The core microbiome of sessile ciliate Stentor coeruleus is not shaped by the environment

Article metrics


Microbiomes of multicellular organisms are one of the hottest topics in microbiology and physiology, while only few studies addressed bacterial communities associated with protists. Protists are widespread in all environments and can be colonized by plethora of different bacteria, including also human pathogens. The aim of this study was to characterize the prokaryotic community associated with the sessile ciliate Stentor coeruleus. 16S rRNA gene metabarcoding was performed on single cells of S. coeruleus and on their environment, water from the sewage stream. Our results showed that the prokaryotic community composition differed significantly between Stentor cells and their environment. The core microbiome common for all ciliate specimens analyzed could be defined, and it was composed mainly by representatives of bacterial genera which include also potential human pathogens and commensals, such as Neisseria, Streptococcus, Capnocytophaga, Porphyromonas. Numerous 16S rRNA gene contigs belonged to endosymbiont “Candidatus Megaira polyxenophila”. Our data suggest that each ciliate cell can be considered as an ecological microniche harboring diverse prokaryotic organisms. Possible benefits for persistence and transmission in nature for bacteria associated with protists are discussed. Our results support the hypothesis that ciliates attract potentially pathogenic bacteria and play the role of natural reservoirs for them.


All possible forms of coexistence of prokaryotes with metazoan organisms became one of the most rapidly developing research fields in microbiology, and also in physiology1,2. Protists may also host associated bacteria, but their microbiomes still are not investigated, and ciliates seem to be perfect candidates for this purpose. They are relatively big and bacterivorous, a number of representatives of this abundant phylum are easily recognizable and can be maintained in cultures. Ciliates have been intensively studied in wastewaters, focusing on their role as indicators of process efficiency3,4,5 and as detectors of heavy metal pollution6,7. Indeed, ciliates as filter-feeders are also efficient removers of some pathogens8,9,10,11, and contribute, together with disinfection systems, in pulling down the microbial load12,13, thus improving the quality of the effluent discharged by wastewater treatment plants14. At the same time, bacteria may benefit in finding protection inside the host cell from disinfection systems and chemical substances used to reduce bacterial load12,13,15. The phagotrophic activity of ciliates allows natural entrance of bacteria into the eukaryotic host, thus enabling the establishment of symbiotic associations with bacteria, also occasionally pathogenic ones16. Indeed, the ciliate cell offers a great variety of intracellular compartments suitable for bacterial colonization, and symbiotic bacteria have been described in the ciliates’ cytoplasm, nuclear apparatus, mitochondria and even in perinuclear space17.

Ciliates often host endosymbionts phylogenetically related to pathogenic bacteria belonging to the families Rickettsiaceae18,19,20 or Francisellaceae21,22. Symbiotic associations can be classified according to the stability of bacteria and ciliate interaction in three categories: permanent, highly infectious, and accidental17. The first two groups comprise all associations, which were recorded more than once in literature, and have been studied from ecological and evolutionary points of view. However, accidental invaders are never considered as true symbionts, such bacteria are likely poorly adapted for symbiotic persistence, and cannot be maintained for a long time within the host cell17. Such invaders usually have been reported just once17, and the study of these temporary occasional associations with “classical” microbiological and molecular techniques is rather difficult, as the associations are very unstable and promptly get lost. Several studies reported that in ciliates the establishment of symbiotic associations with pathogenic bacteria can be experimentally induced23. For example, ciliates under certain conditions may host Legionella pneumophila24,25, that sometimes leads to increase of virulence of these bacteria for human cells26. Internalized Listeria remain infectious in cysts of Tetrahymena27, and pathogenic Escherichia coli survives after passage through Tetrahymena cells28. However, such systems do not appear to be stable. The role of protists as environmental reservoirs of pathogenic bacteria has been elucidated more in amoebae, in which several pathogens have been found29,30, for example Francisella tularensis31,32, Vibrio cholerae33, E. coli29, Chlamydia34, Mycobacterium35,36,37, and Listeria monocytogenes38. Thus, in some cases, pathogenic bacteria also are a part of prokaryotic communities associated to protists, but for most of them the mode of maintenance and propagation in environment remains unclear.

However, contribution of bacteria in survival and environmental adaptation of protists, as well as the role of protists as reservoirs for the pools of microorganisms remains poorly understood. The studies of protistan microbiomes just start to be addressed with the diffusion of new technologies, such as Next Generation Sequencing (NGS). The characterization of the bacterial consortium associated with the Antarctic marine ciliate Euplotes focardii has been reported39. It was shown that even during long maintenance of ciliates in laboratory conditions they still keep a rather variegated set of initial bacterial cohabitants. Recently, Illumina sequencing was applied to discriminate between bacteria of the rumen fluid and symbiotic prokaryotes of ruminant ciliates40. The results of the pilot study showing that marine and freshwater ciliates harbour distinct microbial communities were reported in the latest publication41, and such prokaryotic communities were analyzed for several Paramecium samples42. However, up to now just two latter works remain the only studies of prokaryotic communities associated with single ciliate cells by NGS. This gap in knowledge extends also to other groups of protists, as there were only few attempts to assess the microbiomes of free-living amoebae43,44 applying NGS, and in several works bacterial diversity in association with protists was estimated by cloning and sequencing45,46,47,48,49. Also, few reports analyzed microbial consortia associated to cyanobacteria50,51,52 proving that even bacteria may organize and maintain stable communities of cohabiting prokaryotes. Nevertheless, all these few studies assume that unicellular organisms do have their own microbiomes.

Herein, we investigated the microbiomes of the sessile ciliates Stentor coeruleus (Fig. 1), isolated from a sewage stream, applying 16S rRNA gene metabarcoding approach on single ciliate cells. We propose to apply the term “core microbiome” for ciliates. Core microbiomes are defined as the “assemblages of microorganisms, active or inactive, associated with a certain habitat”53. Other authors suppose that core microbiome is the interacting subset of the total microbiome which fulfill a certain active function54. We suppose that bacteria composing core microbiome of a ciliate are prone to interact with eukaryotic cells. We also provide further support to the hypothesis that ciliates may host some opportunistic bacteria and should be considered as potential reservoirs of human pathogens.

Figure 1

In vivo morphology of Stentor coeruleus. Nikon Eclipse Ni, DIC microscopy. Scale bar = 50 µm.


A total number of 554260 reads were assembled, and after denoising and chimera filtering the obtained contigs clustered at 97% similarity threshold in 473 OTUs (for further information see Supplementary Table 1).

