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edited by H.-D. Gôrtz. Springer Verlag, Berlin, Heidelberg, New York, London, Paris and Tokyo, 1988. Pp. xxii+444. L85. DM248

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J. B. Tucker; Paramecium. J Cell Sci 1 January 1989; 92 (1): 6–7. doi: https://doi.org/10.1242/jcs.92.1.6

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I hope that a good library is included among the Heavenly Facilities because the late Tracy Sonneborn would enjoy reading this book. It is just over half a century since Sonneborn discovered mating types in Paramecium. It was the first demonstration of sex in a unicellular animal and stimulated a wide range of investigative work. Paramecium was rapidly established in a commanding position as ‘top ciliate’ and exploited as a eukaryotic cell model, albeit a tantalizingly peculiar one, for analyses of gene action and cell functions. Paramecium is still number one so far as the overall biology of ciliates is concerned, although Tetrahymena has become pre-eminent in some areas. The book provides a well-coordinated and comprehensive set of reviews that are all of a high standard. It surveys the current state of research with Paramecium and effectively documents what has been achieved in the last 50 years. This volume is a welcome arrival because, as John Preer rightly emphasizes in his Foreword, Paramecium is becoming increasingly valuable as both a model cell and a model organism.

For many cell biologists, ciliates have often seemed too different, and too irrelevant, so far as the mainstream of progress in understanding eukaryotic cell organization is concerned. However, this book demonstrates that analyses of the ‘Paramecium cell’ have entered a new era. The nature of the ‘differences’ are beginning to be established in terms that are familiar to all cell biologists. Most importantly, they are providing fresh insights into a wide range of cell phenomena. Paramecium can now be regarded as a very relevant and useful cell type. For example, extracts of conjugants mimic the in vitro action of maturation-promoting factor by inducing the breakdown of the germinal vesicles at Bufo oocytes (chapter 5), and a soluble calmodulin influences a surface membranebound Ca 2+ -dependent ion channel (chapter 15). The extent to which Paramecium can be exploited as a large eukaryotic cell and subjected to penetrating investigations employing a revealing combination of analytical approaches (such as microinjection, voltage clamping and genetic dissection, as well as the usual array of molecular and ultrastructural techniques) is especially well portrayed in the chapters dealing with ion channel functions and trichocyst exocytosis (chapters 15 and 20).

Advances have been made on an impressively broad front. The 23 main chapters deal with: cytology (Allen), breeding systems (Nyberg), mating-types (Tsukii; Kitamura), conjugation (Fujishima), cell cycle (Berger), nuclear organization (Mikami; Freiburg), aging (Tak-agi), immobilization antigens (Schmidt), mitochondria (Sainsard-Chanet & Cummings), electrophysiology (Machemer), cilia (Machemer; Schultz & Klumpp), ionchannels (Ramanathan, Saimi, Hinrichsen, Burgess-Cassler & Kung), behavioural genetics (Takahashi), chemokinesis (Van Houten & Preston), lysosomes (Fok & Allen), exocytosis (Adoutte), cytoskeleton (Cohen & Beisson), endocytobionts (Gôrtz; Quackenbush) and ecology (Landis). The only obvious omission is a detailed treatment of cortical pattern formation. This might seem odd since it was the famous inverted ciliary rows of Paramecium that did much to attract attention to the topic. However, most of the recent progress has been effected with other ciliates.

Despite a flurry of recent advances, several of the most longstanding and interesting questions remain shrouded in mystery. How is the regular distribution of multiple copies of genetic material effected during macronuclear amitosis? How do killers kill? What is the function of the 8000 or so extrusible trichocysts that occupy up to 10 % of each cell’s total volume? Mating-type substances still elude attempts to isolate them. Judging by progress to date, resolution of these seemingly parochial issues will advance general understanding of evolution, ecology and cell biology.

On the back cover of the book it is claimed that the contents will interest students as well as scientists working on all aspects of cell research. I agree, but with the proviso that the students are advanced and all these potential readers are familiar with the basics of ciliate conjugation and genetics. Such familiarity is particularly necessary for most chapters in the first half of the book where the genetic system and life cycle are substantially involved. Thereafter, one can usually dip straight into individual chapters to browse rewardingly through and Müller have used the freeze-substitution methods, first developed in the early 60s by van Harreveld and Malhotra, to achieve remarkably beautiful preservation of insect sensilla. They discuss these relatively simple methods in chapter 7. Apart from the fear of the unknown, one wonders why this approach has not gained popularity in electron microscopy laboratories! The nine chapters on specialized applications in part III include immunogold labelling, X-ray microanalysis and autoradiography.

This is not a book of recipes but of brief critical reviews written with competence and authority. All the contributors have, of course, extensively reviewed their subjects in other publications; so why publish this volume? A novel feature that distinguishes this collection from its numerous competitors is that, after submitting their manuscripts to the editors, all the authors were interned for three days in Rigberg Castle in Bavaria for a thorough soul-searching exercise induced by mutual criticism and free discussion. This ‘group therapy’ seems to have resulted in much improved versions of their contributions and a uniformity, at least of jargon (given in the Glossary), often lacking in multi-authored volumes, ‘peer refereed’ or not. The book has been very well produced, and the high-quality reproduction of the micrographs is especially commendable. The editors are to be congratulated for doing such a professional job. In my view it is probably the best short volume available on the subject. I hope that it will inspire more biologists to take up cryotechniques for the preparation of their specimens.

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• Published: 27 October 2022

Morphological diversity and molecular phylogeny of five Paramecium bursaria (Alveolata, Ciliophora, Oligohymenophorea) syngens and the identification of their green algal endosymbionts

• Christian Spanner 1 ,
• Tatyana Darienko 1 , 2 ,
• Sabine Filker 3 ,
• Bettina Sonntag 1 &
• Thomas Pröschold 1  

Scientific Reports volume  12 , Article number:  18089 ( 2022 ) Cite this article

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Paramecium bursaria is a mixotrophic ciliate species, which is common in stagnant and slow-flowing, nutrient-rich waters. It is usually found living in symbiosis with zoochlorellae (green algae) of the genera Chlorella or Micractinium . We investigated P. bursaria isolates from around the world, some of which have already been extensively studied in various laboratories, but whose morphological and genetic identity has not yet been completely clarified. Phylogenetic analyses of the SSU and ITS rDNA sequences revealed five highly supported lineages, which corresponded to the syngen and most likely to the biological species assignment. These syngens R1–R5 could also be distinguished by unique synapomorphies in the secondary structures of the SSU and the ITS. Considering these synapomorphies, we could clearly assign the existing GenBank entries of P. bursaria to specific syngens. In addition, we discovered synapomorphies at amino acids of the COI gene for the identification of the syngens. Using the metadata of these entries, most syngens showed a worldwide distribution, however, the syngens R1 and R5 were only found in Europe. From morphology, the syngens did not show any significant deviations. The investigated strains had either Chlorella variabilis , Chlorella vulgaris or Micractinium conductrix as endosymbionts.

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Introduction

Paramecium bursaria has been studied since decades because of its easiness to be kept and experimentally manipulated under manifold cultivation conditions. Some major aspects on this model ciliate were investigated in detail: (i) P. bursaria lives in symbiosis with coccoid green algae belonging to the genera Chlorella and Micractinium (Pröschold et al. 1 and references therein). Advantages of this close relationship include nutritional aspects as the algae provide photosynthetic products and photoprotection to the ciliate 2 , 3 . Accordingly, different aspects such as the process of cell–cell recognition and the symbiont-specificity are of great interest (Fujishima 4 and articles therein). (ii) Complex mating systems in P. bursaria were discovered in mating experiments during (sexual) conjugation processes. So far, six genetic varieties were originally detected by Sonneborn 5 and later designated as syngens 1 to 6, which were considered as biological species 6 , 7 , 8 . Most syngen-types have four (syngens 1 and 3) or eight (syngens 2 and 4–6) mating types 9 . As the strains that Bomford 9 used for his experiments were lost, Greczek-Stachura et al. 10 established a new syngen system, i.e., R1-R5 in principle most likely corresponding to Bomford’s syngens and indicated by a “B” but in a different order (R1–B6, R2–B4, R3–B1, R4–B2, and R5—B3 according to Greczek-Stachura et al. 10 ). Syngen 5 of Bomford 9 was not included. However, the subdivision into syngens was accompanied by phylogenetic analyses of the ITS (internal transcribed spacer regions; partial SSU–ITS1–5.8S–ITS2–partial LSU region) rDNA, the mitochondrial COI (cytochrome oxidase I) and H4 histone genes 10 . The five syngens were recently described as cryptic species based on COI haplotypes and named accordingly as Paramecium primabursaria , P . bibursaria , P . tribursaria , P . tetrabursaria and, P . pentabursaria 11 . Unfortunately, these species were not validly described according to the International Code for Zoological Nomenclature (ICZN), which requires formal descriptions and deposition of holotype specimens to public museums.

Despite such detailed studies in respect to conjugation and endosymbiosis, the morphology and the phenotypic plasticity of P. bursaria has only been rarely investigated. Kreutz et al. 12 compared the morphology and ultrastructure of one P. bursaria strain with another “green” Paramecium , i.e., Paramecium chlorelligerum . However, both species were investigated directly from field samples and the phenotypic plasticity was not studied from cultured material.

The aim of this study was the comparison of 48 P. bursaria strains using an integrative approach to answer the following questions: (i) How many phylogenetic lineages among the investigated strains can be revealed? (ii) Do they correspond to the known syngen affiliations? (iii) Does the morphology of the ciliate strains differ among the syngens? (iv) Do the different syngens show any biogeographic pattern? and, (v) Do all strains bear the same algal endosymbiont? We studied the strains both isolated from diverse geographical regions and acquired from culture collections. First, we sequenced the SSU and ITS rDNA sequences. Subsequently, from each phylogenetic clade, at least one strain was selected to study its morphology and phenotypic plasticity from living and silver-stained specimens. Finally, the green algal endosymbionts were identified both from morphology and a diagnostic PCR approach.

Molecular Phylogeny of Paramecium bursaria and Identification of its Endosymbionts

The SSU and ITS rDNA of the nuclear ribosomal operon were sequenced to infer the genetic variability of the investigated strains. The SSU and ITS rDNA sequences were aligned according to their secondary structure (examples are presented for the strain SAG 27.96; Fig.  1 and Supplementary Fig.  1 ). Additional sequences acquired from GenBank were incorporated into a dataset, which included all syngens also from references known for P. bursaria . The phylogenetic analyses revealed five highly supported lineages among the P. bursaria strains, which corresponded to their syngen assignment. As demonstrated in Fig.  2 , all investigated strains belonging to the syngens R1, R2 and R5 originated from Europe, whereas the others of the syngens R3-R4 showed a worldwide distribution. The three known green algal endosymbionts, i.e., Chlorella variabilis (Cvar), Chlorella vulgaris (Cvul) and Micractinium conductrix (Mcon) showed no or only little affiliation to specific syngens.

ITS‐1 ( A ) and ITS-2 ( B ) secondary structures of Paramecium protobursaria , SAG 27.96 (syngen R1).

Molecular phylogeny of the Paramecium bursaria species complex based on SSU and ITS rDNA sequence comparisons. The phylogenetic tree shown was inferred using the maximum likelihood method based on the datasets (2197 aligned positions of 19 taxa) using the computer program PAUP 4.0a169. For the analyses, the best model was calculated by PAUP 4.0a169. The setting of the best model was given as follows: TVM + I (base frequencies: A 0.2983, C 0.1840, G 0.2271, T 0.2906; rate matrix A–C 2.6501, A–G 8.6851, A–U 5.3270, C–G 0.91732, C–U 8.6851, G–U 1.0000) with the proportion of invariable sites (I = 0.9544). The branches in bold are highly supported in all bootstrap analyses (bootstrap values > 50% calculated with PAUP using the maximum likelihood, neighbour—joining, and maximum parsimony). The clades are named after the syngens (color‐coded) proposed by Greczek‐Stachura et al. 10 and Bomford 9 in brackets. The accession numbers are given after the strain numbers. The endosymbiotic green algae identified are highlighted (Mcon— Micractinium conductrix , Cvar— Chlorella variabilis and Cvul— Chlorella vulgaris ) after the origin of the P. bursaria strains. The reference strain of each syngen is marked with an asterisk. The strains used for morphological comparisons are marked with a green dot next to the strain number.

Synapomorphies of the Paramecium bursaria Syngens

As demonstrated in Fig.  2 , the subdivision of the P. bursaria strains into syngens is supported by the phylogenetic analyses of the SSU and ITS rDNA sequences. To figure out if these splits were also supported by characteristic molecular signatures, we studied the secondary structures of both SSU and ITS of all available sequences. We discovered 30, respectively 23 variable positions among the SSU and ITS sequences (numbers of these positions in the respective alignments are given in Fig.  3 ). All syngens showed characteristic patterns among the SSU and ITS. Only the syngens R1 and R2 could not be distinguished using the SSU only, however, in combination with the ITS, each syngen is characterized by unique synapomorphies as highlighted in yellow (Fig.  3 ). In addition, few variable base positions within syngens (marked in blue in Fig.  3 ) have been recognized in the ITS regions. For comparison with literature data, we also analyzed all available sequences of the mitochondrial COI gene to find synapomorphies for the five syngens. Within this gene, only 18 variable positions at the amino acid level could be discovered of which 13 are diagnostic for the five syngens (Fig.  3 ).

Variable base positions among the SSU, ITS rRNA, and COI sequences of the five syngens among the Paramecium bursaria species complex. The unique synapomorphies are highlighted in yellow, variable positions marked in blue.

The synapomorphies discovered above were used to get insights into the geographical distribution of each P. bursaria syngen. Despite the complete SSU and ITS rDNA sequences included in the phylogeny presented in Fig.  2 , records of the partial SSU or ITS rDNA sequences are available in GenBank (BLASTn search; 100% identity; 13 ). Considering the metadata of our investigated strains and of the entries in GenBank (Supplementary Table 1 ), we constructed three haplotype networks using the Templeton-Crandall-Sing (TCS) approach. The SSU haplotype network (Fig.  4 ) containing 84 records showed that the syngens R1, R2 and R5 were only found in Europe, whereas the other three syngens have been discovered around the world. A similar distribution pattern occurred when using the ITS (101 entries in GenBank). Records of syngens R1 and R5 have only been found in Europe, whereas all other syngens were distributed around the world. The 132 COI records found in GenBank by the BLASTn search were used for the haplotype network, which also showed the similar pattern (Fig.  4 ).

TCS haplotype networks of the five syngens inferred from SSU, ITS rRNA, and COI sequences of the Paramecium bursaria species complex. This network was inferred using the algorithm described by Clement et al. 40 , 41 . Sequence nodes corresponding to samples collected from different geographical regions.

