An integrative approach sheds new light onto the systematics and ecology of the widespread ciliate genus Coleps (Ciliophora, Prostomatea)

Morphology and phenotypic plasticity

The morphological features of the investigated colepid strains differed from those described for C. hirtus, C. spetai and even for N. nolandi¹ (Fig. 1). Characteristics that matched the descriptions were the ciliate cell length and width, the barrel-shaped cell (except for strain CIL-2017/7, which was pear-shaped and strain CCAP 1613/15 that had a cylindrical shape), a number of six armor tiers, the structure of the armor tiers (hirtus-type or nolandia-type, respectively), and one caudal cilium (Table S2). Variations (CV > 20%) were found (i) in the number of plate windows in the posterior/anterior main plates even within individual cells, and (ii) in the presence/absence of anterior and posterior spines (Tables 1 + S2, Fig. 1). This phenotypic plasticity of the ciliate could also be observed in freshly collected Coleps specimens and was therefore not an artifact resulting from cultivation conditions (Fig. 1A). Wickham and Gugenberger⁴³ hypothesized that the formation of the spines was a response to grazing pressure on C. hirtus; however, this could not be confirmed by respective experiments. Nevertheless, spineless specimens of C. hirtus have obviously been found before^44,45,46,47. Luckily, we were able to investigate two strains (CCAP 1613/1 and CCAP 1613/2) that had been kept in the CCAP culture collection since the 1950ies and the 1960ies and which did not bear any spines or symbionts and could be clearly assigned to C. hirtus (Fig. 1T–V). These observations suggest that without predation pressure, colepid ciliates probably do not need to synthesize spines avoiding ingestion by a predator.

The presence/absence of green algal endosymbionts, one of the diagnostic features for the discrimination among C. hirtus subspecies and C. spetai, was also not a stable feature (Table S2). Under culture conditions, some strains lost their endosymbionts completely, other strains consisted of symbiotic and aposymbiotic individuals, and some strains showed only symbiont-bearing individuals (e.g., CCAP 1613/5 and CIL-2017/6). This indicates that the symbiosis is facultative and might be probably influenced by cultivation or environmental conditions (presumably, though not tested, food availability). Consequently, the morphological separation of C. hirtus into the two subspecies may no longer be valid. We clearly demonstrated that the morphological features used for species descriptions can vary and have severe consequences for colepid species identification. Moreover, even the strains belonging to the groups 1 and 2 discovered by the phylogenetic analyses (Fig. 2) cannot discriminate morphotypes because they can neither be assigned to a certain cell morphology nor to the possession of algal endosymbionts. This questions the traditional morphology-based taxonomy. The separation of Coleps hirtus hirtus, C. hirtus viridis and C. spetai, which Foissner et al.¹ differentiated by the presence of zoochlorellae in the latter two species and the number of windows in the armor plates, could not be supported by our analyses. C. hirtus viridis was originally described by Ehrenberg^48,49 as C. viridis and later transferred as synonym of C. hirtus by Kahl⁵⁰ based on almost identical morphological features. However, Foissner²² described C. spetai for the green Coleps because of the morphological discrepancies to the Ehrenberg’s C. viridis (presence of only 11 windows per plate row and smaller cell size in C. viridis; see Table 1 for comparison). Our study has clearly demonstrated that most of the morphological features are variable and the limits for species separation were too narrow. Therefore, we propose the re-establishment of C. viridis for group 1 and C. hirtus for group 2, both with emended descriptions as follows. Considering our findings, the morphological descriptions of C. spetai, C. hirtus viridis and C. hirtus hirtus cannot be applied for (sub-) species separation any more. Consequently, we deal with a cryptic species complex, i.e., two genetically different groups that are fused in a highly variable morphotype including features of all three (sub-) species. To solve this taxonomic problem, two possible scenarios can be proposed: (1) We merge the three morphotypes under C. hirtus, the type species of Coleps. As a consequence, two new species needed to be proposed for both groups 1 and 2, which could be done following the suggestion of Sonneborn⁵¹ for the P. aurelia-complex. However, Sonneborn based his new descriptions on results of mating experiments, which are not applicable for Coleps here because conjugations have not been reported and the conditions for the induction of sexual reproduction are unknown. (2) To avoid confusion by introducing new species names, we propose keeping the already existing names, i.e., C. viridis for group 1 and C. hirtus for group 2 including the synonyms (see below).