Average Shannon diversity index was calculated to assess diversity of the bacterial communities from the environment and those associated with the Stentor cells (Fig. 2). The environmental prokaryotic community was more than twice as much diverse as total prokaryotic community associated with the ciliates: average value of the environmental community richness index was 5.26 ± 0.30, while for the stentors it was 2.51 ± 0.99. Moreover, both rarefaction curves reached plateau indicating that OTUs forming respective communities were almost completely determined.

Figure 2

Rarefaction analysis of the studied samples. The average values of Shannon index are reported for the environmental samples (red) and for the Stentor cells (blue). The curves were generated basing on a 97% threshold level sequence similarity of OTUs.

In total representatives of 11 phyla were found, and among them 8 were abundant in all environment samples, while 6 phyla were the most plentiful in Stentor cells (Fig. 3). Bacterial community composition differed between the environments and the Stentor cells. Indeed, Firmicutes was the most abundant phylum in the environments (45.1%, average percentage of contigs), followed by Cyanobacteria (19.3%), Proteobacteria (14.6%), Actinobacteria (11.7%), Bacteroidetes (1.9%), Planctomycetes (1.7%), Chloroflexi (1.3%), and Euryarcheota (1.2%). On the contrary, the most abundant phylum in Stentor cells was Proteobacteria (66.7%), followed by less copious Bacteroidetes (15.6%), Firmicutes (6.9%), Cyanobacteria (5.3%), Fusobacteria (2.7%), and Actinobacteria (1.0%).

Figure 3

Composition of the prokaryotic communities from the environments and of Stentor coeruleus cells microbiomes. Relative abundances of the prokaryotic phyla are represented for each sample. Unclassified phyla are grouped in “Others”.

All OTUs obtained and assigned to certain bacteria were categorized according to the ecological niche as belonging to free-living bacteria (i.e. all those bacteria which live in the environment and are not known to form any kind of symbiotic relations), human commensals (i.e. bacteria normally associated to healthy human individuals), potential pathogens or bacteria related to pathogens (i.e. those bacteria causing diseases in humans or animals), and symbionts (i.e. known as obligate intracellular bacteria) (Table 1). The set of OTUs in the environmental samples was composed of a number of free-living genera, which are ubiquitous and normally present in the environment, such as bacteria from the order Rhizobiales (uncultured MNG7), uncultured Actinobacteria (bacterium PeM15), uncultured Cyanobacteria, uncultured Clostridiales, and some Firmicutes. Some potential pathogens and opportunistic bacteria like Streptococcus sp. were also detected in the environmental samples. Only few OTUs found in the environments were rather frequent also in the ciliate samples, though showing different average percentages (Table 1). The OTU belonging to uncultured Cyanobacteria, (very prevalent in Stentor4 cell), was identified as the chloroplast sequence of the microalga Mychonastes homosphaera (Chlorophyceae), which once has been found in symbiotic association with Stentor polymorphus55. In the same Stentor cell, OTU belonging to Holospora obtusa, a specific bacterial endosymbiont of the ciliate Paramecium caudatum, was found, but its abundance was very low (0.20%) (for further information, see Supplementary Table 1).

Table 1 The most common OTUs retrieved from the environmental samples and from Stentor cell samples.

Numerous OTUs belonging to some uncultured representatives of Neisseriaceae and to the genus Neisseria, normally associated to human mucosa, were recorded in association with stentors, and never in the environmental controls. Interestingly, Streptococcus sp. appeared to be mostly associated to Stentor cells, despite its presence was detected also in the environment. The non-parametric Kruskal-Wallis test assessed that there was a significant difference in the distribution of OTUs between stentors and their environments (p < 0.05), thus showing that human commensals and potentially pathogenic bacteria were indeed preferentially associated to Stentor cells. However, the most abundant OTU in six of seven Stentor samples belonged to the widespread endosymbiotic bacterium “Candidatus Megaira polyxenophila” (Alphaproteobacteria, Rickettsiales), probably inhabiting Stentor cells (Table 1). OTUs belonging to this endosymbiont were also present in the environmental samples, likely due to the entrapment of some Stentor cells during water filtering.

The microbial communities composition was reanalyzed after removing presumable symbionts OTUs, namely “Ca. Megaira polyxenophila” and the microalga chloroplast sequences, which reached 30–80% of the total number of reads in Stentor samples, thus, hiding the rest of bacterial diversity associated to ciliate cells. The total number of phyla did not change and corresponded to initial analysis, but their abundances shifted (Fig. 4). In the environment, Firmucutes (56.2%) remained the most abundant phylum, it was followed by Proteobacteria (15.8%), Actinobacteria (14.0%), Bacteroidetes (2.6%), Planctomycetes (2.2%), Chloroflexi (1.7%), and Cyanobacteria (1.7%). Removal of “Ca. Megaira polyxenophila” reads significantly reduced the fraction of Proteobacteria in the ciliates samples (32.0%), while the most abundant phyla associated with ciliates became Bacteroidetes (38.5%), then followed by Firmicutes (16.4%), Fusobacteria (6.3%), Actinobacteria (2.3%), and “Ca. Saccharibacteria” (2.1%), while Cyanobacteria (0.6%) became almost absent (Fig. 4).

Figure 4

Composition of the prokaryotic communities from the environments and of Stentor coeruleus cells microbiomes after removal of presumable abundant symbionts. Relative abundances of the prokaryotic phyla are represented for each sample. Unclassified phyla are grouped in “Others”.

After removing reads of the abundant symbionts, no new major taxa appeared in the environmental samples, while relative abundance of some OTUs increased significantly in the stentors samples (Table 2). The most common OTUs defined in the stentors samples belonged to the representatives of genera, which include human commensals or opportunistic pathogens.

Table 2 The most common OTUs retrieved from the environmental samples and from Stentor cell samples after removal of abundant symbionts’ OTUs.

Interestingly, a number of these bacteria were present, though in different percentages, in all seven or at least six Stentor samples (Table 2). We can assume that they form the core microbiome of S. coeruleus cells from the studied waterbody. This core microbiome included more than 10 bacterial genera, and just two of them, representatives of Chitinophagaceae and of “Ca. Saccharibacteria”, belong to the families considered as free-living. Capnocytophaga sp., Streptococcus sp., and representatives of Neisseriaceae, all known as opportunistic commensals of animals56,57,58,59,60, were predominant components of Stentor core microbiome. The ratios between members of core microbiome were different for all analyzed Stentor cells.

However, besides such widespread core microbiome components, some unique major bacteria were detected for some Stentor cells. For example, uncultured bacterium MNG7 was a major microbiome component of Stentor2 and Stentor6 specimens originating from two different sampling points. Xanthobacter sp. was the most abundant bacterium associated with Stentor4, and some bacteria from Leptotrichiaceae family were dominant in Stentor1 community. These bacteria, probably, can be considered as transitory components of Stentor microbiome.

