Available online at www.sciencedirect.com
European Journal of Protistology 49 (2013) 477–486
New Paramecium quadecaurelia strains (P. aurelia spp. complex, Ciliophora) identified by molecular markers (rDNA and mtDNA) Ewa Przybo´sa , Sebastian Tarcza,∗ , Eike Dusib a
Department of Experimental Zoology, Institute of Systematics and Evolution of Animals, Polish Academy of Sciences, Kraków 31-016, Sławkowska 17, Poland b Workgroup Limnology, Institute of Hydrobiology, Technische Universität Dresden, Dresden 01062, Zellescher Weg 40, Germany Received 3 August 2012; received in revised form 2 October 2012; accepted 8 November 2012 Available online 3 January 2013
Abstract Paramecium quadecaurelia is a rare species (previously known only from two locations) belonging to the P. aurelia species complex. In the present paper, fragments of an rDNA gene (ITS1-5.8S-ITS2-5 rDNA) and mtDNA genes (cytochrome oxidase subunit I and cytochrome b regions) were employed to assist in the identification and characterization of three new strains collected from Ecuador and Thailand. Molecular data were confirmed by mating reactions. In rDNA and mtDNA trees constructed for species of the P. aurelia complex, all P. quadecaurelia strains, including the three new strains discussed in this study and two known previously from Australia and Africa, form a monophyletic but differentiated clade. The present study shows that genetic differentiation among the strains of P. quadecaurelia is equal to or even greater than the distances between some other P. aurelia species, e.g., P. primaurelia and P. pentaurelia. Such great intra-specific differentiation may indicate a future splitting of the P. quadecaurelia species into reproductively isolated lines. © 2012 Elsevier GmbH. All rights reserved. Keywords: Paramecium quadecaurelia; P. aurelia species complex; rDNA (ITS1-5.8S-ITS2-5 LSU rDNA); Cytochrome oxidase subunit I (COI) gene; Cytochrome b (CytB) gene; Geographic distribution
Introduction Several ciliate genera reveal a complex structure of their taxonomic species, which consist of a number of reproductively isolated groups termed syngens or sibling species. For the first time the possibility of the existence of sibling species in ciliates was described in Paramecium aurelia by Sonneborn (1937). At present the Paramecium aurelia species complex is composed of fifteen genetic species, 14 of which were named by Sonneborn (1975), and the fifteenth (P. sonneborni) was described separately (Aufderheide
∗ Corresponding
author. E-mail address:
[email protected] (S. Tarcz).
0932-4739/$ – see front matter © 2012 Elsevier GmbH. All rights reserved. http://dx.doi.org/10.1016/j.ejop.2012.11.001
et al. 1983). Although particular members of the complex are morphologically indistinguishable, they are true biological species (identified by strain crosses); moreover, particular species differ according to geographical distribution, temperature preferences, culture conditions necessary for conjugation, and the system of mating type inheritance (Stoeck et al. 2000). The increasing role of molecular biology techniques in taxonomic studies of unicellular eukaryotes seems to be an appropriate tool not only for genetic variability assessment, but also for fast identification of P. aurelia species. Previous studies have shown that particular species of the complex have different levels of intraspecific polymorphism as revealed by molecular methods based on PCR fingerprinting (RAPD – Random Amplified Polymorphic DNA (Stoeck
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et al. 1998, 2000); RAPD, RFLP – Restriction Fragment Length Polymorphism and ARDRA – Amplified Restriction rDNA Analysis (Przybo´s et al. 2007)). The existence of syngens (sibling species) within some Paramecium species has also been revealed or confirmed by the sequencing of gene fragments such as ITS1-5.8S-ITS2-5 LSU rDNA and COI mtDNA, e.g., in Paramecium calkinsi (Przybo´s et al. 2012), P. multimicronucleatum (Tarcz et al. 2012), and P. bursaria (Greczek-Stachura et al. 2012). Haentzsch et al. (2011) has developed specific multiplex PCR primers based on 18S rDNA sequences for the detection of species belonging to the P. aurelia complex and the genus Paramecium. These primers can be used for the fast determination of ciliate communities during water testing, as particular Paramecium species are indicative of water quality. In the present paper, molecular markers, i.e., fragments of an rDNA gene (ITS1-5.8S-ITS2-5 LSU rDNA) and mtDNA genes (cytochrome oxidase subunit I and cytochrome b regions – designated COI and CytB, respectively) were applied for the identification and characterization of new strains belonging to the P. aurelia species complex. We used three markers because at least a two-locus approach should be used in molecular phylogenies (Dunthorn et al. 2011). Here, we present the results of a molecular identification subsequently confirmed by mating reactions.