Clonal cultures of both genetically varying Coleps groups have been deposited in the CCAP culture collection. Future studies may therefore be able to investigate, for example, sibling among strains or predator-prey experiments revealing spine- or wing-formation.

Coleps viridis Ehrenberg 1831 (printed 1832), Abh. Königl. Akad. Wiss. Berlin 1832: 101.

Synonym: Coleps spetai Foissner 1984, Stapfia 12: 21-22, Fig. 7, SP: 1984/10 and 1984/11 (lectotype designated here deposited in LI, see Aescht 2008: Denisia 23: 179), Coleps hirtus sensu Kahl 1930, Tierwelt Deutschlands 18: 134.

Diagnosis: Differed from other colepid ciliates by their SSU and ITS rDNA sequences (MT253680).

Lectotype (designated here): Fig. II, Tab. XXXIII, 3 in Ehrenberg 1838, Infusionsthierchen als vollkommene Organismen, p. 314.

Improved Description (specifications in brackets apply to our reference strain CCAP 1613/7): Coleps with conspicuous armor composed of six tiers with plate windows of the hirtus-type. With or without green algal endosymbionts. Cell size 44–63 × 21–35 μm (52–54 × 35–36 μm). Total number of windows in length rows 12–16 (14–16), number of windows of anterior primary plates 3–6 (4–6), number of windows of anterior secondary plates 2–3 (2), number of windows of posterior primary plates 4–5 (4–5), number of windows of posterior secondary plates 2–3 (2–3). One caudal cilium (1). With 0-2 anterior (0–1) and 0–5 posterior (1–4) spines, respectively.

Reference material (designated here for HTS approaches): The reference strain CCAP 1613/7 permanently cryopreserved at CCAP in a metabolically inactive stage.

Locality of reference strain: Plankton of Lake Mondsee, Upper Austria, Austria (47° 50′ N, 13° 23′ E).

Coleps hirtus (O.F. Müller) Nitzsch ex Ersch & Gruber 1827, Allgemeine Encyclopädie der Wissenschaften und Künste 16: 69, NT (proposed by Foissner 1984, Stapfia 12: 22, fig. 8): 1984/12 and 1984/13 (LI, in Aescht 2008: Denisia 23: 159).

Protonym: Cercaria hirta O.F. Müller 1786, Animalcula Infusoria: 128, tab. XIX, fig. 17, 18 (lectotype designated here).

Diagnosis: Differed from other colepid ciliates by their SSU and ITS rDNA sequences (MT253687).

Improved Description: Coleps with spiny armor composed of six tiers with plates of the hirtus-type. Without green algal endosymbionts. Cell size 42–52 × 23–28 μm. Total number of windows in length rows 12-13, number of windows of anterior primary plates 3-5, number of windows of anterior secondary plates 2, number of windows of posterior primary plates 4-5, number of windows of posterior secondary plates 2. One caudal cilium. Without anterior and 1-4 posterior spines, respectively.

Reference strain (designated here for HTS approaches): The strain CCAP 1613/14 permanently cryopreserved at CCAP in a metabolically inactive stage.

Locality of reference strain: Plankton of Lake Piburg, Tyrol, Austria (47° 11′ N, 10° 53′ E).

Molecular phylogeny of the Colepidae (Prostomatea)

The colepids belonging to the Prostomatea form a monophyletic lineage in the phylogenetic analyses of SSU rDNA sequences (Fig. 2). Mixotrophic as well as heterotrophic Coleps strains that resembled C. hirtus and C. spetai clustered in group 1 whereas group 2 included only two specimens which were identified as C. hirtus. These findings confirm the results of Barth et al.²⁹ with one exception. The authors found a clear separation into mixotrophic and heterotrophic species, which were therefore assigned to a C. spetai-(with endosymbionts) and a C. hirtus-group (without endosymbionts), respectively. Despite the difficulties of identifying these species by morphology, both groups clearly differed in their SSU and ITS rDNA sequences (Fig. 3). The ITS-2/CBC approach introduced for green algae (details in Darienko et al.⁵²) clearly demonstrated that both groups represented two separate ciliate species from a molecular point of view, which was also confirmed by analyses of the V9 region of the SSU, a region commonly used for metabarcoding (Figs. 4 and 5).

