Chemosynthetic symbionts of marine invertebrate
animals are capable of nitrogen fixation
Jillian M. Petersen
1,2
*
, Anna Kemper
2
, Harald Gruber-Vodicka
2
,UlisseCardini
1
,
Matthijs van der Geest
3,4
, Manuel Kleiner
5
, Silvia Bulgheresi
6
, Marc Mußmann
1
,CraigHerbold
1
,
Brandon K.B. Seah
2
, Chakkiath Paul Antony
2
, Dan Liu
5
, Alexandra Belitz
1
and Miriam Weber
7
Chemosynthetic symbioses are partnerships between invertebrate animals and chemosynthetic bacteria. The latter are the
primary producers, providing most of the organic carbon needed for the animal host’s nutrition. We sequenced genomes
of the chemosynthetic symbionts from the lucinid bivalve Loripes lucinalis and the stilbonematid nematode Laxus oneistus.
The symbionts of both host species encoded nitrogen fixation genes. This is remarkable as no marine chemosynthetic
symbiont was previously known to be capable of nitrogen fixation. We detected nitrogenase expression by the symbionts
of lucinid clams at the transcriptomic and proteomic level. Mean stable nitrogen isotope values of Loripes lucinalis were
within the range expected for fixed atmospheric nitrogen, further suggesting active nitrogen fixation by the symbionts.
The ability to fix nitrogen may be widespread among chemosynthetic symbioses in oligotrophic habitats, where nitrogen
availability often limits primary productivity.
S
ymbioses between animals and chemosynthetic bacteria are
widespread in Earth’s oceans
1
. Animals from at least seven
phyla have formed such symbioses, and even more chemosyn-
thetic bacterial lineages have evolved symbioses with animal hosts
1
.
Chemosynthetic symbionts can use a range of chemicals, such as
sulfide, methane, hydrogen and carbon monoxide, to power their
metabolism
2–4
. The hosts of chemosynthetic symbionts dominate
some animal communities. For example, shallow-water lucinid
bivalves, which host sulfur-oxidizing symbionts, often dominate
the macrobenthic infaunal community in seagrass meadows,
where they can reach densities greater than 3,500 individuals per
square metre
5,6
. Their diversity in nature, their persistence over evol-
utionary timescales and their dominance in many habitats attest to
the success of these symbiotic partnerships
1
.
Chemosynthetic symbionts are primarily considered ‘nutritional
symbionts’, meaning their primary role is to provide nutrition for
their hosts
1,7
. So far, most studies have focused on inorganic
carbon fixation by the symbionts and the transfer of fixed organic
carbon compounds to the hosts. In addition to organic carbon, all
animals require a source of fixed nitrogen. However, nitrogen
metabolism in chemosynthetic symbioses has received far less atten-
tion. Chemosynthetic symbionts have been shown to gain their
nitrogen from ammonium or nitrate in their environment
8–10
and
co-occurring nitrogen-fixing and chemosynthetic symbionts have
been found in cold-water corals
11
. Nitrogen fixation by chemo-
synthetic symbionts has long been hypothesized, but so far not
yet shown
12–14
.
Our study focused mainly on the endosymbiosis between
bivalves of the family Lucinidae and sulfur-oxidizing bacteria.
Lucinids are by far the most diverse and widespread group of
bivalves that host chemosynthetic symbionts
15
. There are at least
400 living species, occupying a range of habitats including mangrove
sediments, seagrass beds, coral reef sediments and coastal mud
and sand
16
. In seagrass habitats, lucinid bivalves and their sulfur-
oxidizing symbionts are part of a nested symbiosis with seagrasses,
which may be essential to the health and ecological success of
seagrasses
6
. We focused on the symbiosis between Loripes lucinalis
(Lamarck, 1818) and its endosymbionts. We also investigated a
second symbiosis, that between stilbonematid nematodes and their
sulfur-oxidizing ectosymbionts, because these symbionts are associ-
ated with the family Chromatiaceae, which contains a number of
diazotrophic sulfur oxidizers
17,18
. Nematodes of the subfamily
Stilbonematinae (family Desmodoridae) can be found worldwide
in marine sulfidic habitats
19
. All known species have a dense
coating of ectosymbionts on their cuticle, which are hypothesized
to contribute to their host’s nutrition
19
. The name Candidatus
Thiosymbion oneisti will be proposed elsewhere for the nematode
symbionts (Gruber-Vodicka et al., in preparation). We propose
the name Candida tus Thiodiazotropha endoloripes for the symbiont
of Loripes lucinalis, where ‘Thiodiazotropha’ refers to the sulfur-
oxidizing (‘thio’) and nitrogen-fixing (‘diazotroph’) metabolism of
the symbiont and ‘endoloripes’ (‘Endo-’, Greek from ἔνδον meaning
‘within’, ‘loripes ’) refers to the endosymbiotic association with
Loripes lucinalis, its bivalve host.
