Gut

ofCylindroiulus

fulviceps

(Diplopoda)

16D-3

757

FR848114

Bacteroidia

Unculturedbacterium

cloneIC7

FJ798836.1

731/750(97%)

Gut

ofCylindroiulus

fulviceps

(Diplopoda)

16D-4

708

FR848115

Bacteroidia

Unculturedbacterium

cloneIC95

FJ798924.1

685/713(96%)

Gut

ofCylindroiulus

fulviceps

(Diplopoda)

E.atherica

36D-3

699

FR848133

Gam

maproteobacteria

Unculturedbacterium

isolate

1112

8652

6160

7HQ12

0577

.169

0/700(99%)

Soil

36D-8

712

FR84

8134

Gam

maproteob

acteria

Unculturedbacterium

cloneHC-11

AB25

5115

.170

7/710(99%)

Env

iron

mentalsample

S.an

glica+E.

atherica

56D-1

657

FR84

8150

Serratia

sp.

Serratia

sp.JSC-N

623-1

JF9581

41.1

639/658(97%)

Soil

56D-3

675

FR84

8151

Serratia

sp.

Serratia

sp.JSC-N

623-1

JF9581

41.1

658/676(97%)

Soil

56D-4

652

FR84

8152

Bacteria

Unculturedbacterium

cloneIC50

FJ798

879.1

647/652(99%)

Gut

ofCylindroiulus

fulviceps

(Diplopoda)

LCAlowestcommon

ancestor


observed in individuals fed only on S. anglica and someFirmicutes (with several sequences being further assigned tothe order Clostridiales) occurred in diplopods fed onE. athericaas the only food source. After feeding on both plants, thebacterial consortia only consisted of unidentified Gammapro-teobacteria and members of the Bacteroidetes (Bacteroidia).

In the cases where the animals had been reared in associ-ation with the respective other detritivore species, sequencesfrom the amphipod O. gammarellus were still assigned to thegenus Vibrio after feeding on both plants and to the phylumBacteroidetes (not further identified) after feeding on S.anglica. Regarding C. latestriatus, members of the Gammap-roteobacteria were detected after feeding on E. atherica aloneor on both plants. In the latter case, several sequences werealso assigned to the formerly undetected genus Serratia.Sequences belonging to the phylum Bacteroidetes, some ofwhich being assigned to the class Bacteroidia, were obtainedfrom individuals that had been fed on S. anglica alone.

In addition, with regard to the origin of the sequencesshowing the highest Blast scores, the sequences obtained

from amphipods were similar to sequences obtained frommarine environments (Table 2). Moreover, some of thesesequences represent isolates from the German Wadden Sea(accession numbers AY241563.1 and AY332204.1). In con-trast, the sequences obtained from diplopod hosts were moreclosely related to isolates from either terrestrial habitats (e.g.soil, accession numbers DQ123777.1, EF019854.1,EU881209.1, HQ283405.1 and HQ397111.1) or the gut ofthe closely related diplopod species Cylindroiulus fulviceps(accession numbers FJ798846.1, FJ798879.1, FJ798831.1,FJ798836.1 and FJ798924.1).

Discussion

The composition of the gut microbiota was found to differdepending on the available plant food sources (i.e. Elytrigia,Spartina or a combination of both) in two taxonomicallydistant detritivores, the terrestrial diplopod C. latestriatusand the semi-terrestrial amphipod O. gammarellus. Further-more, microbial richness and diversity were higher afterfeeding on a mixture of both plant species. Based on theseobservations, we hypothesise that a substantial proportion ofthe bacteria present in the gut had been ingested along withthe plant tissue. A scenario that would further explain ourresults could be as follows: The two plant species arecolonised by different bacterial consortia, which areingested with the plant tissue by animals feeding on theseplants and remain at least temporarily viable in the gut. Thiswould result in food source-dependent bacterial communi-ties in the gut, with an increased microbial richness anddiversity when the bacterial consortia from both plant spe-cies are combined, exactly as it is observed here.

Alternatively, the higher bacterial diversity observed afterfeeding on both plants could also be due to rare bacterialstrains, which, despite being present in the gut, remain unde-tected unless their prevalence is enhanced by particularly ben-eficial feeding conditions, e.g. feeding on a less restricted diet.This hypothesis is further supported by the observation thatsome recurrent bacterial taxa occur only after feeding on bothplants, while they are absent from all conditions involving onlyone plant species. Hence, combining different plant food sour-ces results in a bacterial gut community that is different from amere combination of the bacterial communities present afterfeeding on either one of the two plant species alone.

Although the bacterial communities associated with thedifferent plant species were not included in the analysis, differ-ences related to plant microbiota across feeding conditionswere limited since for each plant species, a single sample hadbeen initially collected in the field. Consequently, all feedingconditions involving the same plant species but differentdetritivores received plant substrate from the same initial plantsample. This underlines that the observed differences in

a

b

Figure 3 Rarefaction curves representing the expected species rich-ness depending on the number of sequences for a the amphipod O.gammarellus and b the diplopod C. latestriatus. Error bars correspondto 95 % confidence limits


microbial community structure are more likely due to a plant-related effect than to different host species. If the latter was thecase, different host species fed on the same plant samplewould have shown host-specific bacterial communities ratherthan plant-specific differences as it is the case here.