In order to compare the prokaryotic community composition of the environments and ciliate cells, nmMDS was calculated on unweighted Unifrac distance matrix for all datasets (Fig. 5). The first dataset, comprising all symbiont contigs, showed a bright distinction between the environmental samples and the ciliate cells. Almost all stentors gathered together with the single exception of Stentor4, which located nearby other ciliates, but did not group with them, and was significantly distant from the environments (Fig. 5A). Indeed, this was the only Stentor specimen which had no “Ca. Megaira polyxenophila” endosymbionts, but contained symbiotic microalgae. After removal of symbionts OTUs, the environmental samples and the ciliates samples remained separated, and the ciliates still grouped together, though they became less congregated than before (Fig. 5B).

Figure 5

Results of Non-metric Multi-Dimensional Scaling based on the weighted Unifrac distance matrix. The original dataset (A), and the dataset after removal of the abundant symbionts (B).

Several analyses were carried out to test if the prokaryotic community composition of the environments and of the Stentor cells was statistically different. ANOSIM test showed that prokaryotic community compositions were significantly different between the environmental samples and the Stentor samples in all datasets (p < 0.05), accordingly with the results of NMDS.

PCoA was performed to determine whether prokaryotic communities of all samples were different among them, and across all datasets employed (Fig. 6). In the original dataset, all Stentor cells grouped together, and were separated from the environments (Fig. 6A). The other dataset, after removal of symbionts OTUs, showed that the ciliates microbiomes were in fact less homogeneous among themselves, but still reliably separated from environmental communities (Fig. 6B).

Figure 6

Results of Principal Component Analysis (PCoA). Percent of variation on the axis is indicated with PC. The original dataset (A), and the dataset after removal of the abundant symbionts (B).


In this study, the microbiomes of single cells of free-living ciliate Stentor coeruleus were investigated using 16S rRNA gene metabarcoding. Few studies addressed this topic in the last years using different techniques. Cloning of 16S rRNA gene PCR products obtained from single ciliate cells49,61 allowed to disclose a complex microbial community, but this approach is prone to underestimate the real biodiversity of the sampled microbiome. The complete metagenome of the bacterial consortium associated with a massive culture of the Antarctic ciliate Euplotes focardii was analyzed by NGS, which showed the interactions between microbiome and the host cell39. Very recently, microbiomes of rumen40 and free-living41,42 ciliates were investigated using single-cell 16S rRNA gene metabarcoding. With our study, we provide another example of how NGS technologies can be applied on single cells of cultivable and uncultivable protists to study their microbiomes and to investigate the presence of potential symbionts.

Prokaryotic community disclosed in the environmental samples was diverse and especially rich with several bacterial phyla, first of all Firmicutes, Actinobacteria and Proteobacteria. Many bacteria known to be anaerobic (uncultivable Clostridiales), or previously found in activated sludge (uncultured bacteria from “Ca. Saccharibacteria”62,63) (Tables 1, 2) were detected. The presence of anaerobic microorganisms is probably due to relatively high volume of the bottom sludge which has been collected together with water in the samples. Although some ciliate cells inevitably remained on filters, and their bacteria were counted as “environmental”, bacterial communities associated with Stentor cells were totally different from those of the environments. The only major OTUs in common between environment and stentors were the protist endosymbiont “Ca. Megaira polyxenophila”, Streptococcus sp. (significantly more numerous in stentors), and uncultured Rhizobiales MNG7, probably inhabiting the stream. Majority of environmental bacteria were absent or negligible in the communities associated with Stentor cells.

Different S. coeruleus cells isolated from the neighboring locations of the sewage stream were characterized by rather diverse associated bacterial communities, which included several dozens of different representatives (Supplementary Table 1). The composition of ciliates’ bacterial community looked apparently very similar at a first glance (Fig. 3, Table 1). Indeed, in six Stentor cells of seven, the presence of the bacterial endosymbiont “Ca. Megaira polyxenophila” was detected (Table 1). This obligatory intracellular bacterium was described in several unicellular organisms, mostly ciliates, and is phylogenetically closely related to the pathogen Rickettsia64. Unfortunately, we were not able to confirm directly “Ca. Megaira polyxenophila” presence inside the studied Stentor cells, as at the moment of OTUs classification the initial samples containing Stentor cells were lost. However, it looks very plausible that the ciliate specimens picked for our study were highly infected with these bacteria. The Stentor4 cell seemed to bear another peculiar endosymbiont, as numerous reads from the chloroplasts of the microalga Mychonastes homosphaera were detected in the dataset. This microalga species was documented in highly polluted wastewaters65, and was also found in symbiosis with another Stentor species, S. polymorphus55. Symbiotic Chlorella-like algae have been reported for S. coeruleus as well66. All selected Stentor cells contained blue-green pigment stentorin, specific only for S. coeruleus67, while presence or absence of symbiotic algae is not a reliable taxonomic character for the identification of Stentor species68. Some bacteria are known to have an algicidal effect, preventing algae survival and eventually killing them69, which could explain why the other six stentors heavily infected with “Ca. Megaira polyxenophila” did not contain microalgae. Still, it cannot be excluded that this particular Stentor cell belonged to another species. Anyway, removal of the chloroplast OTUs from the analysis revealed more uniform with other ciliates microbiome composition associated with Stentor4 cell.

Unexpectedly, few reads belonging to Holospora obtusa (Proteobacteria), bacterial intranuclear symbiont strictly host-specific for another ciliate, Paramecium caudatum, were detected in the dataset of the same Stentor cell (Supplementary Table 1). Outside of Paramecium, Holospora and related bacteria were reported in Frontonia70,71, but have never been recorded in Stentor72. Holospora possess an infectious stage in their life cycle, when they are released from the host cell to the environment and need to be ingested by another ciliate72. Probably, that Stentor cell might have engulfed H. obtusa infectious forms from the medium, and bacteria remained intact inside the ciliate up to DNA extraction. Of course, very low number of reads does not provide any basis to suggest that the symbiotic relationship emerged in this case. However, this finding gives interesting inkling to use 16S rRNA gene metabarcoding as molecular tool to investigate symbiont diversity in single cells of protists, and also to test early stages of symbiosis development when symbionts are just a few per host cell. On the other hand, the results of 16S rRNA gene metabarcoding analysis should be manually curated, and the best-matching sequences from the databases should be carefully checked, otherwise the presence of unknown symbiotic bacteria could be missed. Even known true symbionts might be misidentified, as it happened in our case with “Ca. Megaira polyxenophila”, which was misclassified initially with Ribosomal Database Project (RDP) against SILVA database as “uncultured Rickettsiaceae”.