Ciliate Taxonomy

Considering all our findings, P. bursaria is morphologically highly variable, and obviously represents a cryptic species complex (Figs. 5 , 6 ; Supplementary Table 2 ). The known five syngens most likely represent biological species according to Mayr 14 and can be attributed to the cryptic species described by Greczek-Stachura et al. 11 . As mentioned above, the assignments of these cryptic species by Greczek-Stachura et al. 11 have not been validly described according to the ICZN. In addition, the naming using a mixture of Latin prefix and Greek suffix is also not appropriate (the epithet bursa derived from the Greek word byrsa ). Therefore, we describe the five syngens as new species as follows. The general morphological features of these species are summarized in Table 1 .

Ventral views of Paramecium bursaria morphotypes in vivo: P. protobursaria (syngen R1), i.e., strains SAG 2645 ( A ) and PB-25 ( B ); P. deuterobursaria (syngen R2), i.e., strains CCAP 1660/36 ( C ) and CCAP 1660/34 ( D ); P. tritobursaria (syngen R3), i.e., strains CCAP 1660/28 ( E ), CCAP 1660/26 ( F ) and CCAP 1660/31 ( G ); P. tetratobursaria (syngen R4), i.e., strains CCAP 1660/25 ( H ) and CCAP 1660/33 ( I ); P. pentobursaria (syngen R5), i.e., strain CCAP 1660/30 ( J ). Scale bar 20 µm.

Morphological details of the Paramecium bursaria species complex from specimens of strains PB-25 ( A ), CCAP 1660/30 ( B ), SAG 2645 ( C , F , G , I , L – N ), CCAP 1660/36 ( D ), CCAP 1660/26 ( E , H ), CCAP 1660/30 ( J , O ), CCAP 1660/16 ( K ) in vivo ( A – F , H – O ) and after silver nitrate staining ( G ). Adoral membranelles ( A , B ), endosymbiotic algae Micractinium conductrix ( C ), caudal and somatic cilia ( D ), arrows denote excretory pores of the contractile vacuoles: extruded extrusomes are shown and caudal cilia ( E ), ventral views showing the preoral suture and the oral opening ( F ), the ciliary pattern ( G ), arrows denote excretory pores of the contractile vacuoles ( H ), trichocysts and symbiotic algae underneath the pellicula ( I , J ), cell size variations ( K ), radial collecting channels (white arrows) and excretory pores (black arrows) of contractile vacuoles ( L ), macro- and micronucleus ( M ), cytopyge and characteristic rectangular pellicular pattern ( N ), pattern of the pellicula ( O ). AS anterior suture, CC caudal cilia, CP cytopyge (cell after), CV contractile vacuole, EP excretory pore of a contractile vacuole, EX extrusomes, M1–M3 membranelles 1–3, MA macronucleus, MI micronucleus, OO oral opening, S symbiotic algae, SC somatic cilia, SK somatic kineties, UM undulating membrane. Scale bars 10 µm ( A , I ), 20 µm ( B , D – H , J , L – O ), 50 µm ( K ).

Paramecium protobursaria sp. nov.

Synonym: Paramecium primabursaria nom. inval.

Description : The strains SAG 27.96 and PB-25 belong to syngen R1 according to Greczek-Stachura et al. 10 , 11 and differ from other syngens by their SSU and ITS rDNA sequences (MT231333). From morphology, the cells are ellipsoidal to broadly ellipsoidal and dorso-ventrally flattened in vivo. The cells measure 70–164 × 44–65 µm; the single macronucleus is located around mid-cell and measures 25–38 × 11–22 µm; the adjacent single compact micronucleus measures 11–20 × 5–8 µm; the usually two (rarely one) contractile vacuoles, one in the anterior and one in the posterior cell portion have radial collecting channels and 1–3 excretory pores each; the number of ciliary rows/20 µm is 14–22; the length of the caudal cilia is 9–19 µm; the numerous trichocysts located in the cell cortex are 4–6 µm in length. The symbiotic algae belong to M. conductrix ; the larger algae measure 4–7 × 4–7 µm; the smaller algal cells measure 2–5 × 2–5 µm.

Geographic distribution : The investigated strains of syngen R1 were found in Europe: Göttingen, Germany; Lake Mondsee, Austria. In addition, this species has been reported from different places in Europe, Asia and North America (see details in Supplementary Table 1 ).

Reference material : Strain SAG 27.96 and the clonal strain SAG 2645 derived from SAG 27.96 are available at the Culture Collection of Algae (SAG), University of Göttingen, Germany.

Holotype : Two slides (one holotype, one paratype) with protargol-impregnated specimens from the clonal culture SAG 2645, which derived from the reference material SAG 27.96, isolated from the pond of the Old Botanical Garden of the University of Göttingen (Germany), have been deposited in the Oberösterreichisches Landesmuseum at Linz (LI, Austria).

Zoobank Registration LSID : AFD967ED-BC2A-43FD-847E-5DF588BB025C.

Paramecium deuterobursaria sp. nov.

Synonym: Paramecium bibursaria nom. inval.

Description : The strains CCAP 1660/34 and CCAP 1660/36 belong to syngen R2 according to Greczek-Stachura et al. 10 , 11 and differ from other syngens by their SSU and ITS rDNA sequences (OK318487). From morphology, the cells are ellipsoidal to broadly ellipsoidal and dorso-ventrally flattened in vivo. The cells measure 81–167 × 35–83 µm; the single macronucleus is located around mid-cell and measures 24–46 × 10–32 µm; the adjacent single compact micronucleus measures 10–18 × 5–9 µm, no micronucleus seen in live cells of strain CCAP 1660/34; the usually two (rarely one or three) contractile vacuoles, one in the anterior and one in the posterior cell portion have radial collecting channels and 1–3 excretory pores each; the number of ciliary rows/20 µm is 13–22; the length of the caudal cilia is 11–20 µm; the numerous trichocysts located in the cell cortex are 4–6 µm in length. The symbiotic algae belong to M. conductrix ; the larger algae measure 5–7 × 4–7 µm; the smaller algal cells measure 3–5 × 2–5 µm.

Geographic distribution : The investigated strains of syngen R2 were found in Europe: Zurich, Switzerland; Lake Piburg, Austria. In addition, this species has been reported from different places in Europe, Asia and Australia (see details in Supplementary Table 1 ).

Reference material : Strain CCAP 1660/36 is available at the Culture Collection of Algae and Protozoa (CCAP) at the Scottish Association for Marine Science, Oban, Scotland.

Holotype : Two slides (one holotype, one paratype) with protargol-impregnated specimens from the reference material CCAP 1660/36, isolated from Lake Piburg (Tyrol, Austria), have been deposited in the Oberösterreichisches Landesmuseum at Linz (LI, Austria).

Zoobank Registration LSID : D1C20BE6-9A15-4A3D-A7E5-DFC31FF04679.

Paramecium tritobursaria sp. nov.

Synonym: Paramecium tribursaria nom. inval.

Description : The strains CCAP 1660/26, CCAP 1660/28 and CCAP 1660/31 belong to syngen R3 according to Greczek-Stachura et al. 10 , 11 and differ from other syngens by their SSU and ITS rDNA sequences (MT231339). From morphology, the cells are ellipsoidal to broadly ellipsoidal and dorso-ventrally flattened in vivo. The cells measure 80–153 × 49–73 µm; the single macronucleus is located around mid-cell and measures 21–53 × 12–31 µm; the adjacent single compact micronucleus measures 9–17 × 3–6 µm; no micronucleus seen in live cells of strain CCAP 1660/28; the usually two (rarely one or three) contractile vacuoles, one in the anterior and one in the posterior cell portion have radial collecting channels and 1–3 excretory pores each; the number of ciliary rows/20 µm is 12–20; the length of the caudal cilia is 8–19 µm; the numerous trichocysts located in the cell cortex are 4–6 µm in length. The symbiotic algae belong to C. variabilis ; the larger algae measure 4–7 × 3–6 µm; the smaller algal cells measure 3–5 × 2–4 µm.

Geographic distribution : The investigated strains of syngen R3 were found in Europe and Asia: Lake Piburg, Austria; Tokyo, Japan; Khabarovsk region, Amur River, Russia. In addition, this species has been reported from different places in Europe, Asia, North and South America as well as in Australia (see details in Supplementary Table 1 ).

Reference material : Strain CCAP 1660/26 is available at the Culture Collection of Algae and Protozoa (CCAP) at the Scottish Association for Marine Science, Oban, Scotland.

Holotype : Two slides (one holotype, one paratype) with protargol-impregnated specimens from the reference material CCAP 1660/26, isolated from Japan, have been deposited in the Oberösterreichisches Landesmuseum at Linz (LI, Austria).

Zoobank Registration LSID : CC0FBA7E-9E3A-4C37-B424-C9BFF2018EC0.

Paramecium tetratobursaria sp. nov.

Synonym: Paramecium tetrabursaria nom. inval.

Description : The strains CCAP 1660/25 and CCAP 1660/33 belong to syngen R4 according to Greczek-Stachura et al. 10 , 11 and differ from other syngens by their SSU and ITS rDNA sequences (MT231347). From morphology, the cells are ellipsoidal to broadly ellipsoidal and dorso-ventrally flattened in vivo. The cells measure 65–179 × 37–79 µm; the single macronucleus is located around mid-cell and measures 18–53 × 10–29 µm; the adjacent single compact micronucleus measures 8–18 × 4–10 µm; the usually two (rarely one or three) contractile vacuoles, one in the anterior and one in the posterior cell portion have radial collecting channels and 1–3 excretory pores each; the number of ciliary rows/20 µm is 14–19; the length of the caudal cilia is 12–20 µm; the numerous trichocysts located in the cell cortex are 4–7 µm in length. The symbiotic algae belong to C. variabilis (CCAP 1660/25) and M. conductrix (CCAP 1660/33); the larger algae measure 3–6 × 3–6 µm; the smaller algal cells measure 2–5 × 1–4 µm.

Geographic distribution : The investigated strains of syngen R4 are found in North- and South America: Burlington, North Carolina, USA; San Pedro de la Paz, Laguna Grande, Chile. In addition, this species has been reported from Europe (see details in Supplementary Table 1 ).

Reference material : Strain CCAP 1660/25 is available at the Culture Collection of Algae and Protozoa (CCAP) at the Scottish Association for Marine Science, Oban, Scotland.

Holotype : Two slides (one holotype, one paratype) with protargol-impregnated specimens from the reference material CCAP 1660/25, isolated from a pond in Burlington (North Carolina, USA), have been deposited in the Oberösterreichisches Landesmuseum at Linz (LI, Austria).

Zoobank Registration LSID : 78BA9923-07A9-4918-AD7C-9E5E15CC9CDB.

Paramecium pentobursaria sp. nov.

Synonym: Paramecium pentabursaria nom. inval.

Description : The strain CCAP 1660/30 belongs to syngen R5 according to Greczek-Stachura et al. 10 , 11 and differs from other syngens by their SSU and ITS rDNA sequences (MT231348). From morphology, the cells are ellipsoidal to broadly ellipsoidal and dorso-ventrally flattened in vivo. The cells measure 161–194 × 76–99 µm; the single macronucleus is located around mid-cell and measures 24–47 × 19–31 µm; the adjacent single compact micronucleus measures 13–20 × 4–9 µm; the usually two (rarely one or three) contractile vacuoles, one in the anterior and one in the posterior cell portion have radial collecting channels and 1–4 excretory pores each; the number of ciliary rows/20 µm is 13–19; the length of the caudal cilia is 14–25 µm; the numerous trichocysts located in the cell cortex are 5–7 µm in length. The symbiotic algae belong to C. variabilis ; the larger algae measure 5–6 × 5–6 µm; the smaller algal cells measure 4–5 × 3–4 µm.

Geographic distribution : The investigated strain of Syngen R5 was found in Europe: Astrakhan Nature Reserve, Russia.

Reference material : Strain CCAP 1660/30 is available at the Culture Collection of Algae and Protozoa (CCAP) at the Scottish Association for Marine Science, Oban, Scotland.

Holotype : Two slides (one holotype, one paratype) with protargol-impregnated specimens from the reference material CCAP 1660/30, isolated from Astrakhan Nature Reserve (Russia), have been deposited in the Oberösterreichisches Landesmuseum at Linz (LI, Austria).

Zoobank Registration LSID : 6629FA71-E00F-48C6-83AB-61C0CA4823B6.

Syngen Affiliation related to Ciliate Morphology, Endosymbionts and Geographic Distribution

Pearson-correlations of morphometric, syngen-specific and endosymbiont datasets of the P. bursaria strains revealed four significant positive correlations (p < 0.05 and − 0.75 > r > 0.75) between ciliate cell length (BLEN) and width (BWID), BWID and macronucleus width (MACWID), as well as length and width of large symbiotic algae (LSALEN and LSAWID; Fig.  7 ).

Pearson-correlations of morphometric, symbiont and syngen data of Paramecium strains under study. Colored dots indicate the strength of correlation, and the size of dots represent p-values. Bold squares highlight significant correlations, with − 0.75 > r > 0.75 and p < 0.05. Abbreviations: ANVAC number of excretory pores in anterior contractile vacuole, ALSPEC algal species, BLEN body/cell length, BWID body/cell width, CAUCIL caudal cilia length, CILROW number of ciliary rows, EXTLEN extrusome/trichocyst length, GEO geographical region, LSALEN large symbiotic algae length, LSAWID large symbiotic algae width, MACLEN macronucleus length, MACWID macronucleus width, MICLEN micronucleus length, MICWID micronucleus width, POVAC number of excretory pores in posterior contractile vacuole, SSALEN small symbiotic algae length, SSAWID small symbiotic algae width, SYN syngen affiliation.

The results of the principal component analysis (PCA) are summarized in the ordination diagram in Fig.  8 . The first two axes explain 44.4% of the total variation in the investigated features. Only the first five components (out of 18) had eigenvalues > 1, accounting for 73.1% variation in total (Supplementary Table 3 ). Principal component axis 1 (PC1) appears to be most negatively weighted by syngen (SYN) and width of the macronucleus (MACWID), separating CCAP 1660/30 and CCAP 1660/33 from the other strains. Principal component axis 2 (PC2) is primarily positively influenced by symbiotic algae characteristics (LSALEN, LSAWID, small symbiotic algal length (SSALEN) and width (SSAWID)) and, ciliate cell length (BLEN) and width (BWID; Supplementary Table 4 ), partitioning strain PB-25, CCAP 1660/26 and CCAP 1660/36 from CCAP 1660/31 and SAG 27.96 (Fig.  8 ).

PCA of morphometric data of Paramecium bursaria strains. Only the top eight contributing variables are shown.