Our study also confirmed the findings of Chen et al.⁷, Lu et al.⁹, and Moon et al.²⁸, showing that the generic concept of colepid ciliates needs to be revised. None of the genera represented by more than one species is monophyletic. For example, the three species of Nolandia belonged to separate lineages. Nolandia nolandi was a sister to our studied strains, whereas both other species were closely related to taxa of Apocoleps, Pinacocoleps, and Tiarina (Fig. 2). The genus Levicoleps and Coleps amphacanthus formed a monophyletic clade representing another example that the generic conception is artificial and needs to be revised. However, to provide a new generic concept of colepid ciliates, it is necessary to study more of the described species by using an integrative approach including experimental approaches on, e.g., the formation of spines. For example, we clearly demonstrated that one key feature, which is the presence/absence of anterior/posterior spines, is highly variable and can therefore not be used to separate colepid genera as indicated by Foissner et al.¹² (Fig. 1). There is a need for more experimental studies with colepids belonging to the Cyclidium viridis and C. hirtus morphotype. Therefore, we deposited all clones used in this study in the CCAP culture collection. One option would be to incorporate all species into one genus, i.e., Coleps in revised form.

Endosymbiosis in Coleps

Some strains of Coleps are known to bear green algal endosymbionts¹. These green algae have Chlorella-like morphology (Fig. 6) and were identified as Micractinium conductrix (Fig. 7). So far, this alga was only known as endosymbiont of the ciliate Paramecium bursaria³⁴. All green algal endosymbionts of Coleps harbored this Micractinium species. In contrast, Pröschold et al.³⁴ found that one ciliate strain identified as C. hirtus viridis had Chlorella vulgaris as endosymbiont (the algae has been deposited in the Culture Collection of Algae and Protozoa under the number CCAP 211/111). Unfortunately, this ciliate strain is not available anymore⁵³.

Ecology and distribution

For limnological studies, the preservation with Bouin’s solution and QPS is an appropriate method for quantifying and identifying ciliate species in environmental samples⁵⁴. However, the quality in characterization of ciliates at the species level is sometimes limited as, in case of Coleps, the characteristic armored calcium carbonate plates are dissolved by the acidified fixation solution. Therefore, in our study, we could only distinguish between algal-bearing (mixotrophic) and non-algal-bearing (heterotrophic) Coleps. Despite that limitation, we could clearly see that the heterotrophic ones were only found in the deepest zones of both lakes (Fig. 8A). Not surprisingly, Coleps is often observed in nutrient- and ion-rich and also oxygen-depleted freshwater habitats or areas, e.g., sulfurous and crater lakes^{1,5,6,27,55,56} or even in the sludge of wastewater treatment plants⁵⁷. Mixotrophic individuals of Coleps were mainly found in the upper layers of both lakes, whereas in Lake Mondsee we could also detect specimens down to 40 m depth (Fig. 8). In contrast to the mero- and monomictic Lake Zurich^4,10,58,59, Lake Mondsee is holo- and dimictic⁶⁰. During mixis events, algal-bearing Coleps specimens can be transferred passively from the upper layers into the deeper zones and vice versa. Although morphotype countings and HTS analyses reads matched quite well, we found discrepancies that have already been discussed before^10,61 (Fig. S2).

Biogeographic aspects (haplotype network)

Our metabarcoding approach showed that C. viridis was found in both lakes as a common ciliate (Fig. S2). In contrast, C. hirtus could not be detected during the sampling period. To obtain more information about the distribution of both species, we used the BLASTn search algorithm⁶² (100 coverage, >97% identity) for the V4 and the V9 regions of the SSU and the ITS-2 sequences. No records using the V9 and the ITS-2 approaches could be discovered in GenBank, but 25 reference sequences using the V4 (Table S3). Together with the newly sequenced strains, we therefore constructed a V4 haplotype network (Fig. 10). Both groups are obviously widely distributed and subdivided into five (group 1) and four (group 2) haplotypes, respectively. All reference sequences were collected from freshwater habitats except for two marine records⁶³ (EU446361 and EU446396; Mediterranean Sea) and showed no geographical preferences.

Co-occurrence networks

In the sub-networks of C. viridis in both lakes, we found several significant correlations that pointed to either potential prey items, e.g., diverse flagellated autotrophic or heterotrophic protists or co-occurring ciliates (Fig. 9). Also, the smaller ciliates such as Cinetochilum margaritaceum or Cyclidium glaucoma may as well be considered as food for the omnivorous C. viridis (for a compilation of the food spectrum; see Foissner et al.¹). However, we identified the endosymbiont M. conductrix and its host C. viridis from both sub-networks of Lake Mondsee but not of Lake Zurich (Fig. 9). Despite this result, we want to point out that we may probably not find M. conductrix free-living in a water body because outside their ciliate host the algae were immediately attacked and killed by so-called Chlorella-viruses⁶⁶. Therefore, the HTS-detection of M. conductrix was probably only together with a host ciliate. This might further explain why the green algae were detected in Lake Mondsee even in the aphotic 40 m zone where photosynthesis was impossible and individuals probably passively transferred into the deeper area by lake mixis.