Results and discussion
Phylogenomics, and carbon and energy metabolism of
L. lucinalis and Laxus oneistus symbionts. The symbiont draft
genomes from five clam individuals (Ca. Thiodiazotropha
endoloripes A– E) were 100% complete for a set of 281 marker
genes conserved across all Gammaproteobacteria and ranged in
size from 4.46 to 4.88 megabase pairs (Mb) on 12–48 contigs
1
Department of Microbiology and Ecosystem Science, Division of Microbial Ecology, Research Network Chemistry meets Microbiology, University of
Vienna, Althanstrasse 14, Vienna 1090, Austria.
2
Max Planck Institute for Marine Microbiology, Celsiusstrasse 1, Bremen 28359, Germany.
3
Centre for
Marine Biodiversity, Exploitation and Conservation (MARBEC), UMR 9190, IRD-IFREMER-CNRS-UM, Université de Montpellier, Montpellier Cedex 5
34095, France.
4
Department of Coastal Systems and Utrecht University, NIOZ Royal Netherlands Institute for Sea Research, PO Box 59, 1790 AB Den
Burg, Texel, The Netherlands.
5
Department of Geoscience, University of Calgary, 2500 University Drive Northwest, Alberta T2N 1N4, Canada.
6
Archaea
Biology and Ecogenomics Division, Department of Ecogenomics and Systems Biology, University of Vienna, Althanstrasse 14, Vienna 1090, Austria.
7
HYDRA Institute for Marine Sciences, Elba Field Station, Campo nell’Elba, Livorno 54037, Italy.
*
e-mail: petersen@microbial-ecology.net
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PUBLISHED: 24 OCTOBER 2016 | VOLUME: 2 | ARTICLE NUMBER: 16195
OPEN
NATURE MICROBIOLOGY 2, 16195 (2016) | DOI: 10.1038/nmicrobiol.2016.195 | www.nature.com/naturemicrobiology 1
© 2016 Macmillan Publishers Limited, part of Springer Nature. All rights reserved.
(Table 1). Those from two individuals of L. oneistus
(Ca. Thiosymbion oneisti A–B) were 86.75 and 89.11% complete
at sizes of 3.66 and 3.51 Mb on 183 and 193 contigs (Table 1).
We conducted phylogenomic analyses to better understand the
relationships between the bivalve and nematode symbionts and
other symbiotic and free-living gammaproteobacterial sulfur
oxidizers. Consistent with previous analyses based on 16S rRNA
genes
1
, our phylogenomic analysis placed the lucinid symbionts
Ca. Thiodiazotropha endoloripes together in a cluster with
Candidatus Endoriftia persephone, the sulfur-oxidizing symbiont
of the hydrothermal vent tubeworm Riftia pachyptila (Fig. 1).
Also consistent with previous analyses
20
, the nematode symbionts
Ca. Thiosymbion oneisti clustered together with the sulfur-
oxidizing endosymbionts of the gutless oligochaete worm Olavius
algarvensis and were affiliated with free-living sulfur oxidizers
from the family Chromatiaceae (Fig. 1). The L. lucinalis and
R. pachyptila symbiont cluster was not clearly associated with any
known free-living sulfur oxidizers, but formed a sister group to
the clade containing the stilbonematid and oligochaete symbionts
and members of the Chromatiaceae (Fig. 1).
As expected based on previous studies of symbiont metabolism,
genes and pathways for sulfur oxidation and carbon fixation were
found in all five Ca. Thiodiazotropha endoloripes and in both Ca.
Thiosymbion oneisti draft symbiont genomes (Fig. 2). Nematode
Table 1 | Features of bivalve and nematode symbiont genomes.