Material and Methods Identification of species of the P. aurelia complex by mating reactions New strains originating from Ecuador and Thailand (Table 1) as well as previously known strains of P. quadecaurelia, collected in Australia and Namibia, were used in the present study. Data concerning other species of the P. aurelia complex and other species of the genus Paramecium are given in Table 2 . Paramecia were cultured at 27 ◦ C in a medium of dried lettuce in distilled water inoculated with Enterobacter aerogenes and identified according to Sonneborn (1950, 1970) by mating reactions. The new strains from Ecuador and Thailand, mature for conjugation, were mated with reactive complementary mating types (c.m.t.) of the reference (standard) strains of species of the P. aurelia complex. The following standard strains were used:
Strain 90, Pennsylvania, USA, P. primaurelia; Strain Rieff, Scotland, GB, P. biaurelia; Strain 324, Florida, USA, P. triaurelia; Strain 87, Pennsylvania, USA, P. pentaurelia; Strain 159, Puerto Rico, P. sexaurelia; Strain 246, Mississippi, USA, P. dodecaurelia; Strain AN1-1, Namibia, Africa, P. quadecaurelia.
Conjugation was carried out in the following manner: 0.5 mL samples of a strain to be identified and of a strain representing a known species of the P. aurelia complex were taken from cultures of sexually reactive paramecia (clones derived from one specimen) forming rings at the top of the test tubes; the samples were subsequently mixed in depression slides. Species were determined based on the occurrence of 85–95% initial agglutination of paramecia (under optimal conditions for particular species) followed by the presence of tight conjugating pairs formed by paramecia from a studied strain and a particular reference strain. To make sure that no intra-strain conjugation (selfing) occurred within c.m.t., controls of non-crossed c.m.t. were also cultured and observed. The F1 generation of hybrids from inter-strain crosses of P. quadecaurelia (new strain from Ecuador or Thailand × strain from Namibia, Africa; strain from Ecuador × strain from Thailand) was obtained by conjugation, and F2 by autogamy by the daily isolation lines method (according Sonneborn 1950, 1970). The occurrence of autogamy was examined in preparations stained with acetocarmine. The dye stains nuclear apparatuses in paramecia, showing old fragmented macronuclei as well as new developing ones. The survival of clones in both generations was estimated from a total of 100 clones. Clones were considered survivors after undergoing 6–7 fissions during 72 h following the separation of conjugation partners or post-autogamous caryonids (the two products of the first fission of each autogamous paramecium). The procedures were carried out pursuant to Chen (1956). The percentage of surviving hybrid clones in crosses was compared in F1 and F2 (Table 3) because paramecium species were identified not only on the basis of their capacity to conjugate with the reference specimens, but also to produce viable recombinant F2 clones (Sonneborn 1975). All the reference (standard) strains of particular species of the P. aurelia complex (with the exception of the P. quadecaurelia strain AN1-1, cf. Przybo´s et al. 2003) were obtained from the laboratories of Prof. T.M. Sonneborn (Department of Biology, Indiana University, USA) or Prof. G.H. Beale (Institute of Animal Genetics, Edinburgh University, Great Britain). They are kept at the Department of Experimental Zoology, Institute of Systematics and Evolution of Animas, Polish Academy of Sciences, Cracow, Poland. These materials will be available on request.