Outlook

As demonstrated in our study, the combination of traditional morphological investigations, which includes the phenotypic plasticity of the cloned strains, and modern molecular analyses using both SSU and ITS sequencing as well as HTS approaches advise a taxonomic revision of the genus Coleps. This comprehensive and integrative approach is also applicable for other ciliate species and genera and will provide new insights into the ecology and evolution of this important group of protists.

Experimental procedures

Study sites, lake sampling and origin of the Coleps strains

Our main study sites were Lake Mondsee (Austria) and Lake Zurich (Switzerland), two pre-alpine oligo-mesotrophic lakes that were sampled at the deepest point of each lake (Table S4). Water samples were taken monthly from June 2016 through May 2017 over the whole water column and additionally biweekly at two main depths, i.e., 5 m in both lakes, 40 m in Lake Mondsee, and 120 m in Lake Zurich, respectively. A 5-L-Ruttner water sampler was used for Lake Zurich and a 10-L-Schindler-Patalas sampler (both from Uwitec, Austria) for Lake Mondsee. Twelve Coleps strains were isolated from Lake Mondsee and one from Lake Zurich. Another six clones could be obtained either from already successfully cultivated own strains, fresh isolates or from culture collections. Detailed information about sampling sites, dates and strain numbers is given in Table S2.

Seasonal and spatial distribution and abundance

For quantification, subsamples (200-300 mL) were preserved with Bouin’s solution (5% f.c.) containing 15 parts of picric acid, 5 parts of formaldehyde (37%) and 1 part of glacial acetic acid⁵⁴. The samples were filtered through 0.8 μm cellulose nitrate filters (Sartorius, Germany) equipped with counting grids. The ciliates were stained following the protocol of the quantitative protargol staining (QPS) method after Skibbe⁵⁴ with slight modifications after Pfister et al.⁶⁷. The permanent slides were analyzed by light microscopy up to 1600x magnification with a Zeiss Axio Imager.M1 and an Olympus BX51 microscope. For identification of Coleps and Nolandia cells, the identification key of Foissner et al.¹ was used. Microphotographs were taken with a ProgRes C14 plus camera using the ProgRes Capture Pro imaging system (version 2.9.0.1, Jenoptik, Jena, Germany).

Cloning, identification and cultivation of ciliates and endosymbionts

Single cells of Coleps were isolated and washed using the Pasteur pipette method⁶⁸. The isolated strains were cultivated in 400 μl modified Woods Hole medium⁶⁹ (MWC; modified) and Volvic mineral water in a mixture of 5:1 and with the addition of 10 μl of an algal culture (Cryptomonas sp., strain SAG 26.80) as food in microtiter plates. These clonal cultures were transferred into larger volumes after successful enrichment. All cultures were maintained at 15–21 °C under a light: dark cycle of 12:12 h (photon flux rate up 50 mol m⁻² s⁻¹).

For the isolation of their green algal endosymbionts, single ciliates were washed again and transferred into fresh MWC medium. After starvation and digestion of any food, after approx. 24 hrs, cells were washed again and the ciliates transferred onto agar plates containing Basal Medium with Beef Extract (ESFl; medium 1a in Schlösser⁷⁰). Before placement of the ciliates onto agar plates, 50 μm of an antibiotic mix (mixture of 1% penicillin G, 0.25% streptomycin, and 0.25% chloramphenicol) were added to prevent bacterial growth. The agar plates were kept under the same conditions as described. After growth (6–8 weeks), the algal colonies were transferred onto agar slopes (1.5%) containing ESFl medium and kept under the described culture conditions.

For light microscopic investigations of the algae, Olympus BX51 and BX60 microscopes (equipped with Nomarski DIC optics) were used. Microphotographs were taken with a ProgRes C14 plus camera using the ProgRes Capture Pro imaging system (version 2.9.0.1, Jenoptik, Jena, Germany).