The redundancy analysis (RDA; Fig.  9 ) revealed a large difference between morphometric features and the tested set of explanatory variables (i.e., algal species (ALSPEC), LSAWID, SSALEN, SYN and GEO) as only 26.9% of the total variation could be explained.

Ordination diagram for redundancy analysis (RDA) of morphometric data and shown syngen (SYN), geographic region (GEO), and algal features (ALSPEC, LSAWID and SSALEN) as explanatory features.

Among strains of P. bursaria , six syngens have been discovered so far by mating experiments 5 , 9 , 10 . Our phylogenetic analyses using a concatenated dataset of SSU and ITS sequences revealed five highly supported lineages among the investigated P. bursaria strains, which clearly corresponded to the cryptic species assigned to syngens R1-R5 according to Greczek-Stachura et al. 10 , 11 . All syngens could be individually distinguished by their molecular signatures (Fig.  3 ), however, isolates belonging to syngens R1 and R2 could not be recognized by sequencing their SSU rDNA only.

Paramecium bursaria are distributed worldwide (Fig.  4 ). Only the syngens R1 and R5 have been found in Europe, whereas the other syngens have been recorded from Europe, Asia, North and South America and Australia. However, very little is known from other regions of the world such as South America, Australia or Africa.

The available strains of P. bursaria were mostly isolated for studying their green algal endosymbionts. Originally, these endosymbionts were differentiated into two groups: an American (or Southern) and a European (or Northern) type 15 , 16 , 17 , 18 , 19 , 20 . Pröschold et al. 1 taxonomically revised both groups and emended the description of the two species C. variabilis and M. conductrix based on the authentic strains (SAG 211–6 and SAG 241.80). Considering both of these strains, Spanner et al. 21 developed an easy diagnostic PCR approach for the isolation and identification of the zoochlorellae living in P. bursaria revealing that both endosymbiotic species were found in almost all syngens (Fig.  2 ). In syngen R3, only C. variabilis was detected. Interestingly, in the strain CCAP 1660/10, belonging to syngen R4, C. vulgaris has been reported 19 . The assignment to syngen R4 of this strain is surprising because this is the only record from Europe. This has to be taken with caution because the assignment of another strain CCAP 1660/11 to syngen R5 by Hoshina & Imamura 19 is incorrect as demonstrated in our study. This strain belongs to syngen R1 (Fig.  2 ; Table S1 ). Unfortunately, this strain is lost and the syngen assignment cannot be proven. Chlorella vulgaris occurred either free-living or as endosymbiont of ciliates such as Euplotes daidaleos , Coleps hirtus , Climacostomum virens and P. bursaria 1 . Unfortunately, those ciliates are neither available in public culture collections nor their molecular datasets in public databases. Consequently, the ciliate host/syngen from which C. vulgaris had originally been isolated remains unknown. Recently, Greczek-Stachura et al. 22 reported that Chlorella sorokiniana , a free-living species from warm-temperate habitats, also occurred in three isolates of P. bursaria collected in Lake Baikal and the Kamchatka region (Asian part of Russia). The investigations were based on the partial nuclear large subunit (LSU) rDNA and chloroplast genes encoding the ribosomal protein L36 ( rpl 36) and translation initiation factor IF-1 ( inf A). Unfortunately, no ITS of these isolates has been sequenced and, accordingly, the assignment to C. sorokiniana is questionable. For example, the Chinese P. bursaria strain Cs2 (R3) bears the “American” type of endosymbiont as demonstrated by Hoshina et al. 16 , which is C. variabilis and not C. sorokiniana 1 . Moreover, two other reports of Greczek-Stachura et al. 22 were probably incorrect: the strains AZ20-1 (according to CCAP 1660/30 in our study; R5) and Yad1-g (R3), both have C. variabilis , and not C. vulgaris as endosymbiont 21 , 23 (this study; see also Supplementary Table 1 ). Both molecular markers (partial LSU, rpl 36- inf A) do not have the diagnostic power for a discrimination of green algal endosymbionts at the species level.

Ciliate descriptions and taxonomic assignments basically require the detailed study of species-specific diagnostic features, relevant literature, and biogeographical aspects 24 . It is consequently necessary that molecular and microscopic approaches are closely linked for a certain population or strain, especially when the ciliate’s ecology is in the focus of a study 25 , 26 . Nevertheless, as molecular approaches are becoming major tools in ciliate ecology, the morphological identification of a ciliate still remains hidden in many cases 27 . Since the first description by Ehrenberg 28 , P. bursaria was often identified only by the presence of green algal endosymbionts despite reported findings of free-living and naturally algal-free individuals 29 . Moreover, the symbiotic algae can be artificially ‘removed’ from P. bursaria for experimental approaches 30 , 31 . Detailed morphological investigations of this species were lacking for a long time under the assumption that all ‘green’ paramecia were assignable to P. bursaria . Kreutz et al. 12 provided a detailed description on a population of P. bursaria and another green congener, P. chlorelligerum , a rare species that was originally established by Kahl 32 . Despite Kalmus 33 already mentioned a high variability of especially the cell shape among Paramecium species, very little is known about their phenotypic plasticity. However, from our detailed morphometric analyses of the studied strains, we can confirm that the morphological features unequivocally revealed P. bursaria and showed that the characteristics tended to be highly variable (Supplementary Table 2 ) as already reported by Foissner et al. 34 in their identification key.

Conclusions

The P. bursaria species complex is widely distributed around the world. As shown, sequencing and analyzing of the SSU and ITS rDNA of isolated samples and strains can clearly assign them to the syngen level. The five lineages revealed by our phylogenetic analyses clearly corresponded to the syngen affiliations. Unfortunately, the syngens could not be identified by morphology only. Further studies are needed to get more insights about the geographical distribution of the P. bursaria species complex and its endosymbionts, which both can be clearly determined using our molecular tools presented here. The usage of diagnostic PCR approach provided an easy method for identification of the green algal endosymbionts.

Origin of the investigated strains and cultivation of ciliates and their endosymbionts

The origin of the investigated P. bursaria strains is summarized in Table 2 . As the respective strains preferred different media, we used modified Bold Basal Medium (3N-BBM + V; medium 26a in Schlösser 35 ) with the addition of 30 ml of soil extract per liter (S/BBM; see Spanner et al. 21 ), modified Woods Hole MBL (WC) medium 36 mixed with Volvic® (V) mineral water, in various concentrations, V/WC 1:1, and V/WC 5:1 v/v. All cultures were maintained at 15–21 °C under a light:dark cycle of 12:12 h (photon flux rate up to 50 μmol m −2  s −1 ). The isolated green algal endosymbionts were cultivated under the same culture conditions in Basal Medium with beef extract (ESFl; medium 1a in Schlösser 37 ).

DNA extraction, PCR and sequencing

Genomic DNA of the P. bursaria strains 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 as described in Spanner et al. 21 . The datasets generated and analyzed during the current study are available in GenBank ( https://www.ncbi.nlm.nih.gov ). The GenBank accession numbers are given in Table 2 .

Identification of the green algal endosymbionts

The green algal endosymbionts were identified using three different approaches: (i) the diagnostic PCR approach 21 , (ii) direct sequencing using the green algal specific primers G500F and G800R as described by Darienko et al. 38 , and (iii) isolation using the method introduced by Spanner et al. 21 and sequencing of the SSU and ITS rDNA with the green algal specific primers. The respective identification method used is given in Table 2 .

Phylogenetic and network analyses

All sequences were aligned to their secondary structures as demonstrated for strain SAG 27.96 (Fig.  1 ; Supplementary Fig.  1 ). The secondary structures were folded using the software mfold 39 , which uses the thermodynamic model (minimal energy) for RNA folding. The visualization of the structures was manually done using the program Illustrator CS5.1 (Adobe Inc.). For the phylogenetic analyses, we calculated the log-likelihood values of 56 models using the automated selection tool implemented in PAUP version 4.0b169 40 to test which evolutionary model fit best for the dataset. The best model according to the Akaike criterion by PAUP was chosen. The settings of the best model were given in the figure legends. The following methods were used for the phylogenetic analyses: distance, maximum parsimony, and maximum likelihood, all included in PAUP version 4.0b169 40 .

The secondary structures of the SSU and ITS rRNA sequences were compared to find genetic synapomorphies, which were used for the construction of haplotype networks. To establish an overview on the distribution of each syngen, the SSU and ITS haplotypes were used for a BLASTn search (100% coverage, > 97% identity; Altschul et al. 13 ). To construct the haplotype networks, we used the Templeton-Crandall-Sing (TCS) network tool 41 , 42 implemented in PopART 43 . The COI sequences presented in Greczek-Stachura et al. 10 , 11 were analyzed to find synapomorphies at the amino acid level.

Morphological investigations of ciliates and endosymbionts

The morphology of the P. bursaria strains and their endosymbionts was studied mainly from living individuals, which were cloned using the isolation method (steps 1 and 2) described in Spanner et al. 21 . After 24 h of starvation, the single ciliate cells were cultivated in 24-well plates (Biomedica) each in the cultivation media mentioned above. To reveal their ciliary pattern, additionally, a dry silver nitrate impregnation was applied 44 . All protists were studied under bright field and differential interference contrast optics with an Olympus BX51 and an Olympus BX60 microscope (Olympus, Vienna, Austria) with 40–1000 × magnifications. For documentation and measurements, two digital image analysis systems were used (ProgRes SpeedXT core 5 2.9.0.1. and ProgRes Capture Pro imaging system version 2.9.0.1, Jenoptik, Jena, Germany). The ciliates were identified by means of the key of Foissner et al. 34 and Kreutz et al. 12 and standard morphometric calculations were done. The green algae were identified by comparison with the descriptions presented in Pröschold et al. 1 . Type slides (holotypes, paratypes) were stained with protargol (Skibbe method) 45 .

Multivariate analyses of morphometric, symbiont and syngen data of Paramecium  strains

All correlation and multivariate analyses were conducted in R version 4.1.1 using the stats and vegan packages. Statistical analyses included all morphometric, syngen and geographic origin information, as well as algal symbiont features of the Paramecium strains under study (Figs. 7 , 8 , 9 ). Strains CCAP 1660/28 and CCAP 1660/34 were excluded from downstream analyses as no micronucleus data (= no micronucleus could be seen in the ciliates) were available.

All data were first checked for normality with a Shapiro–Wilk test and then used to run standard Pearson correlations between each other to rule out any correlations. Correlations were considered significant if p < 0.05 and − 0.75 > r > 0.75. The overall variation in the dataset was summarized with a PCA (unconstrained ordination). The relationship between morphometric features (response variables) and explanatory variables, representing syngen and symbiont features, was summarized using an RDA (constrained ordination) with centered data. Features GEO, LSALEN and SSAWID were removed from analysis due to multicollinearity with SYN, LSAWID and SSALEN, respectively (Supplementary Tables 3 – 4 ). The significance of the observed relationship was tested with a Monte Carlo permutation test using 999 permutations.

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Acknowledgements

The study was funded by the Austrian Science Fund (FWF): P28333-B25. We thank Hans-Dieter Görtz, Renu Gupta, Thomas Posch, Ulrike Scheffel, Ulrike Koll and Monika Summerer for the collection of strains and help in the laboratory. We thank two anonymous reviewers and Alexey Potekhin for constructive comments on an earlier version of our manuscript.

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Spanner, C., Darienko, T., Filker, S. et al. Morphological diversity and molecular phylogeny of five Paramecium bursaria (Alveolata, Ciliophora, Oligohymenophorea) syngens and the identification of their green algal endosymbionts. Sci Rep 12 , 18089 (2022). https://doi.org/10.1038/s41598-022-22284-z

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DOI : https://doi.org/10.1038/s41598-022-22284-z

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Integrative Neuroscience of Paramecium , a “Swimming Neuron”

Author contributions: R.B. designed research; R.B. performed research; R.B. wrote the paper.

Paramecium is a unicellular organism that swims in fresh water by beating thousands of cilia. When it is stimulated (mechanically, chemically, optically, thermally…), it often swims backward then turns and swims forward again. This “avoiding reaction” is triggered by a calcium-based action potential. For this reason, some authors have called Paramecium a “swimming neuron.” This review summarizes current knowledge about the physiological basis of behavior of Paramecium .

Significance Statement

Introduction.

Even the simplest behavior must engage at least a sensory organ, a large part of the nervous system, the body (muscles, skeleton), and the environment. Thus, understanding the biological basis of behavior requires an integrative approach, which remains highly challenging given the complexity of both the nervous and musculoskeletal systems of vertebrates. A fruitful research strategy is to study model organisms that are structurally simpler and have experimental advantages. For example, the biophysical basis of excitability was studied in the giant axon of the squid ( Hodgkin, 1964 ), the molecular basis of learning and memory was studied in Aplysia ( Kandel, 2009 ). A recent model organism to develop integrative approaches to behavior is Caenorhabditis elegans , with its 302 neurons and a known connectome ( Schafer, 2018 ). In C. Elegans , modeling the entire organism and its interaction with the body and environment seems more feasible in principle ( Cohen and Sanders, 2014 ; Cohen and Denham, 2019 ). Nevertheless, even in this more favorable situation, developing functional and empirically valid neuromechanical models of C. elegans remains very challenging. Two other recently introduced model organisms for this type of integrative work are Hydra , which has a few thousand neurons and the advantage of being transparent ( Dupre and Yuste, 2017 ; Wang et al., 2020 ), and jellyfish Aurelia aurita ( Pallasdies et al., 2019 ). Here, I will present a model organism that is significantly simpler as it consists of a single “neuron.”

Paramecium is a single-cell eukaryote, 100–300 μm long depending on species ( Fokin, 2010 ; Fig. 1 ), which has long been a model organism for many aspects of eukaryotic biology ( Wichterman, 1986 ; Görtz, 1988 ). It is a ciliate that has been living in ponds and lakes all over the world for hundreds of millions of years ( Parfrey et al., 2011 ), a fossil has been discovered in a 200 million-year-old piece of amber ( Schönborn et al., 1999 ). Its abundance and large size made it a popular subject of behavioral study in the late 19th century; Jennings described his culture method as follows ( Jennings, 1897 ): “A handful of hay or grass is placed in a jar and covered with hydrant water. In a few weeks the solution of decaying vegetable matter swarms with paramecia.”

Paramecium morphology. A , Scanning electron microscopy image of P. tetraurelia ; scale bar: 10 μm ( Valentine et al., 2012 ). B , Paramecium caudatum ( Jennings, 1899a ), a large species (∼200 μm) with a pointed posterior end. a, anterior end; p, posterior end; g, oral groove; m, mouth; o, oral side; ab, aboral side; cv, contractile vacuole. The drawing also shows food vacuoles and cilia.