Molecular methods All strains used for DNA isolation were homozygous, as they passed autogamy previously. Paramecium genomic DNA was isolated from vegetative cells at the end of the exponential phase (approx. 1000 cells were used for DNA extraction) using a NucleoSpin Tissue Kit (Macherey-Nagel, Germany) according to the manufacturer’s instructions for DNA isolation from cell cultures. The only modification was centrifugation of the cell culture for 20 min at 13,200 rpm.
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Table 1. Strains of Paramecium quadecaurelia identified to date. No.
Strain designation
1.
E9II
2.
Ta3a
3. 4.
Ta7a AN1-1 (DO-207)b
5.
328 (SO-328)b
Strain geographical origin
Collector’s name
Ecuador, Loyola. small pond on a grazing land Thailand, Samui island, lake
E. Dusi May 2010
Namibia, Vindhoek, Pond in park Australia, Emily Gap near Alice Springs
S. Dobretzov, 2001 G. Beale, pre 1958
Reference
Present paper
T. Fokin September 2010 Przybo´s et al. (2003) Sonneborn (1975)
GenBank accession numbers
ITS1-5.8SITS2-5 LSU rDNA
COI mtDNA
CytB
JX010637
N/A
JX010677
JX010638
JX010659
JX010678
JX010639 JX010636
JX010660 JX010658
JX010679 AM949774c
JX010635
JX010657
AM949773c
a Ta3
and Ta7 originated from the same sample collected in the lake, they may be the sister clones, as their haplotypes are the same (Fig. 1a–c). Genbank accession numbers for cytosol type hsp70 gene of the strain AN1-1 – AB106337, and the strain 328 – AB106336 (Przybo´s et al. 2003). c Designations and GenBank numbers as in (Barth et al. 2008).
b The
The supernatant was removed and the remaining cells were suspended in lysis buffer and proteinase K. Fragments of rDNA, COI, and CytB genes were sequenced and analyzed. First, rDNA fragments were amplified with ITS1 and ITS4 universal eukaryotic primers (White et al. 1990) and ITS3zg and 3pLSU primers developed with OligoAnalyzer 3.1 (http://www.eu.idtdna.com/analyzer/ applications/oligoanalyzer) (Table 4). F388dT and R1184dT primers (Table 4) and the protocol previously described by Strüder-Kypke and Lynn (2010) were used for the amplification of the COI fragment of mitochondrial DNA. In some cases, when the above COI pair of primers did not yield a well-defined product, the internal primer CoxH10176 (Barth et al. 2006) was used instead of R1184dT. To amplify the CytB gene fragment, the primer pair CytBF/PaCytR and the protocol previously described by Barth et al. (2008) were used. PCR amplification for all analyzed DNA fragments was carried out in a final volume of 40 L containing 30 ng of DNA, 1.5 U Taq-Polymerase (EURx, Poland), 0.8 L of 20 M of each primer, 10× PCR buffer, and 0.8 L of 10 mM dNTPs. After amplification, PCR products were electrophoresed in 1% agarose gel for 45 min at 85 V with a DNA molecular weight marker (Mass Ruler Low Range DNA Ladder, Fermentas, Lithuania). NucleoSpin Gel and PCR Clean-up (Macherey-Nagel, Germany) was used for purifying PCR products. In some PCR products, additional sub-bands were obtained apart from the main band. In these cases, 30 L
of each PCR product was separated on 1.8% agarose gel (100 V/60 min) with a DNA molecular weight marker (Mass Ruler Low Range DNA Ladder, Fermentas, Lithuania). Then the band representing the examined fragment was cut out and purified. Sequencing was done in both directions with the application of BigDye Terminator v3.1 chemistry (Applied Biosystems, USA). The primers used in PCR reactions were applied for sequencing the rDNA region, and the primer pair M13F/M13R was used for sequencing the COI fragment (Table 4). The sequencing reaction was carried out in a final volume of 10 L containing 3 L of template, 1 L of BigDye (1/4 of the standard reaction), 1 L of sequencing buffer, and 1 L of 5 M primer. Sequencing products were precipitated using Ex Terminator (A&A Biotechnology, Poland) and separated on an ABI PRISM 377 DNA Sequencer (Applied Biosystems, USA). The sequences are available in the NCBI GenBank database (see Tables 1 and 2).