After removal of reads of the most abundant symbionts, other OTUs became major or considerable enough to conclude that they reflect bacteria preferentially associated with ciliates. Many OTUs related to commensals and potential pathogens displayed high relative abundances within Stentor samples (Table 2). Bacteria discovered in all Stentor samples, according to the concept of core microbiome53, were supposed to form a core microbiome of S. coeruleus. Capnocytophaga sp., Streptococcus sp., some Neisseriaceae and Porphyromonas sp. were the most abundant, and, perhaps, they could be considered as keystone components of S. coeruleus core microbiome. These genera together with other members of Stentor core microbiome (e.g., Veillonella sp., Rothia sp., Lautropia sp., Prevotella sp., Bergeyella sp., and Haemophilus sp.) are a part of human microbiome73. Majority of them are known to colonize the mucosal surfaces of mammals, and are considered as opportunistic bacteria, if not true pathogens, which may cause serious diseases under certain conditions56,57,58,59,60. The interaction between Streptococcus sp. and Veillonella sp. was studied during the formation of biofilms74, which could be interesting since they have been both found in association with Stentor cells and might have a role in bacterial aggregation. The genus Neisseria and other representatives of the family Neisseriaceae were found in rather high abundancy (Table 2). Neisseria species normally colonize the mucosal surface of mammals and rarely invade their host cells56,75, so their presence in association with stentors might reveal novel aspects of this bacterial genus biology. Importantly, all these bacteria were absent or, like Streptococcus sp., present in much lower amounts in the environment.

Some free-living bacteria, like a representative of Chitinophagaceae or “Ca. Saccharibacteria” described in wastewater treatment plants76, were also retrieved in our analysis as a part of Stentor core microbiome (Table 2). Numerous OTUs associated with bacteria occasionally appearing as dominant microbiome components were defined in several Stentor samples. These bacteria belong either to presumably free-living genera (like bacterium MNG7 or Xanthobacter sp.), or to the groups of commensals (Leptotrichiaceae). We suggest to consider such bacteria as transitory components of Stentor microbiome, which by chance manage to colonize some ciliate cells.

We do not have an idea of any microbiome function useful for the host ciliate. Still, there is a common feature for almost all major bacteria in Stentor microbiome. These bacteria are very likely prone to interact with eukaryots, as they belong to genera encompassing numerous commensals or even pathogens of animals. When they appear freely in water, they may use their general skills to interact with available eukaryotic cells, namely, protists. There are two possible ways of bacterial persistence in association with protists: to stay attached on the surface of the host cell, or to resist digestion or even to escape from the food vacuoles to the host cytoplasm. Ciliates are rather big protists (S. coeruleus reaches up to 0.5–1 mm in length, Fig. 1), and their large cell surface, densely covered with cilia, may act as a kind of substrate for bacterial attachment, thus, facilitating them to propagate in a more stable environment. Bacteria propagating on ciliate cell surface have been described77,78. Sessile ciliates may colonize big areas of suitable substrates, and in this way bacterial community associated with ciliate cells may get stabilized in certain general environmental conditions. At the same time, ciliates can swim quickly and use different taxis allowing them to select favorable conditions79, which would be beneficial also for the associated bacteria. Ciliate surface may also offer a shelter to bacteria, protecting them from phagocytosis by other microorganisms. It is known that bacteria have developed numerous strategies to avoid prey-selective grazing by protozoa, namely morphological changes, high speed motility, and production of antagonistic or toxic substances80,81,82,83. One of the most important defense mechanisms is formation of biofilms or aggregations, in which separate bacteria are less vulnerable for predators81. From this point of view, the ciliate cell surface allows bacteria to adhere, aggregate, form biofilms and co-regulate their living activities through a quorum sensing mechanism, which regulates gene expression as a response to fluctuations in bacterial population density84. Thus, even a small substrate may be substantial for propagation of certain bacteria in the environment. To our best knowledge, no one have tried yet to analyze biofilms formation and bacterial quorum sensing mechanisms of regulation at such small-scale level as a surface of unicellular eukaryotes.

The second possibility for bacteria to stay in association with a ciliate is to somehow escape or survive digestion after being phagocytized. It is well known that many pathogenic bacteria enter the host cells by phagocytosis85. Some opportunistic bacteria, if engulfed by a phagocyte, are also able to avoid digestion. This may allow them to find a shelter from predators or from viral lysis inside the protozoan hosts, and to persist there as occasional “endosymbionts”29. For some bacteria, intracellular persistence may even trigger and increase their pathogenic properties and virulence86. Thus, protists may serve as transient reservoirs for many potentially dangerous bacteria. It has been proven that amoebae are the “Trojan horses” of the microbial world, as they act as temporary hosts for many pathogens29. At the same time, the similar role of ciliates has been also shown, but small-scale investigations were focused mainly on pathogens found as endosymbionts of ciliates15,25. In some recent works49,61 diverse digestion-resistant bacteria were described in different ciliates; some of them were enough numerous to be visualized inside the host cells with fluorescence in situ hybridization. However, the abundance of different bacteria forming the microbiome of ciliates is usually low, thus making NGS analysis a much more sensitive technique to study these phenomena.

Together with recently published data41,42 our results allow to presume that free-living ciliates may serve as certain “magnets” accumulating from the environment bacteria searching for hosts, and providing an ecological microniche for them. The ciliate core microbiomes then would be mostly formed by such bacteria, and the ratio between these would be dynamic and varying from cell to cell; sometimes also occasional bacteria could dominate, forming a transitory part of a microbiome. Actually this is what we obtained as a result of our metabarcoding analysis. Thus, ciliates can play in nature the role of reservoirs for potentially opportunistic and pathogenic bacteria.

Statistical analyses showed that sets of OTUs from the environmental controls were significantly different from those associated to stentors (Figs 5, 6). Indeed, statistical analyses, nmMDS and PCoA strongly confirmed that prokaryotic communities of Stentor cells and corresponding environments were clearly separated from each other, being very distant (Figs 5, 6). However, ciliate cells did not display a homogenous microbiome among themselves. When reads of abundant symbionts were removed from the analyses, it became clear that the microbiomes of single cells had some individual differences. At the same time, our data are insufficient to generalize if all specimens of the same ciliate species isolated from one locality possess similar or diverse microbiomes, but allow to suggest that there is a core microbiome for ciliates of the same species, at least isolated from the same locality. Further studies using several species of ciliates from the same location and from different origins should be performed to clarify this aspect. We still cannot rule out the hypothesis that environment determines the ciliates’ microbiomes, as all bacteria attracted and accumulated by ciliates, probably, come from their environment. However, we suppose that environment does not shape the core microbiomes of ciliates, as the latter in the end have almost nothing in common with the environmental prokaryotic community.