The bacterial diversities may seem rather low comparedto previous studies on the microbial gut communities ofother plant-feeding invertebrates, especially termites [36,37]. This might in part be due to the fact of having oven-dried the plant samples: While this measure allowed us tocontrol the amount of plant substrate used for the feedingexperiments, it may have eliminated some plant-associatedbacteria. Hence, we cannot assume to have detected theentire range of plant-derived gut bacteria as they may bepresent in these hosts under natural conditions. This is allthe more important in the light of our observation that themicrobial communities analysed here might to a certaindegree consist of transient bacteria ingested with the plantfood. However, the suspected elimination of some (but mostlikely not all) plant-associated bacteria due to the planttreatment increases the probability of detecting potentialindigenous gut symbionts, i.e. bacteria more closely associ-ated with the host species than transient gut passengers.

Therefore, the main goal of the sequencing analysis wasnot to carry out an exhaustive phylogenetic analysis but rather

to identify predominant bacterial taxa that occur across allconditions. Interestingly, although the majority of the sequen-ces were assigned to the two phyla Proteobacteria and Bacter-oidetes, both detritivores harboured different bacterialassemblages. This is in line with our observations based onthe TGGE profiles, which indicated that microbial communitycomposition was also influenced by the host species, albeitless strongly than by the plant food source. Regarding thehost-specific bacterial taxa in more detail, the amphipod O.gammarellus carried representatives of the Gammaproteobac-teria (mainly the genus Vibrio) across all feeding conditions,as well as Flavobacterium after feeding on the combination ofboth plants. In C. latestriatus, unidentified Gammaproteobac-teria occurred independently of the food source, but represen-tatives of additional phyla varied depending on the feedingconditions and included Alphaproteobacteria (family Aceto-bacteraceae), Bacteroidetes (class Bacteroidia) and Firmicutes(order Clostridiales).

Concerning the hypothesis that the bacterial gut commu-nities originate at least partly from the plant substrate, thefate of the bacteria after ingestion by detritivores should alsobe considered. Most of them might be only transient gutpassengers before being egested. However, it is interestingthat Vibrio is present in amphipods across all feeding con-ditions in equally high abundances but entirely absent in

a

b

Figure 4 Phylogenetic affiliation of the gut bacteria in a the amphipodO. gammarellus and b the diplopod C. latestriatus after rearing insingle-species groups. Circle size indicates the number of sequences

assigned to the respective node and colours refer to the different foodsources (white 0 E. atherica; black 0 S. anglica; grey 0 both plants)


diplopods, even when the two detritivores had been reared insyntopy and fed on the same plant substrate. Thus, presenceand abundance of Vibrio in the amphipods do not seem to beaffected by different diets, nor by other bacterial taxa that dochange depending on the food source. Previous work byKnapp et al. [27] showed that the composition of the gutmicrobiota in a diplopod was only partly affected by changingfood sources in an alpine environment. Instead, several bac-terial taxa seemed to form a stable community resilient to dietshifts. Considering this example as well as our results regard-ing the occurrence of Vibrio, we hypothesise that this bacteri-um is more stably associated with the amphipod and not just atransient passenger. Furthermore, the genera Vibrio and Fla-vobacterium are among the most commonly reported gutbacteria of aquatic invertebrates [38–41], which supports ourfinding that Vibrio occurs in the semi-terrestrial amphipod,while it is absent in the fully terrestrial diplopod. Other bac-terial taxa identified here have also been associated with thegut microbiota of other species. For instance, bacterial strainsbelonging to the Gammaproteobacteria as well as to the Fir-micutes and Bacteroidetes have been isolated from numeroussoil invertebrates [26, 42–44].

The observed host-related variation in recurrent bacterialtaxa is most likely due to the ecological differences betweenthe two detritivores: While most sequences from the semi-terrestrial amphipods could be traced back to isolates frommarine (or even specifically Wadden Sea) environments,sequences obtained from diplopods were more closely relatedto terrestrial (e.g. soil) habitats and/or gut isolates from anotherdiplopod species. Hence, the microbiota of these detritivoresmay, at least partly, be considered host specific. However,independent of the origin, both detritivores were found toharbour bacterial taxa that varied depending on the feedingconditions. This aspect strengthens the significance of thebacterial community profiles by TGGE fingerprinting, whichclearly indicate that different plant food sources have a strongereffect on the gut microbiota than different host species.

In conclusion, the present work indicates that the gutmicrobiota of two saltmarsh detritivores is strongly influ-enced by the available plant food sources, except for thepresence of Vibrio in amphipods across all feeding condi-tions. Hence, bacterial gut communities can be expected tochange along with the on-going succession at the plantcommunity level in this environment. This is of interestsince the two species studied here represent the dominantmacrodetritivores in these ecosystems and their gut micro-biota may thus play an important role at the ecosystem levelvia its contribution to decomposition and nutrient cycling.Since many aspects of the nutritional interactions betweenthe two detritivores and their bacterial consortia remain asyet unexplored, it is impossible to predict how exactly ashift in the gut microbiota might translate into detritivore-mediated decomposition processes in the saltmarshes. For

instance, it is unknown whether the resulting microbial com-munity will be more efficient for the decomposition of thenewly dominant plant substrate and/or nutrient transfer to thedetritivore. However, if the opposite was the case, this wouldnegatively affect the functional role of the detritivores and,eventually, influence nutrient cycling in this ecosystem.

Acknowledgements The authors would like to thank Philipp Eereveldfor providing the animal samples used in this work as well as the technicalstaff at the Laboratoire Écologie, Évolution, Symbiose (LEES) for theirassistance and Gaël Freyssinel for his advice concerning biodiversityanalyses in R.

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Influence of Changing Plant Food Sources on the Gut Microbiota of Saltmarsh Detritivores

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