Data analysis Sequences were examined using Chromas Lite (Technelysium, Australia) to evaluate and correct the chromatograms. The alignment of the studied sequences was performed using ClustalW (Thompson et al. 1994) within the BioEdit software (Hall 1999) and checked manually. All of the obtained sequences were unambiguous and were used for analysis. Phylograms were constructed for the studied fragments with Mega v5.0 (Tamura et al. 2011), using neighbor-joining
480
Table 2. Other strains of Paramecium genus used in present studies. No.
P. primaurelia P. primaurelia P. primaurelia P. biaurelia P. biaurelia P. biaurelia P. biaurelia P. triaurelia P. triaurelia P. triaurelia P. tetraurelia P. tetraurelia P. tetraurelia P. pentaurelia P. pentaurelia P. pentaurelia P. sexaurelia P. sexaurelia P. sexaurelia P. septaurelia P. septaurelia P. septaurelia P. octaurelia P. octaurelia P. novaurelia P. novaurelia P. novaurelia P. decaurelia P. decaurelia P. undecaurelia P. dodecaurelia P. dodecaurelia P. dodecaurelia P. tredecaurelia P. tredecaurelia P. sonneborni
Strain designation
90 (SO-90a ) SS PR-08a Rieff USB PR-34a DB-05a 324 (SO-324a ) SCM HH-05a S (PR-92a ) FP SO-51a 87 (SO-87a ) HBB NA-05a 159 (SO-159a ) SAS FO-128a 38 (SO-38a ) AZ24-4 PO-162a 138 (SO-168a ) IEA (PR-169a ) CS UG (PR-175a ) ED-05a 223 (SO-223a ) JH 219 (SO-219a ) 246 (SO-246a ) GM SK-199a 209 (SO-209a ) IKM (PR-205a ) ATTC 30995 (AU-208a )
Geographical origin
USA, Pennsylvania Spain, Andalusia, Sevilla Russia, Astrakhan Nature Reserve Great Britain, Scotland, Rieff USA, Boston Russia, Irkutsk Germany, Saidenbach Reservoir USA, Florida Spain, Castile Germany, Hannover Australia, Sydney France, Paris USA, Indiana USA, Pennsylvania Hungary, Balatonfüzfo Italy, Naples Puerto Rico Spain, Andalusia, Seville Japan, Yamaguchi USA, Florida Russia, Astrakhan Nature Reserve Russia, Astrakhan Nature Reserve USA, Florida Israel, Ein Effek Czech Republic, Ceske Skalnice Ukraine, Gorgany Mts. UK, Edinburgh USA, Florida Japan, Honshu Island USA, Texas USA, Mississipi Germany, Münster USA, Hawaii France, Paris Israel, Kiryat Motzkin USA, Texas
GenBank accession numbers ITS1-5.8S-ITS2-5 LSU rDNA
COI mtDNA
CytB
JF304163 JN998643 N/A JX010640 JX010641 N/A N/A JX010642 JX010643 N/A JF304164 JN998645 N/A JX010644 JX010645 N/A JX010646 JX010647 N/A JX010648 JX010649 N/A JX010650 JX010651 JX010652 JX010653 N/A JX010654 JX010655 JX010656 JN998639 JN998615 N/A JF304165 JN998647 JF304167
JF304182 JN998685 N/A JX010661 JX010662 N/A N/A JX010663 JX010664 N/A JF304183 JN998687 N/A JX010665 JX010666 N/A JX010667 JX010668 N/A JX010669 JX010670 N/A JX010671 JX010672 EU056250 EU056263 N/A JX010673 JX010674 JX010675 JN998681 JN998657 N/A JF304184 JN998689 JX010676
AM949780a N/A AM949779a N/A N/A AM949784a AM949785a AM949778a N/A AM949777a AM949771a N/A AM949770a AM949782a N/A AM949781a AM949765a N/A AM949764a AM949766a N/A AM949768a AM949767a AM949772a N/A AM949776a AM949775a AM949769a N/A AM949783a AM949763a N/A AM949762a AM949760a AM949761a AM949786a
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1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29. 30. 31. 32. 33. 34. 35. 36.