Sample collection and preparation

Three samples were collected from a sewage stream located in Peterhof, St. Petersburg, Russia (59.879780, 29.864358). The sampling points were located about 10 m downstream one after another, starting with PW1.

Each collected sample (200 ml) was divided in two subsamples: 150 ml were immediately filtered through 0.2 μm nitrocellulose filters (Sartorius, Germany) for further NGS analysis of the environmental controls, and 50 ml were brought to the laboratory within 15 min for isolation of ciliates. The filters were air-dried and stored at −80 °C till DNA extraction.

Stentor cells were observed in vivo, and identified as S. coeruleus (Fig. 1) by presence of blue-green pigment and moniliform macronucleus67. Three ciliate specimens were isolated as replicates for each sampling point with a sterile micropipette, washed thoroughly through several passages in sterile Volvic water and then kept in the last water aliquot overnight to reduce the load of random bacteria present in food vacuoles. Then stentors were washed briefly in autoclaved sterile distilled water, in order to reduce contaminants and microorganisms attached to the cell surface. Finally, single cells were fixed separately in 50 µl of 70% ethanol for further DNA extraction.

DNA extraction, libraries preparation and sequencing

The filters were firstly treated by sonication as described87, then total genomic DNA was washed off and extracted using NucleoSpin® Tissue kit (Macherey-Nagel, Germany).

Just before DNA extraction, the Stentor samples were centrifuged for 30 min at 14000 rpm and 4 °C, and the pellet was dried for 3–5 minutes in the same PCR tube. Then 15 μl of MilliQ water and a mixture of glass beads (0.1 and 0.5 mm in diameter) were added to the tubes in approximate ratio 3:1. The mixture was homogenized with Tissue Lyser LT (QIAGEN, Germany) for 3 minutes at a maximal frequency of 50 Hz. The suspension was vortexed and centrifuged at 14000 rpm for 10 min. The lysate was transferred to a new tube, the glass beads were washed in 15 μl of MilliQ water, centrifuged again, and the supernatant was transferred in the tube with the lysate.

Preparation of the DNA libraries was performed according to the Illumina protocol (Part # 15044223, Rev. B.) using primers for V3 and V4 region of the 16S rRNA gene88. Two of nine Stentor samples did not yield sufficient DNA for library preparation and were discarded. The DNA libraries were sequenced on the MiSeq platform (Illumina, USA).

Preparation of the DNA libraries and sequencing was carried out in the Center of Shared Scientific Equipment “Persistence of microorganisms” at the Institute for Cellular and Intracellular Symbiosis, UrB RAS (Orenburg, Russia).

Sequencing data analysis

Raw FASTQ files were analyzed using the Quantitative Insights Into Microbial Ecology 1.9.1 software package (QIIME)89. De-multiplexing and quality filtering were performed removing any low quality or ambiguous reads, and sequences shorter than 200 nucleotides were discarded. Chimeras were identified using UCHIME90 and subsequently removed from the analysis. Operational Taxonomic Units (OTUs) were clustered with a 97% of similarity cutoff using USEARCH91, and SILVA 119 database92 as reference. The most common sequence was selected in each OTU as representative. Taxonomic classification up to genus level was performed using Ribosomal Database Project Classifier (RDP)93 against the SILVA 119 database.

Prokaryotic community analysis

QIIME was used to evaluate diversity of the prokaryotic communities both within and between samples. Firstly, the relative abundances of prokaryotic taxa were estimated as the percentage of contigs number for each taxon for the environmental and the Stentor cell samples, and the community richness was evaluated calculating the Shannon index. The significance of OTUs presence in the ciliates samples was assessed using the non-parametric Kruskal-Wallis test. Furthermore, similarities between stentors and their environments were assessed using the non-metric Multi-Dimensional Scaling (nmMDS) and Principal Coordinate Analysis (PCoA) based on the unweighted Unifrac distance matrices. In addition, beta diversity was estimated to compare the bacterial communities of the Stentor cells and their environments. To confirm the significance of differences observed between bacterial communities of the environments and stentors, ANOSIM was calculated. The same analyses were carried out also after excluding the most abundant OTUs representing symbiotic microorganisms associated with stentors (see Results).

Data Availability

The raw reads generated and analyzed during the current study are available in the ENA database (study number PRJEB30974). The datasets analyzed during this study are included in the Supplementary Information file.


  1. 1.

    Knight, R. et al. Best practices for analysing microbiomes. Nat. Rev. Microbiol. 16, 410–422 (2018).

  2. 2.

    Ost, K. S. & Round, J. L. Communication between the microbiota and mammalian immunity. Annu. Rev. Microbiol. 72, 399–422 (2018).

  3. 3.

    Madoni, P. Protozoa as indicators of wastewater treatment efficiency in The Handbook of Water and Wastewater Microbiology 361–371 (Academic Press, Amsterdam, 2003).

  4. 4.

    Papadimitriou, C. A. et al. Investigation of protozoa as indicators of wastewater treatment efficiency in constructed wetlands. Desalination 250, 378–382 (2010).

  5. 5.

    Debastiani, C., Meira, B. R., Lansac-Tôha, F. M., Velho, L. F. M. & Lansac-Tôha, F. A. Protozoa ciliates community structure in urban streams and their environmental use as indicators. Braz. J. Biol. 76, 1043–1053 (2016).

  6. 6.

    Martín-González, A., Díaz, S., Borniquel, S., Gallego, A. & Gutiérrez, J. C. Cytotoxicity and bioaccumulation of heavy metals by ciliated protozoa isolated from urban wastewater treatment plants. Res. Microbiol. 157, 108–118 (2006).

  7. 7.

    Gomiero, A., Dagnino, A., Nasci, C. & Viarengo, A. The use of protozoa in ecotoxicology: application of multiple endpoint tests of the ciliate E. crassus for the evaluation of sediment quality in coastal marine ecosystems. Sci. Total Environ. 442, 534–544 (2013).

  8. 8.

    Curds, C. R. & Fey, G. J. The effect of ciliated protozoa on the fate of Escherichia coli in the activated-sludge process. Water Res. 3, 853–867 (1969).

  9. 9.

    Stott, R., May, E., Matsushita, E. & Warren, A. Protozoan predation as a mechanism for the removal of Cryptosporidium oocysts from wastewaters in constructed wetlands. Water Sci. Technol. 44, 191–198 (2001).

  10. 10.

    Ravva, S. V., Sarreal, C. Z. & Mandrell, R. E. Identification of protozoa in dairy lagoon wastewater that consume Escherichia coli O157: H7 preferentially. PLoS ONE 5, e15671 (2010).

  11. 11.