Paramecium swims in fresh water by beating its thousands of cilia, and feeds on smaller microorganisms such as bacteria and algae. It is a prey for other microorganisms such as Didinium . As beautifully described by Jennings more than a century ago (1906), Paramecium lives in a rich sensory environment: it finds food by detecting and following chemicals produced by decaying plants and fellow paramecia; it moves toward the water surface by gravitaxis; it avoids obstacles thanks to its mechanosensitivity; it resists water currents by rheotaxis; it avoids bright light; it avoids hot and cold waters; it even communicates chemically. It typically swims in helicoidal paths interrupted by abrupt changes in direction called avoiding reactions, which form the “trial-and-error” basis of its behavior. When an unfavorable condition is met (obstacle, unwanted chemical), the avoiding reaction is triggered ( Fig. 2 A ): Paramecium swims backward for a brief time, then turns and swims forward in a new direction. By this simple mechanism, Paramecium can navigate in crowded multisensory environments.

The avoiding reaction triggered by an action potential. A , Avoiding reaction against an obstacle, as illustrated by Jennings (1906) . B , Action potential in response to a 2-ms current pulse (top), recorded with the hanging droplet method (bottom; from Naitoh et al., 1972 with permission).

This avoiding reaction is triggered by an action potential produced by voltage-gated calcium channels located in the cilia ( Fig. 2 B ; Eckert, 1972 ). These are L-type calcium channels related to the Ca V 1 family found in neurons, heart and muscles of mammals ( Lodh et al., 2016 ). A number of other ionic currents have been identified ( Eckert and Brehm, 1979 ), and genes for many more ionic channels have been found in the genome, often homologs of mammalian channels ( Martinac et al., 2008 ). Sensitivity to various sensory signals is provided by transduction into ionic currents, which may then trigger action potentials. Piezo channels, which convey mechanosensitivity in many species including mammals ( Coste et al., 2010 ) have been identified in the genome. A rhodopsin-like protein has been identified in Paramecium bursaria , a photosensitive species ( Nakaoka et al., 1991 ). In fact, many signaling pathways of neurons have been found in Paramecium , in particular calcium signaling pathways ( Plattner and Verkhratsky, 2018 ), calcium release channels, pumps, calmodulin, centrin, calcineurin, SNARE proteins, cAMP and cGMP-dependent kinases, etc. For this reason, some authors have called Paramecium a “swimming neuron” ( Kung and Saimi, 1985 ).

Many other motile unicellular organisms have rich behavior ( Wan and Jékely, 2020 ) and produce action potentials, including microalgae ( Eckert and Sibaoka, 1968 ; Harz and Hegemann, 1991 ; Taylor, 2009 ), other ciliates such as Stentor ( Wood, 1991 ), other protists such as Actinocoryne contractilis ( Febvre-Chevalier et al., 1986 ) and even bacteria ( Kralj et al., 2011 ; Masi et al., 2015 ). One advantage of Paramecium is its large size, allowing relatively simple electrophysiological recordings ( Naitoh and Eckert, 1972 ; Kulkarni et al., 2020 ). For this reason, there is a rich literature on Paramecium electrophysiology, mostly from the 1960–1980s ( Eckert, 1972 ; Eckert and Brehm, 1979 ). In addition, Paramecium is still an active model organism in genetics, and benefits from many tools such as RNA interference ( Galvani and Sperling, 2002 ); its genome has also been sequenced ( Aury et al., 2006 ; McGrath et al., 2014 ).

I will first give an overview of the behavior of Paramecium , then I will explain how it moves with its body and cilia, and finally I will describe the physiological basis of behavior, with a special focus on the avoiding reaction. Most studies cited in this review were done on two species, P. caudatum and P. aurelia .

The Life of Paramecium

Swimming, feeding, reproducing.

Behavior has been described in detail in articles and books by Jennings and a few contemporary scientists, in the late 19th and early 20th century ( Jensen, 1893 ; Ludloff, 1895 ; Mendelssohn, 1895 ; Jennings, 1897 , 1906 ); these observations would benefit from precise and systematic measurements with modern techniques. Paramecium lives in fresh water in various kinds of habitats, differing in temperature and composition. It swims in spiral paths at ∼1 mm/s by beating its thousands of cilia, revolving around its long axis at about one cycle per second, the oral groove facing the spiral axis ( Fig. 3 A ; Jennings, 1899a ; Bullington, 1930 ). These paths are occasionally interrupted by abrupt changes in direction, which can be preceded by a short period of backward swimming.

Swimming, feeding and reproducing. A , Spiral swimming, with the oral groove facing the spiral axis ( Jennings, 1899a ). B , Thigmotactic Paramecium resting against a fiber ( Jennings, 1897 ). Arrows show water currents produced by oral cilia. C , Two paramecia in conjugation (sexual reproduction; Jennings, 1904 ).

It is often found near the water surface, as it tends to swim against gravity ( Jensen, 1893 ; p. 18). When it hits a solid surface such as glass or wood, it gives the avoiding reaction ( Fig. 2 A ). But when it encounters some fibrous material such as a decaying plant or a piece of cloth, it may stall ( Jennings, 1897 ). This behavior has been termed contact reaction or thigmotaxis ( Fig. 3 B ). It can also occur on properly coated glass ( Iwatsuki et al., 1996 ). The cilia in contact with the object are immobilized, and all the other cilia are quiet or quivering except the oral cilia, which beat strongly. In this situation, Paramecium may feed, for example on bacteria, yeast or algae. Food is brought into its oral groove by powerful cilia, which have different properties from locomotor cilia ( Jung et al., 2014 ; Aubusson-Fleury et al., 2015 ).

A well-fed Paramecium can reproduce by fission every 6 h ( Beisson et al., 2010a ), depending on temperature ( Krenek et al., 2011 ). Without food, Paramecium can survive for several weeks ( Jackson and Berger, 1984 ). Starvation triggers sexual reproduction, where two individuals of opposite mating types attach to each other by the oral side and exchange genetic material ( Fig. 3 C ). In P. aurelia , sexual reproduction can also occur by autogamy (with itself; Beisson et al., 2010b ).

When Paramecium encounters a solid obstacle, it swims backward for a fraction of second, still revolving around its long axis, then the anterior end turns while the posterior end is still ( Fig. 2 A ). This is called the avoiding reaction; it forms the basis of much of its behavior. According to Jennings, the organism always turns toward the same structurally defined side, the “aboral” side (away from the oral groove; Jennings, 1899a ), although systematic measurements are lacking. But since it also revolves along its long axis, from a fixed viewpoint the change in direction may alternate between left and right. Thus, the change in direction may be considered as pseudo-random.

The avoiding reaction is graded ( Fig. 4 ). A weak stimulus may only trigger a gentle reorientation with no backward swimming ( Fig. 4 A ), while a stronger stimulus induces backward swimming and reorientation ( Fig. 4 B ). A very strong stimulus may trigger long backward swimming followed by turning a complete circle ( Fig. 4 C ). This graded reaction parallels the graded action potential: the duration of backward swimming correlates with the stimulus-induced depolarization ( Machemer and Eckert, 1973 ).

The avoiding reaction is graded ( Jennings, 1904 ): swinging of the anterior end in a weak reaction ( A ), a strong reaction ( B ) and a very strong reaction ( C ).

Paramecium also reacts when the rear is touched, but in a different way ( Fig. 5 A ): it swims forward faster, by beating its cilia up to twice faster ( Machemer, 1974 ). This speed increase is accompanied by a contraction along the longitudinal axis ( Nakaoka and Machemer, 1990 ). This is called the escape reaction, first described by Roesle in 1903 ( Roesle, 1903 ), then by Jennings ( Jennings, 1904 ). Non-localized mechanical stimulation, as when shaking a tube of Paramecium culture, also induces an increase in swimming speed that can last for several minutes.

Paramecium navigation. A , Escape reaction triggered by a heat stimulus (laser) near the posterior end (star; Hamel et al., 2011 ). B , Sideways jumping from a strong heat stimulus (star) by throwing trichocysts ( Hamel et al., 2011 ). C , Trajectory of Paramecium in a 5-mm capillary, showing an increase in backward swimming after 1 min, corresponding to ∼40 avoiding reactions ( Kunita et al., 2014 ). D , Bending of P. caudatum in a 160-μm channel ( Jana et al., 2015 ).

When stimulated by a strong heat using a laser (5–10°C increase), Paramecium can jump away from the stimulus (possibly sideways) within just 5 ms, at ∼10 mm/s ( Hamel et al., 2011 ; Fig. 5 B ). To perform this feat, Paramecium throws trichocysts, which are sorts of needles docked near the membrane, thereby projecting itself in the opposite direction. The same behavior occurs in reaction to an appropriate chemical stimulus and in encounters with the predator Dileptus ( Knoll et al., 1991 ).

When Paramecium swims in a narrow channel that does not allow it to turn, it may be trapped into a dead end, where it will give the avoiding reaction repeatedly, alternatively moving backward and forward against the wall ( Kunita et al., 2014 ). But after a minute, the avoiding reaction suddenly becomes much longer (several millimeters), potentially allowing the organism to escape ( Fig. 5 C ). When the channel is very narrow, Paramecium may also bend itself to move forward ( Jana et al., 2015 ; Smith, 1908 ; Fig. 5 D ). The posterior end anchors onto the wall, presumably because tail cilia do not beat ( Machemer and Machemer-Röhnisch, 1984 ; Ishikawa and Hota, 2006 ), while the anterior end slides along the other wall, causing the cell to bend until it can swim freely. Under some conditions, Paramecium can also slide along surfaces ( Li and Ardekani, 2014 ; Nishigami et al., 2018 ; Ohmura et al., 2018 ). Some of this behavior is due not to physiological responses but to hydrodynamic interactions with surfaces ( Berke et al., 2008 ; Lauga and Powers, 2009 ; Li and Ardekani, 2014 ; Ohmura et al., 2018 ).

Finally, in a water current, Paramecium orients itself with its anterior end directed up stream, a behavior called rheotaxis. According to Jennings (1906) , rheotaxis derives from the avoiding reaction. When Paramecium swims along the water current, its cilia beat backwards and the water current opposes that movement. This acts as a mechanical stimulus which triggers the avoidance reaction. By trial and error, Paramecium turns until it faces the current. However, this remains an untested hypothesis. In a few other microorganisms, rheotaxis has been attributed to hydrodynamic effects ( Bretherton and Rothschild, 1961 ; Marcos et al., 2012 ).

Chemical sensing and social behavior

Paramecium is sensitive to a variety of chemical compounds ( Jennings, 1899b ; Nakatani, 1968 ; Dryl, 1973 ; Valentine et al., 2008 ). It is attracted by some substances, in particular bacterial metabolites (folate, acetate, glutamate, cAMP, biotin, ammonium, etc.), weak acids, carbon dioxide, colloidal solutions. These substances may indicate the distal presence of food, possibly components of the “phycosphere,” the rich interface between phytoplankton and bacteria ( Seymour et al., 2017 ).

Other substances are repellent (e.g., alkaline solutions, quinine, ATP, GTP, GDP, NBT, Alcian Blue, Cibacron blue, Cytochrome c; Francis and Hennessey, 1995 ). Some of these molecules may signal the distal presence of a noxious condition. For example, Hennessey speculated that ATP and GTP are strong repellents because they are “blood-in-the-water signals” ( Hennessey, 2005 ): these molecules are present at high concentrations in cells, and so their presence signals cell lysis, and whatever dangerous condition might have caused it.

Some substances only produce a reaction when Paramecium is subject to toxic doses (cane sugar, dextrose, urea), effectively killing it ( Jennings, 1899b ). For example, after some time, cane sugar induces plasmolysis, and then Paramecium begins to swim backward and forward repeatedly, possibly because of the induced depolarization. But many substances are toxic at doses much larger than the sensitivity threshold. In a number of cases, this sensitivity is conferred by specific membrane receptors, which can depolarize or hyperpolarize the cell ( Van Houten, 1998 ) and possibly modulate ionic channels ( Oami, 1996a , b ).

In the 19th century, Jennings described the behavior of paramecia gathering in a drop of weak acid ( Fig. 6 A ). He linked this behavior to the avoiding reaction. When Paramecium enters a drop of acid, its course is unchanged; but when it reaches the border of the drop, it gives the avoiding reaction and therefore remains in the drop ( Fig. 6 B ). On the contrary, alkaline solutions are repellent: an avoiding reaction is triggered as soon as the alkaline solution is reached ( Fig. 6 C ). More recently, various substances have been characterized as attractant or repellent based on the accumulation of paramecia in a test solution relative to a control solution, using different behavioral assays ( Van Houten et al., 1975 ; Leick and Helle, 1983 ; Levandowsky et al., 1984 ; Nakazato and Naitoh, 1993 ; Valentine and Van Houten, 2016 ).

Chemotaxis and social behavior. A , Gathering of paramecia in a drop of weakly acid solution ( Jennings, 1899a ). B , Path followed by Paramecium in a drop of acid ( Jennings, 1906 ). C , paramecia avoiding a drop of sodium carbonate ( Jennings, 1899a ). D , paramecia gathering in a cloud of carbon dioxide generated by their respiration ( Jennings, 1899a ).

As previously mentioned, when stimulated, Paramecium turns to a structurally defined side (the aboral side, away from the mouth). Therefore, Paramecium is not attracted to a substance because it turns toward it. Rather, its behavior seems to result from trial and error: if attractant concentration increases, then Paramecium keeps on swimming in the same direction; if it decreases, then Paramecium changes direction. Jennings reported that the reaction of the organism is independent of where the chemical substance is applied; however, this may well depend on the compound because some chemoreceptors are spatially organized ( Preston and Van Houten, 1987 ; Oami, 1996b ).

For this reason, this behavior is sometimes named chemokinesis (changes in motility with chemical signals), which is more general than chemotaxis (movements toward a chemical stimulus; Houten, 1979 ). In particular, chemokinesis can result not only from modulation of the avoiding reaction (named klinokinesis), but also of swimming speed (named orthokinesis; Houten, 1978 ). Nonetheless, the chemical modulation of this apparently random motion can lead to motion toward the chemical source, and presumably to a preferred orientation of the body in the direction of the source (since the organism spends more time in the favored direction). There is some similarity with run-and-tumble chemotaxis in bacteria for which there is a dense literature ( Berg, 2008 ; Sourjik and Wingreen, 2012 ), including theoretical ( Berg and Purcell, 1977 ; Kollmann et al., 2005 ; Tu et al., 2008 ; Celani and Vergassola, 2010 ; Tu, 2013 ), and with pirouettes in C. elegans ( Pierce-Shimomura et al., 1999 ).