Species
AM949759a AM949758a N/A N/A AM949757a AM949756a AM072773 JF304188 JF741273 JF304189 N/A N/A JF304168 JF304171 JF741241 JF304172 N/A N/A
Results and Discussion Occurrence and distribution of the P. aurelia spp. complex
a Designations
b GenBank
and GenBank numbers as in (Barth et al. 2008). numbers as in (Barth et al. 2006).
China, Shanghai Saudi Arabia, Al Riyadh USA, Boston USA, Louisiana, Baton Rouge Germany, Martinfeld Italy, Naples Sh1-38 SA (S-AUa ) AB9-20 BR GMA-2a ISN-11a P. shewiakoffi P. jenningsi P. multimicronucleatum P. multimicronucleatum P. multimicronucleatum P. multimicronucleatum
481
(NJ) (Saitou and Nei 1987), maximum parsimony (MP) (Nei and Kumar 2000), and maximum likelihood (ML). All positions containing gaps and missing data were eliminated. NJ analysis was performed using Mega v5.0 program, by bootstrapping with 1000 replicates (Felsenstein 1985). MP analysis was evaluated with the min-mini heuristic parameter (at level 2) and bootstrapping with 1000 replicates. Bayesian inference (BI) was performed with MrBayes 3.1.2 (Ronquist and Huelsenbeck 2003); analysis was run for 5,000,000 generations and trees were sampled every 100 generations. All trees for BI analysis were constructed with TreeView 1.6.6 (Page 1996). Analysis of haplotype diversity, nucleotide diversity, and variable nucleotide positions was done with DnaSP v5.10.01 (Librado and Rozas 2009). Analysis of nucleotide frequencies, uncorrected p-distance estimation, and identification of substitution models (T92+G+I for rDNA and HKY+G+I for mtDNA fragments) for ML analysis were done with Mega v5.0 (Tamura et al. 2004, 2011).
b
37. 38. 39. 40. 41. 42.
ITS1-5.8S-ITS2-5 LSU rDNA
Strain designation Species No.
Table 2 (Continued).
Geographical origin
GenBank accession numbers
COI mtDNA
CytB
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Among the 15 known species of the P. aurelia complex (14 of them characterized by Sonneborn in 1975 and one – P. sonneborni – by Aufderheide et al. in 1983) some are cosmopolitan, e.g., P. primaurelia, P. biaurelia, P. tetraurelia, and P. sexaurelia, while others are limited to certain regions, environments, or even habitats (cf. Przybo´s 2005; Przybo´s and Fokin 2000; Przybo´s et al. 2008; Sonneborn 1975). P. quadecaurelia is an example of a species limited to a certain area. Recently, some new strains of P. quadecaurelia have been found in Ecuador and Thailand; both localities being situated at a similar latitude. The strains were identified as P. quadecaurelia on the basis of conjugation between them and with the reference strain of the species (AN1-1 from Namibia, Africa; Przybo´s et al. 2003). No reaction (conjugation) was observed between P. quadecaurelia strains and the reference strains of other species of the P. aurelia complex. Strains of P. quadecaurelia (Fig. 2) sp. have the typical morphological appearance of a species belonging to the P. aurelia complex, the mean cell length being 150 m, comparable to that of P. biaurelia or P. sexaurelia, larger than P. tetraurelia (117 m) and P. primaurelia (140 m), and smaller than P. undecaurelia (170 m) (data according Sonneborn 1975). Previously, P. quadecaurelia was collected in Emily Gap, Australia, situated 15 km away from Alice Springs (strain 328, Sonneborn 1975) and in Vindhoek, Namibia, Africa (Przybo´s et al. 2003). The five strains of this species known to date have been found only in the tropical zone limited by the tropics of Capricorn and Cancer. It seems that Sonneborn’s
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Table 3. Percentage of surviving clones of P. quadecaurelia inter-strain hybrids. No.