Genome Size (Mb) No. of contigs No. of genes predicted GC content (%) Completeness estimate* (%)
Ca. Thiodiazotropha endoloripes A 4.46 15 4,193 52.1 100
Ca. Thiodiazotropha endoloripes B 4.61 18 4,381 51.9 100
Ca. Thiodiazotropha endoloripes C 4.46 18 4,226 52.1 100
Ca. Thiodiazotropha endoloripes D 4.5 12 4,301 52.0 100
Ca. Thiodiazotropha endoloripes E 4.88 48 4,685 51.7 100
Ca. Thiosymbion oneisti A 4.44 2,026 4,149 58.71 96.63
Ca. Thiosymbion oneisti B 4.33 1,891 4,050 58.84 96.07
*Completeness estimates were calculated based on how many of 281 conserved gammaproteobacterial marker genes were present in each draft genome. See Methods for details.
Olavius algarvensis γ3 symbiont
Alkalilimnicola ehrlichii
Thiothrix flexilis
Ectothiorhodospira haloalkaliphila
Ectothiorhodospira
sp. PHS1
Sedimenticola thiotaurini*
Ca. Endoriftia persephone
Thiomicrospira halophila
Thiomicrospira crunogena
Thioalkalimicrobium cyclicum
Thioalkalimicrobium aerophilum
0.1
Ca. Thiodiazotropha endoloripes E*
Ca. Thiodiazotropha endoloripes D*
Ca. Thiodiazotropha endoloripes B*
Ca. Thiodiazotropha endoloripes A*
Ca. Thiodiazotropha endoloripes C*
Ca. Thiosymbion oneisti A*
Ca. Thiosymbion oneisti
B*
Thioflavicoccus mobilis
*
Allochromatium vinosum*
Marichromatium purpuratum*
Thiorhodovibrio sp. 970*
Olavius a
l
g
arvensis γ
1 symbiont
L
amprocystis purpurea
Thiorhodococcus drewsii
Chrysomallon squamiferum symbiont
Thiolapillus brandeum
Solemya velum symbiont
Thiomicrospira chilensis
Thiomictospira kuenenii
Ca. Vesicomyosocius okutanii
Ca. Ruthia magnifica
Ca. Thioglobus singularis
Thiobacillus deni
trificans
Ca. Thioglobus autotrophica
Bathymodiolus azoricus symbiont
Bathymodiol
us
sp. SMAR symbiont
SUP05
Hydrogenovibrio marinus
Thio
t
hrix disciformis
Thiothrix lacustris
Thioalkalivibrio nitratireducens
Thioalkalivibrio thiocyanodenitrificans
*Thiothrix nivea
*Thiorhodospira sibrica
*
Be
ggiat
oa alba
Figure 1 | Phylogenomic tree of sulfur-oxidizing Gammaproteobacteria. Phylogenomic tree of free-living and symbiotic sulfur oxidizers from the
Gammaproteobacteria. The 25 single-copy genes used in the analysis were defined based on the AMPHORA2 core bacterial phylogenetic marker database
66
.
SUP05 bin refers to the genomic assembly from the metagenome study by Walsh and co-authors
81
. The betaproteoba cter ial sulfur oxidizer Thiobacillus denitrificans
was used as the outgroup. SH-like support values were above 90% for all nodes of the tree. Genomes encoding nitrogenase genes are indicated with an
asterisk and bold text. Sequences from this study are shown in red.
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and bivalve symbionts encoded a complete tricarboxylic acid (TCA)
cycle and transporters for uptake of organic compounds and thus
have the potential for heterotrophic growth. Both are capable of
using oxygen and oxidized nitrogenous compounds such as nitrate
as terminal electron acceptors, but only the bivalve symbiont draft
genomes encoded genes for uptake hydrogenases, which would
allow them to use hydrogen as an energy source. Bivalve and nema-
tode symbionts both produce intracellular elemental sulfur gran-
ules
21,22
. The genomes also revealed the potential to store organic
carbon in the form of polyhydroxyalkanoates (PHAs) and phosphor-
ous in the form of polyphosphate granules. A detailed comparative
genomics study of the lucinid and stilbonematine symbionts is
beyond the scope of this study and will be published elsewhere.
Symbionts of L. lucinalis and L. oneistus encode nitrogenases.