PCR, sequencing and phylogenetic methods

Single-cell PCR was used to obtain the sequences of the Coleps strains. Before PCR amplification, single cells of Coleps were washed as described above. After starvation followed by additional washing steps, cells were transferred into 5 μm sterile water in PCR tubes and the prepared PCR mastermix containing the primers EAF3 and ITS055R⁷¹ was added. After this primary PCR amplification and subsequent PCR purification, a nested PCR was conducted using the primer combinations EAF3/N1400R and N920F/ITS055R⁷¹.

The sequences of the Coleps strains were aligned according to their secondary structures of the SSU and ITS rDNA (see detailed folding protocol described in Darienko et al.⁵²) and included into two data sets: (i) 34 SSU rDNA sequences (1,750 bp) of representatives of all members of the Prostomatea and (ii) 19 ITS rDNA sequences (538 bp) of the investigated strains. Genomic DNA of the green algae was extracted using the DNeasy Plant Mini Kit (Qiagen GmbH, Hilden, Germany). The SSU and ITS rDNA were amplified using the Taq PCR Mastermix Kit (Qiagen GmbH, Hilden, Germany) with the primers EAF3 and ITS055R. The SSU and ITS rDNA sequences of the isolated green algae (aligned according to the secondary structures) were included into a data set of 31 sequences (2,604 bp) of representatives of the Chlorellaceae (Trebouxiophyceae).

GenBank accession numbers of all newly deposited sequences can be found in Table S2 and in Fig. 7, respectively. For the phylogenetic analyses, the datasets with unambiguously aligned base positions were used. To test which evolutionary model fit best for both data sets, we calculated the log-likelihood values of 56 models using Modeltest 3.7⁷² and the best models according to the Akaike criterion by Modeltest were chosen for the analyses. The settings of the best models are given in the figure legends. The following methods were used for the phylogenetic analyses: distance, maximum parsimony, maximum likelihood, and Bayesian inference. Programs used included PAUP version 4.0b164⁷³, and MrBayes version 3.2.3⁷⁴.

The secondary structures were folded using the software mfold⁴², which uses the thermodynamic model (minimal energy) for RNA folding.

Haplotype networks

The haplotypes of the V4 region were identified among the groups of Coleps (see Fig. S1). The present haplotypes and the metadata (geographical origin and habitat) of each strain belonging to the different haplotypes are given in Table S3. To establish an overview on the distribution of the Coleps groups, the V4 haplotypes were used for a BLASTn search⁶² (100% coverage, >97% identity). To construct the haplotype networks, we used the TCS network tool^64,65 implemented in PopART⁷⁵.

High-throughput sequencing of the V9 18S rDNA region and subsequent bioinformatic analyses

On each sampling date, water samples for a high-throughput sequencing approach (HTS) were taken in depths of 5 m and 40 m at Lake Mondsee and 5 m and 120 m depths in Lake Zurich. DNA extraction, amplification of the V9 SSU rDNA, HTS and quality filtering of the obtained raw reads was conducted as described in Pitsch et al.¹⁰. After quality filtering, all remaining reads were subjected to a two-level clustering strategy⁷⁶. In the first level, replicated reads were clustered in SWARM version 2.2.2 using d=1⁷⁷. In the second level, the representative sequences of all SWARM OTUs were subjected to pairwise sequence alignments in VSEARCH version 2.11.0⁷⁸ to construct sequence similarity networks at 97% sequence similarity. The network sequence clusters (NSCs) resulting from the second level of clustering were then taxonomically assigned by running BLASTn analyses against NCBI’s GenBank flat-file release version 230.0 and the Coleps SSU sequences obtained from single-cell sequencing. Network sequence clusters were assigned to Coleps, if the closest BLAST hit of the NSC representative sequence was a Coleps reference sequence. Furthermore, the NSC representative sequence had to share a fragment of at least 48 consecutive nucleotides and at least 90% sequence similarity to a reference sequence in order to be assigned to Coleps.

Co-occurrence networks

With the protist community data matrix resulting from HTS, we further conducted co-occurrence network analyses to assess biotic and abiotic interactions of Coleps. For each lake and depth, we ran network analyses with NetworkNullHPC (https://github.com/lentendu/NetworkNullHPC) following the null model strategy developed by Connor et al.⁷⁹. This strategy was especially designed for dealing with HTS datasets and allows for inferring statistically significant correlations between NSCs while minimizing false positive correlation signals. We screened the resulting networks for Coleps nodes and extracted their subnetworks including all directly neighbouring co-occurrence partners as well as all edges between Coleps and its neighbours and the neighbours themselves.

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