    Siqueira-Castro, I. C. V., Greinert-Goulart, J. A., Bonatti, T. R., Yamashiro, S. & Franco, R. M. B. First report of predation of Giardia sp. cysts by ciliated protozoa and confirmation of predation of Cryptosporidium spp. oocysts by ciliate species. Environ. Sci. Pollut. Res. Int. 23, 11357–11362 (2016).

  12. 12.

    King, C. H., Shotts, E. B., Wooley, R. E. & Porter, K. G. Survival of coliforms and bacterial pathogens within protozoa during chlorination. Appl. Environ. Microbiol. 54, 3023–3033 (1988).

  13. 13.

    Barker, J. & Brown, M. R. W. Trojan horses of the microbial world: protozoa and the survival of bacterial pathogens in the environment. Microbiology 140, 1253–1259 (1994).

  14. 14.

    Salvadò, H., Gracia, M. P. & Amigó, J. M. Capability of ciliated protozoa as indicators of effluent quality in activated sludge plants. Water Res. 29, 1041–1050 (1995).

  15. 15.

    Fields, B. S. The molecular ecology of legionellae. Trends Microbiol. 4, 286–290 (1996).

  16. 16.

    Görtz, H. D. & Brigge, T. Intracellular bacteria in protozoa. Naturwissenschaften 85, 359–368 (1998).

  17. 17.

    Fokin, S. I. Bacterial endocytobionts of ciliophora and their interactions with the host cell. Int. Rev. Cytol. 236, 181–249 (2004).

  18. 18.

    Schrallhammer, M. et al. ‘Candidatus Megaira polyxenophila’ gen. nov., sp. nov.: considerations on evolutionary history, host range and shift of early divergent rickettsiae. PLoS ONE 8, e72581 (2013).

  19. 19.

    Sabaneyeva, E. et al. Host and symbiont intraspecific variability: The case of Paramecium calkinsi and “Candidatus Trichorickettsia mobilis”. Eur. J. Protistol. 62, 79–94 (2018).

  20. 20.

    Castelli, M. et al. The hidden world of Rickettsiales symbionts: “Candidatus Spectririckettsia obscura,” a novel bacterium found in Brazilian and Indian Paramecium caudatum. Microb. Ecol. 77, 748–758 (2019).

  21. 21.

    Schrallhammer, M., Schweikert, M., Vallesi, A., Verni, F. & Petroni, G. Detection of a novel subspecies of Francisella noatunensis as endosymbiont of the ciliate Euplotes raikovi. Microb. Ecol. 61, 455–464 (2011).

  22. 22.

    Vallesi, A. et al. A new species of the γ-Proteobacterium Francisella, F. adeliensis sp. nov., endocytobiont in an Antarctic marine ciliate and potential evolutionary forerunner of pathogenic species. Microb. Ecol. 77, 587–596 (2019).

  23. 23.

    Fields, B. S., Shotts, E. B., Feeley, J. C., Gorman, G. W. & Martin, W. T. Proliferation of Legionella pneumophila as an intracellular parasite of the ciliated protozoan Tetrahymena pyriformis. Appl. Environ. Microbiol. 47, 467–471 (1984).

  24. 24.

    Berk, S. G. & Garduño, R. A. The Tetrahymena and Acanthamoeba model systems. Methods Mol. Biol. 954, 393–416 (2013).

  25. 25.

    Watanabe, K. et al. Ciliate Paramecium is a natural reservoir of Legionella pneumophila. Sci. Rep. 6 (2016).

  26. 26.

    Koubar, M., Rodier, M. H., Garduño, R. A. & Frère, J. Passage through Tetrahymena tropicalis enhances the resistance to stress and the infectivity of Legionella pneumophila. FEMS Microbiol. Lett. 325, 10–15 (2011).

  27. 27.

    Pushkareva, V. I. & Ermolaeva, S. A. Listeria monocytogenes virulence factor Listeriolysin O favors bacterial growth in co-culture with the ciliate Tetrahymena pyriformis, causes protozoan encystment and promotes bacterial survival inside cysts. BMC microbiology 10, 26 (2010).

  28. 28.

    Smith, C. D., Berk, S. G., Brandl, M. T. & Riley, L. W. Survival characteristics of diarrheagenic Escherichia coli pathotypes and Helicobacter pylori during passage through the free-living ciliate, Tetrahymena sp. FEMS Microbiol. Ecol. 82, 574–583 (2012).

  29. 29.

    Molmeret, M., Horn, M., Wagner, M., Santic, M. & Kwaik, Y. A. Amoebae as training grounds for intracellular bacterial pathogens. Appl. Environ. Microbiol. 71, 20–28 (2005).

  30. 30.

    Corsaro, D., Pages, G. S., Catalan, V., Loret, J. F. & Greub, G. Biodiversity of amoebae and amoeba-associated bacteria in water treatment plants. Int. J. Hyg. Env. Health 213, 158–166 (2010).

  31. 31.

    Kantardjiev, T. & Velinov, T. Z. Interaction between protozoa and microorganisms of the genus Francisella. Problems of Infectious Diseases 22, 34–35 (1995).

  32. 32.

    Abd, H., Johansson, T., Golovliov, I., Sandström, G. & Forsman, M. Survival and growth of Francisella tularensis in Acanthamoeba castellanii. Appl. Environ. Microbiol. 69, 600–606 (2003).

  33. 33.

    Thom, S., Warhurst, D. & Drasar, B. S. Association of Vibrio cholerae with fresh water amoebae. J. Med. Microbiol. 36, 303–306 (1992).

  34. 34.

    Birtles, R. J., Rowbotham, T. J., Storey, C., Marrie, T. J. & Raoult, D. Chlamydia-like obligate parasite of free-living amoebae. Lancet 349, 925–926 (1997).

  35. 35.

    Mura, M. et al. Replication and long-term persistence of bovine and human strains of Mycobacterium avium subsp. paratuberculosis within Acanthamoeba polyphaga. Appl. Environ. Microbiol. 72, 854–859 (2006).

  36. 36.

    Yu, H. S. et al. Natural occurrence of Mycobacterium as an endosymbiont of Acanthamoeba isolated from a contact lens storage case. Korean J. Parasitol. 45, 11 (2007).

  37. 37.

    Thomas, V., Loret, J. F., Jousset, M. & Greub, G. Biodiversity of amoebae and amoebae‐resisting bacteria in a drinking water treatment plant. Environ. Microbiol. 10, 2728–2745 (2008).

  38. 38.

    Ly, T. M. C. & Müller, H. E. Ingested Listeria monocytogenes survive and multiply in protozoa. J. Med. Microbiol. 33, 51–54 (1990).

  39. 39.

    Pucciarelli, S. et al. Microbial consortium associated with the Antarctic marine ciliate Euplotes focardii. Microb. Ecol. 70, 484–497 (2015).

  40. 40.