Surprisingly, the five draft genomes of Ca. Thiodiazotropha
endoloripes and both draft genomes of Ca. Thiosymbion oneisti
contained large clusters of genes involved in nitrogen fixation,
including the structural genes for the iron-molybdenum
dinitrogenase (nifD and nifK), the dinitrogenase reductase subunit
(nifH), ferredoxins, and maturation and regulatory factors (Fig. 3,
Supplementary Discussion and Supplementary Figs 1 and 2). The
nifH gene is commonly used as a functional marker for nitrogen
fixation, so many nifH genes are available in public databases. Our
phylogenetic analyses showed that the sulfur-oxidizing symbiont
sequences clustered together with Group 1 molybdenum-dependent
NifH sequences as defined by Raymond
23
(Fig. 3). The symbiont
sequences fell into a clade containing mainly Gammaproteobacteria
and some Betaproteobacteria. The NifH sequences from both
Ca. Thiodiazotropha endoloripes and Ca. Thiosymbion oneisti
grouped separately to those from other members of the
Chromatiaceae, which could indicate that this gene was acquired by
horizontal gene transfer in these two symbionts. To gain more
insight into the evolutionary history of nitrogen fixation in
Ca. Thiodiazotropha endoloripes and Ca. Thiosymbion oneisti, we
analysed the phylogeny of the NifD proteins, which make up one of
the two structural subunits of the nitrogenase enzyme. The
placement of NifD proteins from the chemosynthetic symbionts was
different to the placement of their NifH proteins (Supplementary
Fig. 3). The NifD from Ca. Thiodiazotropha endoloripes
grouped together with the NifD from Sedimenticola thiotaurini
(Supplementary Fig. 3). This was similar to our phylogenomic
Calvin cycle
(RuBisCO form I)
CO
2
H
+
Biomass
PP
P
HPP
H
2
H
2
ase
e
−
Dsr Apr Sat
HS
−
FCC
S
0
Sox
S
2
O
3
2−
SO
4
2−
SO
4
2−
Dsr
AprM
e
−
e
−
O
2
resp
O
2
Nresp
NO
3
−
NH
4
+
Glutamine
NH
4
+
ABC
Urea
Urea
TCA cycle
PHA
Pyruvate
Intermediates
ActP
Acetate
Nif
N
2
H
2
PolyP
*
*
OM
IM
Flagellum*
Type IV pilus
TRAP
Organic compounds
Nass
NO
3
−
ABC
GlnA
Figure 2 | Overview of the major cellular features and metabolic pathways encoded in bivalve and nematode symbiont genomes. Metabolic enzymes and
enzyme systems are shown in yellow, transporters in brown, storage granules in pink and structural features in grey. Features only encoded in the draft
genomes of Ca. Thiodiazotropha endoloripes and not yet found in the draft genomes of Ca. Thiosymbion oneisti are indicated by an asterisk. IM, inner
membrane; OM, outer membrane; HPP, proton-translocating pyrophosphatase; H
2
ase, uptake hydrogenase; FCC, flavocytochrome c;S
0
, elemental sulfur
granule; Sox, sox enzyme system for sulfur oxidation; Dsr, reverse dissimilatory sulfite reductase; Apr, adenosine phosphosulfate reductase; AprM, adenosine
5′-phosphosulfate membrane anchor; Sat, sulfate adenylyltransferase; O
2
resp, genes for oxygen respiration (cytochrome c oxidases); Nresp, nitrate
respiration (denitrification, pathway complete to N
2
); Nass, assimilatory nitrate reduction; ActP, acetate transporter; ABC, ABC transporter; TRAP, TRAP
transporter; PolyP, polyphosphate granule; PHA, polyhydroxyalkanoate granule; GlnA, glutamine synthetase; Nif, nitrogenase.