Strain designation
Strain origin
Percentage of surviving clones F1 generation
1.
Ta7 x AN1-1
2.
E9II x Ta7
3.
E9II x AN1-1
4.
AN1-1 x 328
Thailand, Samui Island x Africa, Namibia Ecuador x Thailand, Samui Island Ecuador x Africa, Namibia Africa, Namibia x Australia
Reference
F2 generation
93
62
Present paper
90
58
Present paper
91
57
Present paper
100
96
Przybo´s et al. (2003)
Table 4. Primers used for amplification and sequencing of studied DNA fragments. DNA fragment
Primer
Sequence 5 –3
References
ITS1-5.8S-ITS2 ITS1-5.8S-ITS2 LSU rDNA LSU rDNA CytB mtDNA CytB mtDNA COI mtDNA
ITS1 ITS4 ITS3zg 3pLSU CytF PaCytR F388dTa
TCCGTAGGTGAACCTGCGG TCCTCCGCTTATTGATATGC CryAwCGATGAAGAACGCAGCC CAAGACGGGTCAGTAGAAGCC GGWACMATGCTRGCTTTYAG GGYCTAAAATATCAATGRGGTGC TGTAAAACGACGGCCAGTGGwkCbAAAGATGTwGC
COI mtDNA
R1184dTa
CAGGAAACAGCTATGACTAdACyTCAGGGTGACCrAAAAATCA
COI mtDNA sequencing primer
CoxH10176 M13F
GAAGTTTGTCAGTGTCTATCC TGTAAAACGACGGCCAGT
sequencing primer
M13R
CAGGAAACAGCTATGAC
White et al. (1990) White et al. (1990) Tarcz et al. (2012) Tarcz et al. (2012) Barth et al. (2008) Barth et al. (2008) Strüder-Kypke and Lynn (2010) Strüder-Kypke and Lynn (2010) Barth et al. (2006) Strüder-Kypke and Lynn (2010) Strüder-Kypke and Lynn (2010)
a Primers
used for amplification of COI fragment are composed of two parts – the first one is a degenerate primer (Forward or Reverse), specific to amplified COI sequence and the second one is a sequencing primer M13 (Forward or Reverse) (in bold face).
opinion (1957) that climatic zones are the main factor restricting the occurrence of some species of the P. aurelia complex has turned out to be true. Essays by Foissner (2006) and Foissner et al. (2008) on the geographical distribution of protists put forth the idea that ciliate biogeography is similar to that of plants and animals, but with an increased proportion of cosmopolitan species, favoring “the moderate endemicity model.” This also pertains to the question of dispersal of Paramecium species. Due to the fact that this genus does not produce cysts (Beale and Preer 2008), some water is necessary for its dispersal. At present it is difficult to explain whether the current distribution of P. quadecaurelia strains is the result of an ancient expansion or a recent dispersal due to Paramecium transfer by natural or anthropogenic factors. Based on the obtained results, P. quadecaurelia appears to be widely distributed but limited to the warm zone.