A consequence of Paramecium attraction to weak acids is social behavior, as observed by Jennings ( Jennings, 1897 ). By their respiration, Paramecium produces CO 2 , which is acid in solutions. At low concentration, Paramecium is attracted to CO 2 . It follows that paramecia tend to attract each other ( Fig. 6 D ). This explains why gatherings can be observed at the bottom of a watch glass or at random positions in a tube. This may play an important role in feeding behavior, as it allows paramecia to collectively search for food.

Finally, Paramecium also has GABA A and GABA B receptors that can influence its behavior ( Bucci et al., 2005 ; Ramoino et al., 2003 , 2004 ). For example, the activation of GABA B receptors inhibits the avoiding reaction. In addition, Paramecium releases GABA on stimulation. This release might act as a signal for other paramecia, or perhaps as an externalized spatial memory for exploration (as observed in slime mold; Reid et al., 2012 ), making the organism take a different action when it comes back to the same location. NMDA-like receptors have also been identified ( Ramoino et al., 2014 ).

The logic of Paramecium behavior

Many aspects of Paramecium behavior can be described as trial and error (1906). If its path is blocked by an obstacle, Paramecium withdraws then tries a new direction. If it encounters an undesirable chemical signal, it changes direction. If it leaves a desirable region, it withdraws and tries a new direction. This logic also applies to other sensory modalities. For example, when placed in a gradient of temperature, Paramecium accumulates in regions with temperature close to their culture temperature ( Mendelssohn, 1895 ; Jennings, 1906 ). Again, this occurs by temperature-triggered avoiding reactions. When temperature changes away from culture temperature (whether this corresponds to a decrease or an increase), the avoiding reaction rate transiently increases; conversely, the avoiding reaction rate decreases when temperature gets closer to culture temperature ( Nakaoka and Oosawa, 1977 ). This behavior is mediated by membrane potential changes ( Tominaga and Naitoh, 1992 ) produced by cold-sensitive and heat-sensitive thermoreceptors ( Tominaga and Naitoh, 1994 ; Kuriu et al., 1996 , 1998 ).

Paramecium also shows photophobic responses to large changes in the intensity of visible light (mainly green, and red; Iwatsuki and Naitoh, 1982 , 1983a , b ; Hinrichsen and Peters, 2013 ). When Paramecium is kept in the dark and a bright light is turned on, it displays the avoiding reaction with a latency of around a second, then adapts over ∼15 s. As a result, Paramecium tends to accumulate in shaded regions. A related species, P. bursaria , is naturally highly sensitive to light and accumulates in lighted regions ( Saji and Oosawa, 1974 ). This species harbors a symbiotic green alga named Chlorella : the alga provides photosynthetic products to its host while the host brings the alga in suitable light conditions. A moderate decrease in light intensity triggers an avoiding reaction, which makes P. bursaria seek light.

This trial-and-error behavior shares some similarity with the run-and-tumble behavior of bacteria ( Berg, 1975 ). Macroscopically, trajectories of Escherichia coli resemble Paramecium trajectories, with helicoidal “runs” interrupted by “tumbles” where the cell changes direction randomly. Bacterial chemotaxis is enabled by concentration-dependent modulation of the tumbling rate: the tumbling rate decreases when concentration increases, while it is unchanged when concentration decreases. Thus, tumbling is not an avoiding reaction (it is not triggered by a concentration decrease). In Paramecium , the new direction is somewhat (pseudo-)random, but the turning event seems more deterministically related to environmental conditions than in bacteria. In other words, the avoiding reaction of Paramecium is more akin to a decision based on sensory inputs, than to a modulation of the spontaneous turning rate. This difference with bacteria may be because of a difference in scale: compared with bacteria, the membrane surface of Paramecium is at least two orders of magnitude larger, so that the signal-to-noise ratio is at least one order of magnitude larger; membrane potential fluctuations are ∼1–3 mV ( Moolenaar et al., 1976 ; Nakaoka et al., 2009 ).

This simple logic of behavior calls for a couple of remarks in the context of neuroscience. First, it is somewhat surprising that a single spiking “neuron” can control relatively complex navigation in crowded multisensory environments, social behavior, and perhaps spatial memory. In terms of connectionism ( Seung, 2012 ), Paramecium is a zero-connectome organism, and yet it can accomplish a variety of ecologically relevant tasks. This arises not from the complexity of the cell, which is electrically much simpler than a single pyramidal cortical neuron (it is isopotential), but rather from the interaction between this spiking cell and the environment, together with the exploratory properties conferred by the pseudo-random nature of the effect of a spike. This highlights the importance of embodiment and coupling with the environment, which are increasingly appreciated in cognitive science and philosophy of mind ( Maturana and Varela, 1973 ; Powers, 1973 ; Gibson, 1979 ; Brooks, 1991 ; Bickhard and Terveen, 1996 ; Hurley, 2001 ; O’Regan and Noë, 2001 ; Ahissar and Assa, 2016 ; Pezzulo and Cisek, 2016 ; Brette, 2019 ).

Second, “control” may not be the right term to describe the relation between spiking and behavior. Motor control is classically described as feedforward or feedback ( Wolpert and Ghahramani, 2000 ). In feedforward control, internal models are used to plan movements, and specific sets of neurons are recruited to trigger the appropriate movements. In Paramecium , spiking produces a single type of movement, regardless of the goal or stimulus: it does not move by planning specific movements. In feedback control, actions are taken that reduce the difference between the observed state and a desired state. In Paramecium , an action is also taken when the observed state is undesirable, but that action is not directed toward the goal, rather, it is (pseudo-)random. Thus, Paramecium movements are based neither on feedforward nor on feedback control, but rather on exploration and selection (trial and error). This is reminiscent of the Darwinian insight that an apparently goal-directed process can occur through random exploration and elimination of unsuccessful choices, rather than by either planning or steering.

Paramecium lives in habitats of diverse ionic composition. Changes in ionic composition directly affect ionic currents and reversal potentials, and therefore can potentially interfere with behavior. For example, moderate changes in cation concentration can alter swimming velocity ( Machemer, 1989 ; Nakaoka et al., 1983 ). More critically, an increase in potassium concentration can inhibit the avoiding reaction through depolarization-induced inactivation of the ciliary calcium channels ( Oka and Nakaoka, 1989 ), making the organism unresponsive to stimulation. Remarkably, after a couple of hours, behavior returns to its normal state before the medium changed ( Oka et al., 1986 ; Fig. 7 A ). In parallel, the resting potential changes after a medium change then decays back to its original value ( Fig. 7 B ). This homeostatic regulation appears to be mediated by changes in channel permeability. With a more prolonged (48 h) exposure to a high potassium solution, more complex changes in excitability can occur, with enhanced responses to Mg 2+ and Na + ( Preston and Hammond, 1998 ).

Adaptation. A , Change in swimming velocity when Paramecium adapted to a solution of 0.25 m m CaCl 2 and 4 m m KCl is transferred to a solution of 0.25 m m CaCl 2 and 1 m m (open circles), 2 m m (closed circles), 4 m m (squares), or 16 m m (triangles) KCl (from Oka et al., 1986 ). B , Resting potential versus KCl concentration for cells adapted to 2 m m , 4 m m , 8 m m , and 16 m m KCl (top to bottom; Oka et al., 1986 , with permission). Arrows indicate the adapted state. C , Accumulation of Paramecium in a warm region ( Jennings, 1899a ). The top of the slide is placed on a 40°C bath while the bottom rests on ice. D , Change in avoiding reaction rate after paramecia cultured at 25°C are transferred to 30°C (from Nakaoka et al., 1982 , with permission). Note the change in time scale.

Temperature also affects ionic channel properties and the entire metabolism of the organism, as well as hydrodynamic properties (viscosity of water). For example, when temperature is lowered, the ciliary calcium current is smaller and slower, action potentials are smaller and broader, cilia reverse with longer latency and for a longer time ( Machemer, 1974 ). As previously discussed, Paramecium has a thermoregulation mechanism based on movement: using the avoiding reaction, it navigates toward waters of a preferred temperature ( Fig. 7 C ). However, this mechanism is insufficient if the medium changes temperature globally. Remarkably, in this case, Paramecium adapts over a couple of hours: behavior returns to normal and the new temperature becomes the preferred temperature ( Nakaoka et al., 1982 ; Fig. 7 D ). This behavioral adaptation correlates with changes in electrophysiological properties, in particular of the ciliary calcium conductance ( Martinac and Machemer, 1984 ).

Beyond adaptation, there is an important literature on learning in Paramecium and other ciliates. Unfortunately, as reviewed by Applewhite (1979) , many of those studies are difficult to interpret as they lack appropriate controls or observations. In a series of papers ( Gelber, 1952 , 1956 , 1957 , 1958 , 1962a , b ), Gelber showed an apparent reinforcement of behavior with a food reward (see ( Gershman et al., 2021 ) for a recent commentary). A platinum wire is lowered repeatedly into a depression slide with paramecia. If the wire is intermittently baited with bacteria, then more and more paramecia cling to the wire, even when a clean wire is finally lowered into the slide. What might be the stimulus? Gelber (1956) noted that the behavior was not observed when paramecia were tested in the dark, suggesting that perhaps paramecia, with permission developed an attraction to a reflection or shadow cast by the wire.

These observations were controversial, because it was objected that lowering the baited wire introduces bacteria in the fluid, to which paramecia are then attracted even when the wire is removed or cleaned ( Jensen, 1957 ). In support of this interpretation, Katz and Deterline (1958) replicated Gelber’s main findings but found that stirring before the final test destroyed the observed behavior. Naturally, this could be interpreted as an erasure of learning because of the mechanical disturbance, but perhaps more crucially, they found that Gelber’s observations could be reproduced when the entire experiment (not just the test) was done in the dark, effectively removing any distal sensory stimulus by which paramecia may be able to recognize the wire. A plausible explanation, in line with informal observations reported in this set of studies, is that feeding reduces the activity of paramecia so that they tend to stay near the wire, and promotes thigmotaxis so that they tend to adhere more easily to the wire. In this case, the procedure would indeed reinforce a behavior, namely the feeding behavior, but not a stimulus-specific behavior. More detailed observations seem necessary to understand the phenomenon.

Another phenomenon that has attracted some attention is tube escape learning, first described by French in 1940 ( French, 1940 ). A single Paramecium is placed in a drop and a thin tube is lowered into it. The organism is drawn into the tube by capillarity. It then escapes from the bottom after ∼30 s. When the experiment is repeated, escape time decreases to around 15 s after a few trials. French states that after the initial trials, paramecia go and back and forth in the tube only a few times then take “one long dive to the bottom.” The faster escape persists for at least 2 h ( Huber et al., 1974 ), which seems to rule out the possibility that Paramecium simply adapts to the mechanical stimulus of capillary suction. This phenomenon has been robustly reproduced by several authors ( Hanzel and Rucker, 1972 ; Applewhite and Gardner, 1973 ), but its basis is unclear. Applewhite and Gardner (1973) proposed that Paramecium released some substance in the tube that then influences future behavior, but this hypothesis contradicts earlier results by Hanzel and Rucker (1972) showing the same performance improvement in multiple paramecia with the same tube. Studies of tube escape learning in Stentor , another ciliate, suggest that the phenomenon is related to gravitaxis ( Bennett and Francis, 1972 ; Hinkle and Wood, 1994 ). Performance improvement is seen only when the tube is vertical, not when it is horizontal, where escape is fast from the first trial. This suggests the following (speculative) explanation: in a vertical tube, paramecia are trapped near the top because of negative gravitaxis, then prolonged confinement (perhaps signaled by frequent avoiding reactions) inhibits the normal gravitactic behavior, so that the organism can escape to the bottom.

Finally, Hennessey et al. (1979) managed to train Paramecium to react to sounds. When a tone is played by a speaker below the slide, Paramecium shows no reaction. However, when the tone is paired with electrical stimulation triggered in the middle of the tone, Paramecium reacts to the stimulus with an avoiding reaction, then after a few trials gives an avoiding reaction at the onset of the tone, in anticipation of the electrical stimulus. The authors demonstrate extinction (reaction disappears when sound is presented alone), retention and specificity (reacting specifically to a 300-Hz tone or to a 500-Hz tone). The physiological basis is not known.

Armus and colleagues ( Armus et al., 2006a , b ; Mingee and Armus, 2009 ) trained paramecia to go to a lighted region. The bath is split into two compartments, one in the dark, the other one in light. Initially, Paramecium spends more time in the dark compartment, because of photophobia. Training consists in electrically stimulating the cell when it enters the compartment of the cathode. After training, Paramecium spends more time than before in the cathodal half, which now only differs by its lighting. However, if stimulation is triggered in the anodal half, then after training Paramecium spends less time in that half. Therefore, the phenomenon does not seem to be based on an association between the electrical stimulus and the light stimulus. A plausible interpretation is the following. As is known from studies of galvanotaxis ( Ludloff, 1895 ; Dale, 1901 ), electrical stimulation makes Paramecium move toward the cathode. Stimulation in the lighted cathodal compartment then makes Paramecium spend more time in light, which results in adaptation of the photophobic behavior. Thus, after training, Paramecium spends more time than before in the lighted compartment. This interpretation is supported by the observation that the “trained” behavior only occurs when the cathodal compartment is lighted during training ( Alipour et al., 2018 ), and by the absence of retention ( Mingee, 2013 ).

In summary, although the existing literature is complex, there is clear evidence of behavioral plasticity in Paramecium . Some can be categorized as adaptation, and there is at least one documented case of learning ( Hennessey et al., 1979 ), understood as a persistent stimulus-specific change in behavior.

The Motor System of Paramecium

How paramecium swims.

In the absence of any stimulus, Paramecium swims in spirals. Paramecium is covered by several thousand cilia ( Fig. 8 A ; ∼4000 cilia in Paramecium tetraurelia ; Aubusson-Fleury et al., 2015 ; for precise counts and spatial pattern, see Iftode et al., 1989 ), each ∼10 μm long and 0.2 μm thick ( Eckert and Naitoh, 1970 ), similar to other motile cilia of eukaryotes, including mammals ( Ishikawa, 2017 ). In forward swimming, each cilium beats at 10–20 Hz ( Fig. 8 B ), with a power stroke toward the right and rear on the visible surface ( Fig. 8 C ). Thus, on the hidden surface (further from the observer), cilia beat toward the left and rear. This results in a forward movement with a rotation around the longitudinal axis, as in unscrewing (over to the left; Fig. 8 D ). The typical velocity is ∼1 mm/s ( Machemer et al., 1991 ).