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Alphaproteobacteria I (EJC75087.1) 65
Alphaproteobacteria II (EGD60366.1) 27
Burkholderiales spp. (EHR69902.1) 24
EKF67409.1 Herbaspirillum frisingense
ACH83478.1 Acidithiobacillus ferrooxidans ATCC 53993
BAL23130.1 Azoarcus sp. KH32C
EGW62911.1 Dechlorosoma suillum
EGJ10941.1 Rubrivivax benzoatilyticus
ZP_10016521.1 Methylacidiphilum fumariolicum
EES53484.1 Leptospirillum ferrodiazotrophum
EQD24965.1 Leptospirillum sp. Group IV ’UBA BS’
Campylobacterales (ADR34162.1)
6
EFC65041.1 Uncultured bacterium S5
ADR18595.1 Calditerrivibrio nitroreducens DSM 19672
EEJ24463.1 Denitrovibrio acetiphilus DSM 12809
Enterobacteriales (EJI87739.1) 22
ACQ92279.1 Tolumonas auensis DSM 9187
EPM41546.1 Vibrio natriegens ATCC 14048
ACR12580.1 Teredinibacter turnerae T7901
EGZ49204.1 Thiorhodospira sibirica ATCC 700588
AAZ46164.1 Dechloromonas aromatica
ADE11119.1 Sideroxydans lithotrophicus
ACV37675.1 Candidatus Accumulibacter phosphatis
Betaproteobacteria
EIJ42511.1 Beggiatoa alba
AGA91923.1 Thioflavicoccus mobilis
ABP79021.1 Pseudomonas stutzeri A1501
ZP_21946170.1 Pseudomonas stutzeri NF13
ACO76403.1 Azotobacter vinelandii
EIJ36507.1 Thiothrix nivea DSM 5205
EGV19816.1 Thiocapsa marina
EER67145.1 Allochromatium vinosum DSM 180
EGV30156.1 Thiorhodococcus drewsii
EGV20999.1 Marichromatium purpuratum
EGZ55571.1 Thiorhodovibrio sp.
CP011412 Sedimenticola thiotaurini SIP−G1
AEF99883.1 Methylomonas methanica
ABK43713.1 Magnetococcus sp.
12
ABM61068.1 Halorhodospira halophila
Gammaproteobacteria
Desulfuromonadales (EAT15955.1)
12
ACG74424.1 Anaeromyxobacter sp.
ABS27227.1 Anaeromyxobacter sp.
Peptococcaceae (AET70418.1)
9
BAI69593.1 Hydrogenobacter thermophilus
ADC89059.1 Thermocrinis albus DSM 14484
ABZ83695.1 Heliobacterium modesticaldum
AET59369.1 Paenibacillus terrae
CCQ92501.1 Clostridium ultunense
Cyanobacteria (ABA23658.1) 52
Frankia spp. (EFC83885.1)
6
0.10
90−100%
80−89.9%
70−79.9%
Candidatus Thiosymbion oneisti (FLUZ00000000)
Candidatus Thiodiazotropha endoloripes (LVJW00000000)
b
a
Ca. Thiodiazotropha endoloripes
Azoarcus sp. BH72
nifHDKT
Figure 3 | Clusters of nif genes and NifH phylogeny. a, Schematic representation of the nifHDKT gene cluster in the Ca. Thiodiazotropha endoloripes draft
genomes and in the model nitrogen-fixing bacterium Azoarcus sp. BH72. See Supplementary Fig. 2 for an overview of the complete nif cluster. b,Maximum-
likelihood phylogeny of full-length Group 1 NifH proteins. Percentages refer to SH-like support values from aLRT. Sequences from this study are showninbold
red text. Numbers in wedges indicate how many sequences are contained in that collapsed clade. Brackets contain examples of specific protein sequences.
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results, which showed that S. thiotaurini grouped together with the
Chromatiaceae (Fig. 1). The NifD of Ca.Thiosymbiononeisti
grouped together with the NifD from Methylomonas methanica
and not with those from other members of the Chromatiaceae
(Supplementary Fig. 3). In summary, there appears to be a history
of horizontal transfer of the genes encoding both NifH and
NifD, so these proteins could have been horizontally acquired
by the chemosynthetic symbionts of lucinid clams and
stilbonematid nematodes.
Using PCR primers specific for nifH, we screened DNA extracted
from six L. lucinalis individuals, three from France and three from
Mauritania, and DNA from five additional lucinid species from
sampling sites around the world (Supplementary Tables 1 and 2).
We obtained bands of the correct size from all individuals of
L. lucinalis and from three other lucinid species (Euanodontia
ovum, Codakia orbicularis and Clathrolucina costata)
(Supplementary Table 2). We could not obtain a PCR product
from one individual each of Lucinoma borealis and Epidulcina cf
delphinae, although the symbiont 16S rRNA gene could be ampli-
fied from these samples (Supplementary Table 2). The PCR pro-
ducts were directly sequenced and, although they were too short
to determine their phylogenetic position reliably, they were highly
similar to the sequences we obtained from the symbiont genomes.