    Park, T. & Yu, Z. Do ruminal ciliates select their preys and prokaryotic symbionts? Front. Microbiol. 9, 1710 (2018).

  41. 41.

    Rossi, A., Bellone, A., Fokin, S. I., Boscaro, V. & Vannini, C. Detecting associations between ciliated protists and prokaryotes with culture-independent single-cell microbiomics: a proof-of-concept study. Microb. Ecol. 78, 232–242 (2019).

  42. 42.

    Plotnikov, A. O. et al. High-throughput sequencing of the 16S rRNA gene as a survey to analyze the microbiomes of free-living ciliates Paramecium. Microb. Ecol. 78, 286–298 (2019).

  43. 43.

    Delafont, V., Brouke, A., Bouchon, D., Moulin, L. & Héchard, Y. Microbiome of free-living amoebae isolated from drinking water. Water Res. 47, 6958–6965 (2013).

  44. 44.

    Delafont, V., Bouchon, D., Héchard, Y. & Moulin, L. Environmental factors shaping cultured free-living amoebae and their associated bacterial community within drinking water network. Water Res. 100, 382–392 (2016).

  45. 45.

    Yoon, H. S. et al. Single-cell genomics reveals organismal interactions in uncultivated marine protists. Science 332, 714–717 (2011).

  46. 46.

    Martinez-Garcia, M. et al. Unveiling in situ interactions between marine protists and bacteria through single cell sequencing. ISME J. 6, 703 (2012).

  47. 47.

    Kuo, R. C. & Lin, S. Ectobiotic and endobiotic bacteria associated with Eutreptiella sp. isolated from Long Island Sound. Protist 164, 60–74 (2013).

  48. 48.

    Brock, D. A., Read, S., Bozhchenko, A., Queller, D. C. & Strassmann, J. E. Social amoeba farmers carry defensive symbionts to protect and privatize their crops. Nat. Comm. 4, 2385 (2013).

  49. 49.

    Gong, J. et al. Protist-bacteria associations: Gammaproteobacteria and Alphaproteobacteria are prevalent as digestion-resistant bacteria in ciliated protozoa. Front. Microbiol. 7, 498 (2016).

  50. 50.

    Velichko, N. et al. Consortium of the ‘bichlorophyllous’ cyanobacterium Prochlorothrix hollandica and chemoheterotrophic partner bacteria: culture and metagenome‐based description. Environ. Microbiol. Rep. 7, 623–633 (2015).

  51. 51.

    Callieri, C. et al. The microbiome associated with two Synechococcus ribotypes at different levels of ecological interaction. J. Phycol. 53, 1151–1158 (2017).

  52. 52.

    Alvarenga, D. O., Fiore, M. F. & Varani, A. M. A metagenomic approach to cyanobacterial genomics. Front. Microbiol. 8, 809 (2017).

  53. 53.

    Shade, A. & Handelsman, J. Beyond the Venn diagram: the hunt for a core microbiome. Environ. Microbiol. 14, 4–12 (2012).

  54. 54.

    Hernandez-Agreda, A., Gates, R. D. & Ainsworth, T. D. Defining the core microbiome in corals’ microbial soup. Trends Microbiol. 25, 125–140 (2017).

  55. 55.

    Hoshina, R. et al. Cytological, genetic, and biochemical characteristics of an unusual non‐Chlorella photobiont of Stentor polymorphus collected from an artificial pond close to the shore of Lake Biwa, Japan. Phycol. Res. 61, 7–14 (2013).

  56. 56.

    Nikulin, J., Panzner, U., Frosch, M. & Schubert-Unkmeir, A. Intracellular survival and replication of Neisseria meningitidis in human brain microvascular endothelial cells. Int. J. Med. Microbiol 296, 553–558 (2006).

  57. 57.

    Wang, K. et al. Preliminary analysis of salivary microbiome and their potential roles in oral lichen planus. Sci. Rep. 6, 22943 (2016).

  58. 58.

    Lombardo Bedran, T. B. et al. Porphyromonas endodontalis in chronic periodontitis: a clinical and microbiological cross-sectional study. J. Oral Microbiol. 4, 10123 (2012).

  59. 59.

    Ma, A. & Goetz, M. B. Capnocytophaga canimorsus sepsis with associated thrombotic thrombocytopenic purpura. Am. J. Med. Sci. 345, 78–80 (2013).

  60. 60.

    Cadre, B., Al Oraimi, M., Grinholtz‐Haddad, J. & Benkhatar, H. “My dog deafened me!”: case report of Capnocytophaga canimorsus infection and literature review. Laryngoscope 129, E41–E43 (2018).

  61. 61.

    Omar, A., Zhang, Q., Zou, S. & Gong, J. Morphology and phylogeny of the soil ciliate Metopus yantaiensis n. sp. (Ciliophora, Metopida), with identification of the intracellular bacteria. J. Eukaryot. Microbiol. 64, 792–805 (2017).

  62. 62.

    Guo, F. & Zhang, T. Profiling bulking and foaming bacteria in activated sludge by high throughput sequencing. Water Res. 46, 2772–2782 (2012).

  63. 63.

    Hugenholtz, P., Tyson, G. W., Webb, R. I., Wagner, A. M. & Blackall, L. L. Investigation of candidate division TM7, a recently recognized major lineage of the domain Bacteria with no known pure-culture representatives. Appl. Environ. Microbiol. 67, 411–419 (2001).

  64. 64.

    Lanzoni, O. et al. Diversity and environmental distribution of the cosmopolitan endosymbiont “Candidatus Megaira”. Sci. Rep. 9, 1179 (2019).

  65. 65.

    Deng, D. & Tam, N. F. Y. Isolation of microalgae tolerant to polybrominated diphenyl ethers (PBDEs) from wastewater treatment plants and their removal ability. Bioresour. Technol. 177, 289–297 (2015).

  66. 66.

    Fernández-Leborans, G. & Zaldumbide, M. C. Chlorella sp. (Chlorophyta) endosymbiotic of Stentor coeruleus (Protozoa, Ciliophora). Arch. Protistenk. 127, 193–200 (1983).

  67. 67.

    Foissner, W. & Wölfl, S. Revision of the genus Stentor Oken (Protozoa, Ciliophora) and description of S. araucanus nov. spec. from South American lakes. J. Plankton Res. 16, 255–289 (1994).

  68. 68.

    Fernandes, N. M., da Silva Neto, I. D. & Schrago, C. G. Morphology and phylogenetic position of an unusual Stentor polymorphus (Ciliophora: Heterotrichea) without symbiotic algae. J. Eukaryot. Microbiol. 61, 305–312 (2014).

  69. 69.

    Mayali, X. Metabolic interactions between bacteria and phytoplankton. Front. Microbiol. 9, 727 (2018).