Genetic variation within P. quadecaurelia In the current analyses, based on a comparison of rDNA and mtDNA (COI and CytB) fragments, the average
genetic distance (uncorrected p-distance) between the studied strains of P. quadecaurelia was 0.004/0.052/0.093 (rDNA/COI/CytB). A previous molecular study of two P. quadecaurelia strains from Namibia, Africa (AN1-1) and Australia (328) (Przybo´s et al. 2003) revealed that the genetic distance of the studied hsp70 gene fragment was p = 0.008. It was identical to the genetic variability between the strains 328 and AN1-1 obtained for the currently studied rDNA fragment (p = 0.008). It is significant that the genetic variation of the three currently studied DNA fragments (rDNA/COI/CytB) is equal or even greater among strains of P. quadecaurelia than between some other P. aurelia species, e.g., P. primaurelia and P. pentaurelia (0.001/0.040/0.059 – rDNA/COI/CytB). Nevertheless, all P. quadecaurelia strains conjugate with each other, yielding viable hybrids (Table 3) with a high percentage of surviving clones of inter-strain hybrids in F1 and F2 generations (90–100% and 57–96%, respectively). Therefore, the studied strains form one species with a common gene pool. It is also worth emphasizing that the new P. quadecaurelia strains from Ecuador and Thailand were first identified by molecular markers. They were not reactive for a long time, perhaps because of long immaturity, so initially
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Fig. 1. Phylograms constructed for 35 (a, c) or 34 (b) Paramecium strains including for 5 (a, c) or 4 (b) strains of P. quadecaurelia studied in the present work. Two strains of P. multimicronucleatum were used as an outgroup. The construction of trees was based on a comparison of sequences from ITS1-5.8S-ITS2-5 LSU rDNA (a), COI mtDNA (b), CytB mtDNA (c) fragments using the neighbor joining method. Bootstrap values for neighbor joining, maximum parsimony analysis, maximum likelihood and posterior probabilities for Bayesian Inference are shown. All positions containing gaps were eliminated from the analysis. There were a total of 1060 positions in the final dataset. Phylogenetic analyses were conducted in MEGA 5.0 (NJ/MP/ML) and Mr Bayes 3.1.2 (BI).
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DNA fragments as a tool for identification of closely related taxons
Fig. 2. Paramecium quadecaurelia, strain 328, Australia. (a) Vegetative individual, giemsa stain. ×200. (b) Autogamous individual, giemsa stain. ×200.
their identification by the classical Sonneborn method was not possible. All P. quadecaurelia strains, those recently discovered in Ecuador, South America and Thailand, Asia and those previously known from Australia and Africa, appear in ribosomal and mitochondrial trees (Fig. 1a–c) as a differentiated but monophyletic cluster with high bootstrap support. Such great differentiation between remote strains of P. quadecaurelia might indicate a future splitting of the species into different, reproductively isolated lines. In all the trees (Fig. 1a–c), the studied strains of P. quadecaurelia present very similar relationships, despite the greater variability of mitochondrial markers (longer branches in Fig. 1b and c). Two strains from Thailand (Ta3 and Ta7) appear together and form a monophyletic cluster with the strain from Namibia, Africa (AN1-1). Furthermore, the Australian strain (328) appears as a separate branch, which is situated between the three above-mentioned ones (AN1-1, Ta3, Ta7) and the most distant strain from Ecuador (E9II) (Fig. 1a–c). It is worth mentioning that the DNA sequences of the two strains from Thailand (Ta3 and Ta7) are identical even if the highly variable mitochondrial fragments are compared. They originated from the same water sample, so it is highly probable that they may be sister clones. In the case of COI analysis, we were not able to obtain a DNA sequence for the strain from Ecuador, so it is absent from the tree (Fig. 1b). The P. quadecaurelia cluster appears in all the obtained trees in different positions (low bootstrap support for internal nodes), but always near P. primaurelia, P. pentaurelia, and P. novaurelia. This seems to be connected with the fact that all these sibling species have the same, caryonidal system of mating type inheritance.