Spiral swimming. A , Organization of ciliary basal bodies on the oral (ventral) and aboral (dorsal) side (from Iftode et al., 1989 , with permission). B , Ciliary beat cycle: power stroke (or effective stroke) and recovery stroke ( Omori et al., 2020 ). C , Water currents produced by cilia for different orientations of Paramecium ( Jennings, 1904 ). In the oral groove, currents are oriented toward the mouth. D , Metachronal waves represented by parallel lines, progressing transversally, with cilia’s power stroke oriented toward the right and rear (from Machemer, 1972 , with permission). Cilia on parallel lines are at the same phase of the beat cycle. The curved arrow shows the direction of movement.

The spiral is wider than the cell’s width, as first described by Jennings (1901) and later by Bullington (1930) . A possible reason is that cilia in the oral groove beat in a specific direction, toward the mouth, which counters the movement produced by the other cilia. A recent study has shown indeed that properties of oral cilia differ from other cilia ( Jung et al., 2014 ). This may explain why the trajectory describes a wide spiral, with the oral side always facing its axis ( Fig. 3 A ; Párducz, 1967 ).

Properties of spiral swimming can vary, in particular its speed and width. Paramecium can also swim backward, with an effective stroke toward the front and slightly to the right. Thus, in backward swimming, the movement is not the symmetrical of forward swimming: the cell still rotates in the same direction.

Cilia beating is coordinated over the cell in the form of metachronal waves , which progress over the surface at ∼1 μm/ms ( Párducz, 1967 ; Fig. 8 D ). These waves encircle the body in spirals ( Párducz, 1967 ; Machemer, 1969 ). Cilia beat against the direction of the wave, but not at 180°, a pattern called “dexio-antiplectic.” This particular kind of motor coordination is functionally important. A key characteristic of swimming microorganisms is they live at low Reynolds number (R ≈ 0.1 for Paramecium ; Purcell, 1977 ), that is, inertial forces are small compared with viscous forces (as if a human were trying to swim in honey). As a consequence, the swimmer stops as soon as cilia stop beating. Therefore, if cilia beating were synchronized over the entire body, then the swimmer would move forward in regular discontinuous steps. In fact, this can happen in the escape reaction: a strong heat stimulus near the posterior end induces a synchronous power stroke of the cilia (as in the butterfly stroke; Hamel et al., 2011 ), which results in a transient speed increase immediately followed by an almost complete stop, before the metachronal pattern is reestablished. If on the contrary cilia beating were completely disorganized (which can happen transiently in the avoiding reaction), then neighboring cilia might beat in inconsistent directions and this is not an efficient way of swimming. In fact, it has been shown that the metachronal pattern optimizes the energetic efficiency of swimming ( Gueron and Levit-Gurevich, 1999 ; Osterman and Vilfan, 2011 ).

It was once postulated that ciliary coordination might be electrically controlled by the cell, but Paramecium is essentially isopotential ( Eckert and Naitoh, 1970 ). Instead, cilia coordination is mediated by hydrodynamic interactions ( Machemer, 1972 ; Guirao and Joanny, 2007 ) and mechanical coupling through the compliant body ( Narematsu et al., 2015 ), in the absence of any central agency. This illustrates the concept of embodiment in motor neuroscience: part of the problem of efficient coordination is solved not by manipulating body representations, but by direct physical interaction of the body with its immediate environment ( Tytell et al., 2011 ). In the case of microorganisms such as Paramecium , the results of this physical interaction can be understood precisely, thanks to an abundant literature on the mechanics of cilia and flagella ( Blake and Sleigh, 1974 ; Sartori et al., 2016 ; Wan, 2018 ) including mathematical models ( Dillon et al., 2007 ; Yang et al., 2008 ), as well as on the hydrodynamics of swimming microorganisms ( Keller and Wu, 1977 ; Lauga and Powers, 2009 ; Jung et al., 2014 ).

How Paramecium moves upward

As many other microorganisms ( Häder and Hemmersbach, 2018 ), Paramecium tends to aggregate near the water surface, despite the fact that it is slightly heavier than water (∼4%), a puzzling phenomenon which has attracted an abundant literature, first described in detail by Jensen in 1893 ( Jensen, 1893 ). When observed in a vertical plane, trajectories are curved upward ( Roberts, 2010 ; Fig. 9 A ). The earliest explanation, the gravity-buoyancy model, postulates a mismatch between the buoyancy center and the gravity center ( Verworn, 1889 ): this could generate a torque making the body align with gravity. Roberts ( Roberts, 1970 , 2010 ) argued that density inhomogeneities are unlikely to be sufficient to account for the observations, and instead proposed a drag-gravity model: as the posterior end is larger than the anterior end, the viscous drag differs and the posterior end falls more rapidly than the anterior end; thus, the cell turns upward. However, Jensen (1893) and later Kuznicki (1968) observed that dead or immobilized cells fall with no preferred orientation, although this is questioned by Roberts ( Roberts, 1970 ). This would discard both passive orientation mechanisms. The propulsion-gravity model ( Winet and Jahn, 1974 ) is a more complex proposition, which links gravitaxis with ciliary beating: sedimentation introduces viscous resistance to beating that is stronger in the up phase of the helicoidal cycle than in the down phase, resulting in velocity-dependent reorientation.

Gravitactic behavior of Paramecium . A , Upwardly curved trajectories of Paramecium in a vertical chamber (from Roberts, 2010 , with permission). B , Velocity change (corrected for sedimentation) as a function of cell orientation (from Nagel and Machemer, 2000 , with permission), open circles correspond to a morphologic mutant. C , Avoiding reaction frequency as a function of acceleration in a centrifuge microscope, after 4 h of equilibration (from Nagel and Machemer, 2000 , with permission). Triangles indicate cell direction.

In addition to these hydrodynamic mechanisms, physiological mechanisms have been postulated. It has been observed that Paramecium swims slightly faster upwards than downwards, once sedimentation has been subtracted ( Machemer et al., 1991 ; Ooya et al., 1992 ; Fig. 8 B ), and the avoiding reaction is triggered more often when it swims backwards than upwards, although this bias tends to disappear after some time ( Nagel and Machemer, 2000 ; Fig. 8 C ). Although spurious correlations should be ruled out (e.g., cells that swim more slowly may tend to fall), Machemer and colleagues have proposed that this is because of pressure differences between the top and bottom ends of the cell, which are sensed by mechanoreceptors. As there is a spatial gradient of mechanosensitivity between the front and rear, the transduced current would be hyperpolarizing when the anterior end is upward (increased pressure on the rear end) and depolarizing when the anterior end is downward. In support of this hypothesis, a cell vertically immobilized between two horizontal electrodes can spontaneously turn upward or downward, and small membrane potential changes with the expected sign are observed, although with long latency (on the order of 20 s; Gebauer et al., 1999 ). These physiologically induced changes in mean velocity and avoiding reaction rate likely represent a small contribution to gravitaxis, compared with the reorientation of the cell ( Roberts, 2010 ), but it is conceivable that reorientation itself occurs by physiological modulation of velocity within the helicoidal cycle ( Mogami and Baba, 1998 ).

How Paramecium turns

In the avoiding reaction, Paramecium swims backward (if the reaction is strong) then turns before it swims forward again. Backward swimming occurs because cilia reorient, with the power stroke oriented toward the anterior end instead of the posterior end, but how can Paramecium turn? Turning requires some inhomogeneity in the ciliary beating pattern.

First, anterior and posterior cilia do not revert synchronously during the avoiding reaction ( Fig. 10 A ; Párducz, 1967 ; Machemer, 1969 ). When the avoiding reaction is initiated, all cilia simultaneously strike forward, which moves the cell backward (2). The beating pattern then progressively reorganizes into the metachronal pattern as the cell swims backward (3–5). Reorientation of the cell starts when the anterior end reverts to the forward metachronal pattern (6–8). Thus, anterior and posterior ends show different metachronal patterns, respectively, of forward and backward swimming.

Details of the avoiding reaction. A , Reorganization of the ciliary beating pattern during the avoiding reaction (after Machemer, 1969 ). B , Cross-section of Paramecium seen from the anterior end, during forward swimming ( a , corresponding to step 1) and during reorientation ( b , corresponding to step 6), according to Jennings (1904) . The arrows correspond to the induced movement of the body (opposite to the beating direction).

It is not obvious, however, how this asynchronous pattern would make the cell turn. If the beating pattern were axisymmetric, then the net force produced by either group of cilia (anterior or posterior) should be directed along the main axis. Jennings claims that cilia in the oral groove may also reverse, i.e., they expel fluid from the mouth ( Jennings, 1899a ; Fig. 8 C ). This could make Paramecium turn toward its aboral (dorsal) side, as observed, but Jennings and Jamieson observed that when Paramecium was cut in two pieces below the oral groove, both pieces could turn in a similar way ( Jennings and Jamieson, 1902 ). Jennings also mentions that cilia of the anterior end do not all strike to the right: instead, they strike toward the oral groove ( Jennings, 1904 ; Fig. 10 B ). As a result, the cell turns toward the aboral side. This is supported by more recent observations in a flattened ciliary sheet from Paramecium ( Noguchi et al., 1991 ). Thus, turning likely results from inhomogeneity in the response of different groups of cilia, but details are still lacking.

The Physiologic Basis of Behavior

The action potential.

When Paramecium touches an obstacle, mechanosensitive channels open, depolarize the membrane and trigger a calcium-based action potential ( Eckert, 1972 ). The entry of calcium then triggers the reorientation of cilia, so that Paramecium swims backwards. Then calcium is buffered or pumped out ( Plattner et al., 2006 ; Yano et al., 2015 ) and the cilia reorient in the original direction.

Historically, Paramecium electrophysiology has been studied by placing the cell in a tiny droplet, letting the fluid evaporate until the cell is captured by surface tension, then inserting sharp microelectrodes and covering with extracellular medium ( Naitoh and Eckert, 1972 ). A recent method immobilizes the cell by suction against a filter ( Kulkarni et al., 2020 ).

Paramecium is an isopotential cell, as demonstrated with two-electrode measurements ( Eckert and Naitoh, 1970 ; Dunlap, 1977 ; Satow and Kung, 1979 ), which is a particularly favorable situation for electrophysiological modeling. This can be sensed from an estimation of the electrotonic length λ = d r m 4 R i , where d is diameter, r m is specific membrane resistance, and R i is intracellular resistivity. For P. tetraurelia , cell width is 34 μm ( Nagel and Machemer, 2000 ), with r m = 64,000 Ω . cm 2 ( Dunlap, 1977 ) and R i = 500 Ω . cm (conservative estimate based on the ∼5 lower intracellular ionic content compared with mammals), we obtain λ ≈ 330 mm, much larger than the cell’s length (115 μm). In the same way, for a 200 nm wide cilium, we obtain λ ≈ 260 μm, much larger than its 10-μm length.

Paramecium has a resting potential of about −30 to −20 mV (more depolarized than neurons), depending on the extracellular medium ( Naitoh and Eckert, 1968a ). P. caudatum has a capacitance of ∼700 pF, half of which is due to the cilia ( Machemer and Ogura, 1979 ), and a resistance of ∼65 M Ω (again depending on the extracellular medium), giving a membrane time constant of ∼45 ms. P. tetraurelia , which is smaller, has a resistance of ∼45–60 M Ω ( Satow and Kung, 1976 ; Nagel and Machemer, 2000 ). Capacitance is not documented, but a simple scaling based on membrane area ( Machemer and Ogura, 1979 ; Nagel and Machemer, 2000 ) gives ∼300 pF. These values are consistent with the surfacic capacitance of other cells including neurons (∼1 μF/cm 2 ).

The negative resting potential is due to a high intracellular concentration of K + ions (18–34 m m depending on studies; Naitoh and Eckert, 1969 , 1973 ; Hansma, 1974 ; Oertel et al., 1978 ; Ogura and Machemer, 1980 ; Oka et al., 1986 ), much larger than the extracellular concentration (typically ∼1–4 m m KCl in experiments; Machemer and Ogura, 1979 ; Machemer, 1998 ). Conversely, there is a low intracellular concentration of Ca 2+ ions at rest (50–200 n m ; Klauke and Plattner, 1997 ; Iwadate, 2003 ), while the extracellular concentration is orders of magnitude higher (the minimal viable concentration is ∼0.1 m m ; Naitoh and Eckert, 1968a ). At rest, the membrane is permeable to many cations ( Naitoh and Eckert, 1968a ). Thus, the ionic content of the cytosol is approximately five times lower than metazoan cells (where intracellular K + concentration is ∼150 m m ). One reason might be that the extracellular medium (fresh water) typically has very low ionic content, so that the cytosolic ions exert a large osmotic pressure on the membrane. In Paramecium and other protozoa, this osmotic imbalance is regulated by specialized organelles, the contractile vacuoles, which expel water that invades the cell by osmosis ( Allen and Naitoh, 2002 ).

When Paramecium is mechanically stimulated on the front, or a current is injected, the membrane is depolarized ( Fig. 11 ). If the stimulus is strong enough, this depolarization triggers a graded action potential, with a stimulus-dependent amplitude (all-or-none spikes can occur if extracellular calcium is partially replaced by barium; Naitoh and Eckert, 1968b ). This action potential is due to calcium voltage-gated channels distributed over the cilia and delayed rectifier potassium channels located in the somatic membrane; this can be demonstrated by removing the cilia with ethanol and shaking ( Machemer and Ogura, 1979 ). In response to a voltage step, the cell produces a current consisting of two phases: a fast inward current carried by Ca 2+ , and a slower outward current carried by K + ( Fig. 12 ), which have been separated using behavioral mutants ( Saimi and Kung, 1987 ).

Membrane potential responses to mechanical stimulation with a glass stylus on the front ( A ) and on the rear ( B ; from Naitoh and Eckert, 1969 , with permission; top traces: voltage command to the piezoelectric actuator).

Action potential currents in P. caudatum (from Brehm and Eckert, 1978a , with permission). A , Current recorded in voltage-clamp with different depolarization steps above resting potential. The first and last peaks are capacitive transients. The early negative transient is mediated by calcium; the late positive current is mediated by potassium. B , Early and late currents versus membrane potential (relative to rest).

The Ca 2+ current inactivates quickly (a few milliseconds) by a calcium-dependent mechanism: the entry of calcium (rather than voltage) inactivates the channels ( Eckert and Brehm, 1979 ; Brehm et al., 1980 ; Eckert and Chad, 1984 ), there is also a slower voltage-gated inactivation acting over tens of seconds ( Hennessey and Kung, 1985 ). Recovery from inactivation takes a few tens to a hundred of milliseconds ( Naitoh et al., 1972 ; Brehm et al., 1980 ). This is a common form of inactivation of calcium channels in neurons, which has been discovered first in Paramecium ( Brehm and Eckert, 1978a ). It involves calmodulin, a highly conserved calcium sensor that is found across all species ( Ben-Johny and Yue, 2014 ). Genetically, three related α units have been identified in the cilia ( Lodh et al., 2016 ), which are similar to the Ca V 1 mammalian family (L-type).