  70. 70.

    Lanzoni, O. et al. Rare freshwater ciliate Paramecium chlorelligerum Kahl, 1935 and its macronuclear symbiotic bacterium “Candidatus Holospora parva”. PloS ONE 11, e0167928 (2016).

  71. 71.

    Fokin, S. I., Serra, V., Ferrantini, F., Modeo, L. & Petroni, G. “Candidatus Hafkinia simulans” gen. nov., sp. nov., a novel Holospora-like bacterium from the macronucleus of the rare brackish water ciliate Frontonia salmastra (Oligohymenophorea, Ciliophora): multidisciplinary characterization of the new endosymbiont and its host. Microb. Ecol. 77, 1092–1106 (2019).

  72. 72.

    Fokin, S. I. & Görtz, H. D. Diversity of Holospora bacteria in Paramecium and their characterization. In Endosymbionts in Paramecium (ed. Fujishima, M.) 161–199 (Berlin, Heidelberg: Springer 2009).

  73. 73.

    Lloyd-Price, J. et al. Strains, functions and dynamics in the expanded Human Microbiome Project. Nature 550, 61–66 (2017).

  74. 74.

    Mashima I, Nakazawa F. The interaction between Streptococcus spp. and Veillonella tobetsuensis in the early stages of oral biofilm formation. J. Bacteriol. JB–02512 (2015).

  75. 75.

    Spinosa, M. R. et al. The Neisseria meningitidis capsule is important for intracellular survival in human cells. Infect. Immun. 75, 3594–3603 (2007).

  76. 76.

    Kindaichi, T. et al. Phylogenetic diversity and ecophysiology of Candidate phylum Saccharibacteria in activated sludge. FEMS Microbiol. Ecol. 92, fiw078 (2016).

  77. 77.

    Radek, R. Adhesion of bacteria to protists. In Prokaryotic Cell Wall Compounds (eds König, H. et al.), 429–456 (Berlin, Heidelberg: Springer 2010).

  78. 78.

    Castelli, M. et al. Deianiraea, an extracellular bacterium associated with the ciliate Paramecium, suggests an alternative scenario for the evolution of Rickettsiales. ISME J, (2019).

  79. 79.

    Gong, J., Song, W. & Warren, A. Periphytic ciliate colonization: annual cycle and responses to environmental conditions. Aquatic Microb. Ecol. 39, 159–170 (2005).

  80. 80.

    Hahn, M. W. & Höfle, M. G. Grazing of protozoa and its effect on populations of aquatic bacteria. FEMS Microbiol. Ecol. 35, 113–121 (2001).

  81. 81.

    Matz, C. & Kjelleberg, S. Off the hook–how bacteria survive protozoan grazing. Trends Microbiol. 13, 302–307 (2005).

  82. 82.

    Pernthaler, J. Predation on prokaryotes in the water column and its ecological implications. Nat. Rev. Microbiol. 3, 537–546 (2005).

  83. 83.

    Jousset, A. Ecological and evolutive implications of bacterial defences against predators. Environ. Microbiol. 14, 1830–1843 (2012).

  84. 84.

    Miller, M. B. & Bassler, B. L. Quorum sensing in bacteria. Annu. Rev. Microbiol. 55, 165–199 (2001).

  85. 85.

    Ribet, D. & Cossart, P. How bacterial pathogens colonize their hosts and invade deeper tissues. Microb. Infect. 17, 173–183 (2015).

  86. 86.

    Cirillo, J. D., Falkow, S. & Tompkins, L. S. Growth of Legionella pneumophila in Acanthamoeba castellanii enhances invasion. Infect. Immun. 62, 3254–3261 (1994).

  87. 87.

    Kesberg, A. I. & Schleheck, D. Improved protocol for recovery of bacterial DNA from water filters: sonication and backflushing of commercial syringe filters. J. Microbiol. Methods 93, 55–57 (2013).

  88. 88.

    Klindworth, A. et al. Evaluation of general 16S ribosomal RNA gene PCR primers for classical and next-generation sequencing-based diversity studies. Nucl. Acids Res. 41, e1–e1 (2013).

  89. 89.

    Caporaso, J. G. et al. QIIME allows analysis of high-throughput community sequencing data. Nat. Methods 7, 335–336 (2010).

  90. 90.

    Edgar, R. C., Haas, B. J., Clemente, J. C., Quince, C. & Knight, R. UCHIME improves sensitivity and speed of chimera detection. Bioinformatics 27, 2194–2200 (2011).

  91. 91.

    Edgar, R. C. Search and clustering orders of magnitude faster than BLAST. Bioinformatics 26, 2460–2461 (2010).

  92. 92.

    Quast, C. et al. The SILVA ribosomal RNA gene database project: improved data processing and web-based tools. Nucl. Acids Res. gks1219 (2013).

  93. 93.

    Wang, Q., Garrity, G. M., Tiedje, J. M. & Cole, J. R. Naive Bayesian classifier for rapid assignment of rRNA sequences into the new bacterial taxonomy. Appl. Environ. Microbiol. 73, 5261–5267 (2007).

Download references


Research was supported by RSF grant 16-14-10157 to AL.P. Travel grant 28875817 of St Petersburg State University supported mobility to AL.P and European Commission FP7-PEOPLE-2009-IRSES project CARBALA (295176) supported mobility to O.L., AN.P. Experiments were partly performed using the equipment of the Core Facility Center “Cultivation of Microorganisms” of St. Petersburg State University, Saint Petersburg, Russia.

Author information

Conceived and designed the experiments: O.L. AL.P. Field work: O.L. AL.P. Prepared samples for NGS. O.L. AL.P. NGS sequencing: AN.P. Bioinformatic analysis: O.L. Y.K. Analyzed data: O.L. AL.P. Contributed reagents/materials/analysis tools: AL.P. Mobility support and international network organization: G.M., G.P. Wrote the paper: O.L., AL.P. with contributions from AN.P. and G.P. All authors revised the final version of the manuscript.

Correspondence to Alexey Potekhin.

Ethics declarations

Competing Interests

The authors declare no competing interests.

Additional information

Publisher’s note: Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Supplementary information

Rights and permissions

Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The images or other third party material in this article are included in the article’s Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this license, visit

Reprints and Permissions

About this article

Verify currency and authenticity via CrossMark

Cite this article

Lanzoni, O., Plotnikov, A., Khlopko, Y. et al. The core microbiome of sessile ciliate Stentor coeruleus is not shaped by the environment. Sci Rep 9, 11356 (2019) doi:10.1038/s41598-019-47701-8

Download citation


By submitting a comment you agree to abide by our Terms and Community Guidelines. If you find something abusive or that does not comply with our terms or guidelines please flag it as inappropriate.