All nifH sequences from L. lucinalis symbionts were between 92
and 100% identical at the nucleotide level (97–100% identity at
the amino acid level). Among symbionts of different lucinid
species, the nifH sequence identity ranged from 83 to 88% (91–
98% at the amino acid level). Genome sequencing of these symbiont
species would confirm whether they also encode all genes necessary
for nitrogen fixation. However, the presence of the nifH gene raises
the possibility that the symbionts of many lucinid species might be
capable of nitrogen fixation.
Nitrogen-fixing symbioses are common in marine ecosystems,
particularly in habitats where nitrogen availability limits primary
production, such as oligotrophic coral reefs and open ocean
water
24,25
. Chemosynthetic symbionts are well known for their con-
tribution to host nutrition through carbon fixation, but so far, no
marine chemosynthetic symbiont was known to be capable of
nitrogen fixation. Nitrogen fixation has been hypothesized in the
only known chemosynthetic symbiosis in the terrestrial environ-
ment between ectosymbiotic Thiothrix-related bacteria and
Niphargus amphipods, because nifH transcripts could be PCR-
amplified from the Niphargus ectosymbiotic community
26
. In con-
trast to previous PCR-based studies, we could unambiguously
associate the nifH sequence with the rest of the symbiont genome.
The presence of these genes in the symbionts of nematodes and
bivalves, two unrelated hosts from different animal phyla, shows
that the ability to fix nitrogen is not restricted to one phylogenetic
group of symbionts, or to the symbionts of only one animal
group, but may be widespread in chemosynthetic symbioses.
Nitrogen fixation genes are expressed by L. lucinalis symbionts.
To test whether nitrogen fixation genes are actively expressed by
the lucinid symbionts when living in their hosts, we sequenced
the gill metatranscriptomes of five individuals and analysed gill
metaproteomes of another six individuals of L. lucinalis from Elba
(Italy). Transcripts from genes involved in nitrogen fixation were
among the 30 most abundantly expressed genes in two of these
five individuals (Supplementary Fig. 4 and Supplementary Data
set 1). In gill metaproteomes from six L. lucinalis individuals,
between 892 and 1,377 symbiont proteins could be detected
(Supplementary Data set 2). Nitrogenase proteins were detected in
five of these six individuals (Fig. 4 and Supplementary Data set 2).
Nitrogen fixation is therefore one of the metabolic pathways
actively expressed by the symbionts in some L. lucinalis individuals
(see Supplementary Discussion for further details).
It is remarkable that the symbionts in the animal tissue actively
express nitrogenases and seem to be nitrogen-limited (Supplementary
Discussion). Because nitrogen fixation is metabolically costly, it is
often downregulated when other nitrogen sources are available in
the environment
27,28
. As far as we are aware, the concentrations of
nitrate, ammonium and urea have not yet been measured in
lucinid tissues. Animals are known to produce ammonia and urea
as nitrogenous waste products and both of these could be used as
nitrogen sources by the lucinid and stilbonematid symbionts
(Fig. 2). It is possible that symbiont nitrogen fixation is regulated
0.00 0.05 0.10 0.500.450.40
NSAF (%)
P1
P2
P3
P4
P5
P6
Nitrogenase molybdenum-iron protein
subunit beta (A3193_07820)
Nitrogenase molybdenum-iron protein
subunit alpha (A3193_07825)
Nitrogenase iron protein (A3193_07830)
Urea ABC transporter substrate-binding
protein (A3193_00820)
Urease subunit alpha (A3193_00780)
Ammonia channel protein (A3193_00530)
Type I glutamate-ammonia ligase
(A3193_16060)
Figure 4 | Expression of proteins for nitrogen assimilation in Loripes lucinalis. Bar chart showing the abundance of seven symbiont proteins involved in
assimilation of nitrogen from urea, ammonia and dinitrogen gas that were identified by proteomics in the gills of six L. lucinalis individuals (P1–P6). See
Supplementary Data set 2 for the complete data set. NSAF, normalized spectral abundance factor.
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