Over the last decade, the application of molecular methods in the taxonomy of unicellular eukaryotes has increased. There are many examples of the use of molecular data on their own and in combination with other taxonomic methods, e.g., for the identification of planktonic protists in environmental samples (Duff et al. 2008), species of Oligotrichia, Ciliophora (Liu et al. 2011), and several species of the genus Paramecium, as well as in molecular phylogeny (Barth et al. 2006, 2008; Catania et al. 2009; Coleman 2005; Hori et al. 2006; Przybo´s et al. 2007). One purpose of the studies mentioned above is to gain the ability to identify species or cryptic species using standardized DNA fragments called DNA barcodes. For the first time a fragment of the mitochondrial COI gene was proposed as a universal DNA barcode (Hebert et al. 2003a,b) and successfully used for the identification of cryptic species in butterflies, in the Astraptes fulgerator species complex (Hebert et al. 2004). Similar attempts were made in different ciliate taxons. For example, Chantangsi et al. (2007) showed the potential utility of the COI mtDNA fragment by identifying the closely related species of the genus Tetrahymena. Similarly, Kher et al. (2011) applied the DNA barcoding technique for the assignment of unknown Tetrahymena isolates at the species level, but postulated that there was a need for the development of more universal PCR primers which could amplify the COI region in other Protista lineages. Broader analysis was conducted by Strüder-Kypke and Lynn (2010), who developed a set of universal primers for COI amplification in ciliates. The above primers were successfully used by Greczek-Stachura et al. (2012), which showed that the COI fragment can be used for the determination of Paramecium bursaria syngens. Moreover, the COI gene seems to be a good marker not only for barcoding analysis but also for phylogeographical studies (Gentekaki and Lynn 2012), which are still rare in unicellular eukaryotes. However, in other Protista groups, the application of the COI fragment has caused problems with setting universal PCR primes for the amplification of homologous COI sequences. For example, Hamsher et al. (2011) proposed as an alternative two other fragments for diatom species delineation: fragments of the rbcL gene and a highly variable part of LSU rDNA. Moreover, the latter seems to be a good alternative for microorganisms without typical mitochondria (and thus the COI gene) such as pelobionts, entamoebae, and diplomonads (Wylezich et al. 2010). In turn, Dunthorn et al. (2011) proposed a two-locus approach for phylogenetic analysis of Colpodea which uses both nuclear SSU rDNA and mitochondrial SSU genes as molecular markers. For this reason, the identification of P. quadecaurelia using three loci seems to be unambiguous (Fig. 1a–c). Additionally, subsequent species assignment by mating reactions (Table 1) confirms this fact. Furthermore, our results are concordant with previous observations of Sonneborn (1975), who found that strains of P. quadecaurelia do not conjugate witch other
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species of the P. aurelia species complex in contrast to, e.g., P. tetraurelia and P. octaurelia, which can interbreed, but their offspring are sterile (Sonneborn 1975). The above findings strengthen our hypothesis that P. quadecaurelia might differentiate completely from other species of the P. aurelia complex.
Conclusions In the case of the Paramecium aurelia species complex, as well as other ciliate sibling species, strain crosses are the standard way of species identification. For this reason, it is worth noting that the species membership of the new strains of P. quadecaurelia from Ecuador and Thailand was first identified by molecular markers. Subsequently, clear taxonomic interpretation was possible with mating tests, where all of the studied P. quadecaurelia strains reacted (conjugated) with each other, yielding viable hybrids. Given the time-consuming procedures of the above method, molecular data have become an alternative enabling an easy and reliable technique of identification of several ciliate species. This seems particularly important for field biologists, who encounter difficulties if exact and fast taxonomic determination is necessary.
Acknowledgements The authors are greatly indebted to Ms. Tatiana Fokin for collecting water samples in Thailand and to Prof. Sergei I. Fokin, Department of Biology, University of Pisa, Italy for providing the established strains Ta3 and Ta7. This research was partly supported by Grant No. N N303 415636 awarded to S. Tarcz by the Ministry of Science and Higher Education, Warsaw, Poland. Project carried as part of the European Cooperation in Science and Technology (COST BM1102).
Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/ j.ejop.2012.11.001.
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