The voltage-gated K + current is a delayed rectifier current, which activates quickly (a few milliseconds; Eckert and Brehm, 1979 ) and inactivates slowly (a few seconds; Satow and Kung, 1980 ; Saimi et al., 1983 ). There is also a calcium-activated current, which develops more slowly ( Satow and Kung, 1980 ). It is involved in repolarization after sustained stimulation ( Saimi et al., 1983 ). Genetic analysis has identified in particular SK channels located in the cilia ( Valentine et al., 2012 ; Yano et al., 2013 ). All these channels have homologs in mammalian neurons.

Currents selective for Na + ( Saimi, 1986 ; Saimi and Ling, 1990 ) and Mg 2+ ( Preston, 1990 , 1998 ) have also been identified.

Electromotor coupling

Cilia are highly conserved structures. Motile cilia are found not only in swimming microorganisms but also in multicellular organisms including humans, where they are involved in moving fluids, for example the cerebrospinal fluid ( Faubel et al., 2016 ). The cilium contains a cytoskeleton called the axoneme, composed of nine microtubule doublets arranged in a ring around a central pair of microtubules ( Porter and Sale, 2000 ). Dynein motors make microtubule doublets slide on each other, which bends the cilium ( Walczak and Nelson, 1994 ). The activity of these motors is regulated by second messengers, in particular calcium and cyclic nucleotides (cAMP and cGMP).

In the absence of stimulation, cilia beat at ∼10–20 Hz with a power stroke toward the right and rear of the cell. Ciliary reversal is triggered by calcium entering the cell through voltage-gated calcium channels distributed over the cilia: this has been shown by direct intracellular exposure of cilia to [Ca] i > 1 μ m ( Naitoh and Kaneko, 1972 ; by making the membrane permeable with a detergent) and calcium uncaging in the cilia ( Iwadate, 2003 ; Fig. 13 A ). In P. tetraurelia , Oertel et al. (1977) estimated that the largest calcium current triggered by a short voltage step increases the ciliary calcium concentration by ∼20 μ m , which then decays because of buffering and pumping. Thus, stronger current pulses trigger larger and faster spikes, resulting in larger calcium increase and therefore longer reversed beating ( Machemer and Eckert, 1973 ). Cyclic nucleotides (cAMP and cGMP) antagonize ciliary reorientation, i.e., an increased cAMP concentration raises the voltage threshold for ciliary reorientation ( Nakaoka and Machemer, 1990 ).

Electromotor coupling. A , Calcium uncaging in cilia (circle) triggers local ciliary reversal (from Iwadate, 2003 , with permission). B , Beating frequency (filled: positive; open: negative) as a function of membrane potential in voltage clamp (from Machemer, 1976 , with permission). Reversal is indicated by dots. C , Beating frequency versus pCa (-log 10 [Ca 2+ ]) in a permeabilized cell (from Nakaoka et al., 1984 , with permission). Squares and circles are two different permeabilized models, circles being more physiological. Cilia reverse at the minimum beating frequency. D , Cell length versus pCa in a permeabilized cell (from Nakaoka et al., 1984 , with permission).

Beating frequency also changes with voltage ( Machemer and Eckert, 1975 ; Fig. 13 B ). In particular, cilia beat faster when the command voltage is increased above resting potential. Early work in permeabilized cells indicated that calcium controls ciliary reorientation but not beating frequency ( Naitoh and Kaneko, 1972 ), but this was later argued to be because of unphysiological aspects of the permeable models ( Nakaoka et al., 1984 ). In more physiological permeabilized cells, an increase in ciliary calcium concentration above the resting level triggers ciliary reorientation and an increase in beating frequency, matching the effect of depolarization ( Fig. 13 C ). Note that swimming velocity does not exactly follow this frequency increase, because it also depends on the coordination of cilia, which is disrupted when cilia reorient. For small depolarizations, not all cilia reorient ( Machemer and Eckert, 1975 ), which may explain how the organism turns. The cell also contracts when calcium concentration increases ( Fig. 13 D ).

Mechanotransduction

Mechanoreception in Paramecium and other ciliates has been the object of several reviews ( Naitoh, 1984 ; Machemer, 1985 ; Machemer and Deitmer, 1985 ; Deitmer, 1992 ). Touching the anterior part of Paramecium results in membrane depolarization, while touching the posterior part results in membrane hyperpolarization ( Naitoh and Eckert, 1969 ). Six genes of the Piezo family ( Coste et al., 2010 ) have been identified in the genome, similar to those mediating mechanosensitivity in many species including mammals. Ionic channels mediating mechanosensitivity are located on the basal membrane; a deciliated cell is still mechanosensitive ( Ogura and Machemer, 1980 ). Cilia are not directly involved in transduction, but they are involved in the mechanical transfer and filtering of stimuli. For example, the tail has long immobile cilia, which may enhance mechanical sensitivity (in particular to current flows) by spreading the mechanical stimulation over a larger membrane area ( Machemer and Machemer-Röhnisch, 1984 ; Machemer-Röhnisch and Machemer, 1984 ).

Mechanosensitive currents change gradually from the posterior to anterior part because of overlapping spatial gradients of K + and Ca 2+ mechanosensitive channels ( Ogura and Machemer, 1980 ; Satow et al., 1983 ). Currents in the posterior part are mostly carried by fast K + currents (time constant in the 10-ms range), whereas currents in the anterior part are carried mainly by Ca 2+ (and possibly other divalent cations; Satow et al., 1983 ) and are slower (in the 20-ms range; Fig. 14 ). In the middle region, a mixed current can be observed, with an outward then inward component, indicative of a superposition of two ionic channel responses. There are also graded changes in mechanosensitivity along the oral-aboral (dorsoventral) axis.

Mechanosensitive responses measured as a function of stimulation position (A: anterior; P: posterior) in a P. aurelia mutant with no action potential (from Satow et al., 1983 , with permission). Below, K + currents are blocked with TEA.

Mechanical responses have been studied mainly by deflecting a thin glass stylus onto the membrane with a piezo-electric actuator. In another ciliate, Stylonichia , the transduced current increases linearly with the deflection amplitude of the probe; the resulting potential may saturate for strong stimuli, near the reversal potential. Faster deflections reduce response latency without changing the amplitude. When a mutant with defective ciliary calcium channels is mechanically stimulated, ciliary reversal is observed only at the site of stimulation on the anterior membrane ( Takahashi and Naitoh, 1978 ): this indicates that mechanical stimulation only recruits local mechanoreceptors (these can trigger ciliary reversal because the transduced current is carried by calcium). Stimulations integrate both spatially and temporally, with no sign of refractoriness. Finally, the duration of the deflection has no effect on the response. The termination of the current may be due to an adaptation process and/or to the membrane passively retreating from the probe.

Thus, the integration of mechanical stimuli is analog to synaptic integration in a neuron: stimulation at a site produces a transient current through ionic channels, transduced currents are integrated both spatially and temporally, and the resulting potential response may trigger an action potential if it is large enough.

Electrophysiology of the escape reaction

When Paramecium is mechanically stimulated on the rear, the membrane is hyperpolarized ( Fig. 14 ), which then triggers the escape reaction: swimming velocity increases. The electrophysiological response is shaped by several hyperpolarization-activated channels.

A fast inward rectifier K + current is activated by hyperpolarization, most strongly below E K ( Oertel et al., 1978 ), and partially inactivates over a few hundred milliseconds ( Preston et al., 1990 ). Thus, the activation voltage depends on extracellular K + concentration. If that concentration is very low, a regenerative hyperpolarization can be obtained ( Satow and Kung, 1977 ). This current is similar to inward rectifiers found in other species ( Doupnik et al., 1995 ). Another K + current activates with calcium ( Preston et al., 1990 ).

A calcium current activates with hyperpolarization, and the entry of calcium then mediates an increase in beating frequency ( Nakaoka and Iwatsuki, 1992 ; Preston et al., 1992a , b ). This current actually activates within a few tens of ms, and decays more slowly through calcium-dependent inactivation ( Preston et al., 1992b ). It actually consists of two pharmacologically distinct currents located in the somatic membrane, one of which is sustained ( Nakaoka and Iwatsuki, 1992 ). The magnitude of the hyperpolarization-activated calcium current is directly related to the increase in beating frequency, and blocking this current also blocks the frequency increase ( Nakaoka and Iwatsuki, 1992 ). Thus, it appears that beating frequency is controlled by calcium concentration in the somatic membrane, presumably at the base of cilia, in line with studies in other ciliary systems ( Tamm, 1994 ). This contradicts several earlier hypotheses: that beating frequency increases with a hyperpolarization-induced decrease in ciliary calcium concentration ( Machemer, 1974 ), by a iontophoretic mechanism in the cilia ( Brehm and Eckert, 1978b ), or by regulation by cyclic nucleotides ( Satir et al., 1993 ; Pech, 1995 ). The latter hypothesis did receive some support ( Bonini et al., 1986 ; Hamasaki et al., 1991 ; Schultz et al., 1992 ), as raising cAMP concentration makes cilia beat faster, but it has been disproven by the demonstration that, when the cell’s voltage is maintained constant, injecting high levels of cAMP has no effect on beating frequency ( Hennessey et al., 1985 ; Nakaoka and Machemer, 1990 ). Thus, the effect of cAMP was likely indirectly due to the hyperpolarization induced by cAMP ( Bonini et al., 1986 ).

Gomez-Marin and Ghazanfar described three fundamental biological principles of behavior that highlight the need for integrated approaches in neuroscience: materiality, agency and historicity ( Gomez-Marin and Ghazanfar, 2019 ). Materiality refers to the role of body and environment in behavior. That is, the relation between neural activity and behavior is not just a case of correspondence (the coding view; Brette, 2019 ), but also of physical causality: spikes cause particular physiological effects, the results of which are determined by the structure of the body and the environment it interacts with ( Tytell et al., 2011 ). For example, in Paramecium , cilia are under electrical control but efficient motor coordination is partly achieved by hydrodynamic interactions between cilia. Agency refers to the fact that action and perception form a closed loop in the service of goals, rather than a linear stimulus-reaction chain. For example, when Paramecium meets an obstacle, the mechanosensory signal is determined not just by the object but also by the motor response that the signal causes, in a closed loop. This concept is increasingly appreciated in cognitive science, philosophy of mind and more recently neuroscience ( Maturana and Varela, 1973 ; Powers, 1973 ; Gibson, 1979 ; Brooks, 1991 ; Bickhard and Terveen, 1996 ; Hurley, 2001 ; O’Regan and Noë, 2001 ; Ahissar and Assa, 2016 ; Pezzulo and Cisek, 2016 ; Brette, 2019 ). Historicity refers to the fact that organisms are individuals: variability is best understood not as a noisy deviation around a norm but as a functional result of their history. In Paramecium , this is evident for example in long-term adaptation to new environments, but also in some exploratory behaviors (such as tube escape).

Addressing these three principles requires studying an entire organism in an environment, rather than isolated subsystems. Computational neuroethology is a subfield of computational neuroscience focusing on the modeling of autonomous behavior ( Beer, 1990 ), which has been investigated in particular artificial organisms ( Beer and Gallagher, 1992 ) and robots ( Webb, 2001 ). More recently, integrated models of C. elegans ( Izquierdo and Beer, 2016 ; Cohen and Denham, 2019 ), Hydra ( Dupre and Yuste, 2017 ; Wang et al., 2020 ), and jellyfish Aurelia aurita ( Pallasdies et al., 2019 ) have been developed. Those model organisms have certain obvious advantages over Paramecium , namely the fact that they have a nervous system, with interacting neurons. But Paramecium has great assets for integrative modeling of a whole organism, relating physiology and behavior.

First, there is an extensive literature on Paramecium , covering detailed aspects of behavior, genetics, electrophysiology, cell and molecular biology. This literature has highlighted similarities with metazoans, in particular nervous systems, not only functionally but also at genetic and molecular levels ( Connolly and Kerkut, 1983 ; Hinrichsen and Schultz, 1988 ; Beisson et al., 2010b ; Yano et al., 2015 ; Plattner and Verkhratsky, 2018 ), with similar ionic channels, pumps, signaling pathways (calcium, cyclic nucleotides), sensory receptors, even GABA receptors. Second, it benefits from various tools, for example genetic tools such as RNA interference ( Galvani and Sperling, 2002 ), proteomics ( Yano et al., 2013 ), and whole genome sequencing ( Aury et al., 2006 ; Arnaiz et al., 2010 ; Arnaiz and Sperling, 2011 ; McGrath et al., 2014 ), behavioral monitoring ( Drescher et al., 2009 ), immobilization for electrophysiology ( Kulkarni et al., 2020 ). Finally, it is easy to culture ( Beisson et al., 2010a ), it has a rich behavior that can be easily observed and quantified, and it allows intracellular electrophysiology in an intact organism, while monitoring its behavior.

As outlined in this review, a number of neuroscientific themes can be addressed and revisited in Paramecium. One such theme is the physiological basis of behavior and the relation between perception and action. A classical way to frame this problem is what Susan Hurley called the “classical sandwich” ( Hurley, 2001 ): at the periphery, a perceptual system transforms stimuli into representations and a motor system transforms motor representations into actions; sandwiched between perception and action, cognition manipulates representations. As noted by many authors, the classical sandwich has many conceptual issues ( Gibson, 1979 ; Brooks, 1991 ; O’Regan and Noë, 2001 ; Pezzulo and Cisek, 2016 ; Brette, 2019 ). Cisek, for example, noted that it leaves the cartesian dualistic view essentially unchanged, replacing the non-physical mind by “cognition” while preserving problematic homuncular concepts ( Cisek, 1999 ). Another key issue is that framing neural activity as responses to stimuli denies any autonomy to the organism. As Dewey pointed out ( Dewey, 1896 ), sensory signals are as much causes as consequences of the organism’s activity, because the relation between organism and environment is one of coupling rather than command. By its relative simplicity, Paramecium offers the possibility to study the physiological basis of autonomous behavior outside the frame of the classical sandwich, because it seems feasible to develop closed-loop dynamical systems models of the organism behaving autonomously in an environment, where spikes are not symbols but actions ( Brette, 2019 ).

Motor control is a related theme where Paramecium may provide some insights. Embodiment is the idea that the body can contribute to motor control, beyond the mere execution of central commands. In Paramecium , cilia beat in a coordinated fashion in the absence of central command, by hydrodynamic and mechanical interactions, yielding efficient swimming. More generally, the mechanical properties of its body contribute to its navigation abilities, as when navigating in confined spaces, and more generally when interacting with surfaces. As it turns out, Paramecium appears to use neither of the two mainstream concepts in motor control, planning (or feedforward control; Wolpert and Ghahramani, 2000 ) and feedback control ( Powers, 1973 ). Instead, it uses another way to produce goal-directed behavior, based on the Darwinian insight that random exploration and elimination of unsuccessful attempts can produce adapted behavior. This simple principle allows Paramecium to perform non-trivial sensorimotor tasks with a single “neuron”.

While the physiological basis of learning is classically framed in terms of stimulus association, Paramecium may offer the possibility to address it in a more ecological context, that is, autonomous learning of a task. Tube escape might be such a task; however, the learning capabilities of Paramecium are still somewhat unclear.

As Paramecium is both an organism and a cell, it also offers the opportunity to investigate the relation between cellular plasticity and behavioral plasticity. Intrinsic plasticity is well documented in neurons ( Daoudal and Debanne, 2003 ), but it remains very challenging to understand its functional implications for the organism. Thus, it is classically interpreted in terms of homeostasis of cellular properties (e.g., firing rate), or of abstract information-theoretical properties. In Paramecium , since the relation between cellular physiology and behavior is more direct than in brains, it becomes possible to relate intrinsic plasticity with behavioral plasticity. For example, ionic channel properties adapt to changes in temperature in such a way as to preserve normal motor behavior ( Nakaoka et al., 1982 ; Martinac and Machemer, 1984 ). Similarly, developmental plasticity can be addressed by investigating the physiological and behavioral changes after fission ( Iftode et al., 1989 ). Indeed, as ionic channels are spatially organized (for example depolarizing mechanoreceptors at the front), this organization is disrupted by fission and must be somehow restored.

This opens exciting perspectives for the development of integrated models of a “swimming neuron”.

Reviewing Editor: Juan Burrone, King’s College London

Decisions are customarily a result of the Reviewing Editor and the peer reviewers coming together and discussing their recommendations until a consensus is reached. When revisions are invited, a fact-based synthesis statement explaining their decision and outlining what is needed to prepare a revision will be listed below. The following reviewer(s) agreed to reveal their identity: Kirsty Wan.

This manuscript provides a detailed review of what can be considered an extreme version of a reductionist neuronal system - a single-cell organism. When searching for simpler systems to study physiology and behaivour, none can be simpler than the paramecium. And yet both the behaviour and physiology of this single-celled organism are also very rich. This work describes in detail the many behaviours of mainly two paramecia P. caudatum and P. aurelia. and links this behaviour to their physiology, which includes a number of voltage-gated channels that decode external inputs and convert it into motion. This extensive review is well written, thought provoking and should of interest to a wide scientific community. Below are some specific comments on the manuscript.

Neuroscience is often, almost by definition, predicated upon knowledge and insights gained from various animal models with well-studied nervous systems. The simplest organisms are often overlooked, yet are capable of surprisingly complex behaviours. The ethology of unicells had been the subject of intense study several decades ago, but is now experiencing something of a resurgence in interest.

This comprehensive review summarises and synthesises much of the data available in the literature about the bioelectrical basis of behaviour in the model ciliate Paramecium. The work will be of interest to a broad audience, particularly biophysicists working on cell motility, while serving to (re)acquaint neuroscientists with this unique model system. Some mechanisms are vague or only hypothetical at this stage, but this is understandable given lack of data.

I have some recommendations that could improve the paper.

1. The work is very much a synthesis of old results - all figures are taken directly from the old literature. I would have liked to see a more elaborate discussion of what this could mean from the ‘integrative neuroscience’ perspective. How might the (motor) behaviour of paramecium be interpreted from the perspective/language of neuronal control principles? There is some mention of this in the last section, but this can be expanded, especially for the intended neuroscience readership.

2. The entire paper focuses on Paramecium, while this may be justified, it would be useful to highlight any conserved features or comparisons with other systems (in the discussions but also throughout) - this may include other ciliated organisms, or other model organisms typically featured in neuroscience. For instance, details of the capacitance of the cell are given (3.1), but how do these values compare with other cell types - ‘true’ neurons? What about the nature, identity, directionality of ionic currents, resting state of membranes etc, how do all these compare with ‘true’ neurons? Which receptors are shared by paramecium and metazoan neuron-types?

3. ‘many signalling pathways of neurons have been found in Paramecium’ - can these connection be expanded? again with above comments in in mind.

4. section 1 - characterisation of the swimming mechanism/trajectories seem to be based on old literature from Jennings et al, is there anything more recent? particularly, is it certain that the oral grooves always faces the inside of the spirals? what is known about the handedness - do all paramecia have the same handedness - is it fixed or can the same individual switch, for example when reversing or responding to cues?

5. Again, many descriptions originate from stylised accounts by Jennings, it would be good to check if these observations indeed hold true, e.g. provide more recent references, if any. Fig 4 - is there other evidence of this type of graded response? what happens when organisms interact with boundaries - where does that fit into the classification? Similarly does turning always happen on the same side?

6. Ciliates often contract - which provides another route for turning/other reactions, this isn’t really mentioned - a contractile cytoskeleton may also respond to membrane potential? is there any evidence of this?

7. The authors may wish to revisit and clarify what they mean by task-driven “trial and error”. After all, bacterial chemotaxis can also be considered a form of trial and error. What is different here (1.4)? If repeated encounters with an obstacle leads to a ‘reversal’, then it can just repeat this until *by chance* it manages to escape. So the question really is whether these patterns of reversal, or orientations say, actually change over successive trials?

minor points

Some sentences/concepts are repeated - the authors are advised to proofread to remove any redundancies.

are calcium channels located exclusively in the cilia?

the authors speak of a dorsal-ventral axis, is this the correct term for unicellular organisms, what about oral-aboral?

does ejection of trichocysts really lead to propulsion? (section 1.2)

It is stated that the cell should be isopotential, a few explanatory sentences may be useful to explain why this should necessarily be the case.

fig 11, in BW it’s very hard to tell the [Ca2+] and [K+] curves apart.

On page 7, instead of ‘releases GABAB’ it should read ‘releases GABA’

Author Response

Response to reviewers

Dear editor and reviewers,

Thank you for these very relevant remarks. In this revision, I have expanded the discussion of the bridges between Paramecium neuroscience and mainstream neuroscience. Regarding the behavior, much of what is known is indeed very old and in need of more modern studies. I have tried to emphasize this fact. I have also responded to each remark in detail. Unrelated to these remarks, I redrew Fig. 10A for copyright reasons.

I hope you will like this revised version.

Best regards,

Romain Brette

Detailed response

Regarding motor control, there are two paragraphs in section 1.4, which I have now expanded a little. I have also expanded the discussion to address the broader issue.

I have added some comparisons throughout the paper. There is work on other ciliates, for example Stentor, but I do not know it as well. Generally, electrophysiology is much more developed in Paramecium than in other ciliates, but there is very interesting behavioral work on various ciliates.

I have added some detail.

Indeed, most behavioral work has been done by Jennings and contemporary scientists (note however that there are a number of more recent references in this section). There are a number of 20th century papers showing trajectories, but not so much their fine structure. Specifically, the observation that the oral groove faces the spiral axis has been reported after Jennings by a couple of authors, in particular by Bullington (ref. 153), but this is also quite old (1930) and to my knowledge it has not been quantified. This is clearly an area where further work would be useful, in particular with high-quality 3D recordings. Regarding the handedness, as far as I know it is always the same, but again this has not been quantified systematically. I added some comments about this issue.

Regarding graded responses, there is evidence about the gradedness of backward swimming in the electrophysiological literature, that is, in observations of ciliary reversal in immobilized cells. I have added a reference (which was previously only mentioned in section 3). It is easy to observe in swimming cells, but to my knowledge it has not been quantified systematically. About turning: Jennings is categorical in his writings that it always happens on the same side, but to my knowledge this also remains to be measured systematically. In brief, there are gaps in the literature regarding detailed aspects of behavior (there is much more detailed knowledge about electrophysiology).

Regarding boundaries (I assume the reviewer means mechanical boundaries), as I wrote at the beginning of the section, Paramecium often gives the avoiding reaction when it hits an obstacle. However, it is not clear how this encounter relates to the intensity of the reaction. Then in addition to this active response, there are also hydromechanical interactions. Clearly sometimes paramecia slide along an object without giving the avoiding reaction (our observations). There is some literature on these effects in other ciliates, and in the physics literature.

I have added comments and a few references in the revised text.

Absolutely. This is very briefly mentioned at the end of section 3.2 and in 1.2 (“This speed increase is accompanied by a contraction along the longitudinal axis (52)”). To my knowledge, it has only been reported in electrophysiological studies of immobilized cells. This is presumably because it happens during the escape reaction, and therefore is probably very difficult to notice in motile cells.

Yes, run-and-tumble behavior of bacteria shares some similarity with Paramecium chemotaxis, except that Paramecium withdraws before changing direction, and has graded reactions. Berg (1975) describes the difference in the following way: “It was thought for many years that bacteria back up or choose new directions at random on entering regions which are unfavourable, that is, that the motor reflex is an avoidance reaction. This has not proved to be the case for E. coli. [...] As the concentration increases, the bacteria change direction less frequently; as it decreases, they swim as they do in the absence of a stimulus”.

In particular, it appears that Paramecium’s behavior is more deterministic than the behavior of bacteria. Biophysically, the membrane area is roughly 2 orders of magnitude larger in Paramecium than in E. Coli, so we expect that signal-to-noise ratio is about one order of magnitude larger. This certainly makes a difference in terms of sensing. Schematically speaking, bacterial run-and-tumble might be described as biased randomness (modulation of the rate of tumbling) whereas Paramecium behavior is perhaps better described as exploration and decision.

Another difference, which is maybe more a difference of degree than nature, is that the avoiding reaction is used in a variety of tasks; avoiding obstacles, thermal homeostasis, avoiding chemicals, etc. I am not sure bacterial behavior has the same diversity (but I am not an expert).

Here “trial-and-error” is just meant as: it tries a swimming direction, and if this leads to a bad or suboptimal situation, then it goes back and tries a different one. I assume the last question refers to learning, i.e., whether the “error” leads to a more persistent change in behavior. This is the aim of section 1.6. The situation described by the reviewer is closest to the behavior described briefly in section 1.2: when trapped in a narrow capillary with a dead end, Paramecium first does a few avoiding reactions, then after a minute, it suddenly starts making much stronger reactions (longer backward swimming).

My reading of Jennings is that to explain the behavior of Paramecium, one may call on the Darwinian insight that the selection of random “explorations” can lead to apparently goal-directed behavior, which at first sight one might be tempted to explain by planning (“intelligent design”) or by steering (Lamarckism) - somewhat analogs of feedforward and feedback control in motor neuroscience. Jennings speculates that the selection of actions is somehow stabilized physiologically, but I don’t think this has been established (we may speculate that this is what happens in the tube escape behavior).

I have expanded this section to address these issues.

Thank you, I have tried to eliminate these redundancies.

As far as I know, the calcium channels responsible for the avoiding reaction are only (or at least essentially) in the ciliary membrane, but there are other calcium channels in the somatic membrane (e.g. those described in section 3.4).

Indeed, Jennings uses the term aboral. I thought dorsal and ventral might be more intuitive for many readers. I changed to oral/aboral, with dorsal/ventral in brackets.

Yes, at a speed of 1-10 mm/s according to Hamel et al. In that paper (ref. 52), they explain the physics of the phenomenon.

I added a calculation of electrotonic length.

That’s a misunderstanding, actually the legends for [Ca2+] and [K+] do not refer to two curves, but describe the composition of the extracellular medium (1 mM CaCl2 and 1 mM KCl). I removed it as it is confusing, and clarified the caption.

Indeed, thank you.

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What is the life cycle of Paramecium ?

Paramecium: paramecium or paramoecium is a genus of unicellular ciliated protozoa. they are characterized by the presence of thousands of cilia covering their body. they are found in freshwater, marine and brackish water and are also found attached to the surface. life cycle of paramecium has asexual and sexual stages. 1.asexual stage: asexual reproduction in paramecium is by binary fission. the mature cell divides into two cells and each grows rapidly and develops into new organism. under favorable conditions. paramecium multiplies rapidly up to three times a day. 2.sexual stage: sexual reproduction in paramecium takes place only when there is scarcity of food. sexual reproduction is by conjugation or autogamy. in conjugation , two complementary paramecia come together and form the conjugation bridge. united paramecia are known as conjugants. macronuclei of both the cells disappear. the micronucleus of each conjugant form 4 haploid nuclei by meiosis. three of them nuclei degenerate. the haploid nuclei of each conjugant then fuse together to form diploid micronuclei and cross fertilization takes place resulting in formation of new daughter cells. in autogamy i.e., self fertilization, a new macronucleus is produced which increases their vitality and rejuvenates them..

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Biocommunication of Ciliates pp 277–304 Cite as

Paramecium as a Model Organism for Studies on Primary and Secondary Endosymbioses

• Yuuki Kodama 3 &
• Masahiro Fujishima 4  
• First Online: 24 May 2016

866 Accesses

2 Citations

Endosymbiosis is a driving force in eukaryotic cell evolution. This phenomenon has occurred several times and has yielded a wide diversity of eukaryotic cells. Despite the importance of endosymbiosis, however, molecular mechanisms for its induction between different microorganisms are not so well known. To elucidate these mechanisms, experiments for synchronous induction of the endosymbiosis by symbionts isolated from the symbiont-bearing host cells and the symbiont-free host cells are indispensable. Also, the infection process needs to be easily observable under a microscope. In many endosymbiotic communities, however, both the endosymbionts and the symbiont-free host cells have already lost the ability to survive and grow independently. Consequently, re-induction of the endosymbiosis was difficult. We have developed optimum experimental conditions for the induction of primary and secondary endosymbiosis using the ciliate Paramecium and their endosymbionts.

• Host Gene Expression
• Symbiotic Alga
• Secondary Endosymbiosis
• Host Nucleus
• Host Cytoplasm

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Acknowledgements

This work was supported by a JSPS KAKENHI Grant-in-Aid for Young Scientists (B) Grant Number 26840119 to Y.K., and also by a JSPS KAKENHI Grant Number 26291073 and a MEXT TOKUBETSUKEIHI to M.F. Paramecium strains used in this chapter were provided by the Symbiosis Laboratory, Yamaguchi University, with support in part by the National Bio-Resource Project of the Japan Agency for Medical Research and Development (AMED).

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Yuuki Kodama

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Masahiro Fujishima

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Kodama, Y., Fujishima, M. (2016). Paramecium as a Model Organism for Studies on Primary and Secondary Endosymbioses. In: Witzany, G., Nowacki, M. (eds) Biocommunication of Ciliates. Springer, Cham. https://doi.org/10.1007/978-3-319-32211-7_16

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