Protist, Vol. 165, 317–329, May 2014 http://www.elsevier.de/protis Published online date 1 April 2014

ORIGINAL PAPER

Haplomyxa saranae gen. nov. et sp. nov., a New Naked Freshwater Foraminifer Marc Dellingera,1 , Amandine Labata , Loïc Perrouaultb , and Philippe Grelliera a“Molécules

de Communication et Adaptation des Microorganismes”, UMR 7245 CNRS MNHN, Muséum national d’Histoire naturelle, CP 52, 57 rue Cuvier, 75005 Paris, France b“Structure et Instabilité des Génomes”, UMR7196 CNRS - MNHN and INSERM U565, Muséum national d’Histoire naturelle, CP 26, 57 rue Cuvier, 75005 Paris, France Submitted November 20, 2012; Accepted March 14, 2014 Monitoring Editor: David Moreira

A new naked foraminifer, Haplomyxa saranae gen. nov. sp. nov., is described from an established cell line made from a single cell isolated from a freshwater garden pond. The new species was morphologically close to Reticulomyxa filosa, the only valid naked freshwater foraminifer species. However the two species differed when it came to the morphology of the cell body, the number of cysts, and the nutrition. The 18S rRNA gene had one of the longest sequences to date (4863 nucleotides), and it contained many insertions that are typical of Foraminifera. The size of this gene was 45% longer than the one of R. filosa due to the elongation of A+T rich regions, but molecular phylogeny based on conserved regions of the 3 -end placed the new species in the same morphological clade K. This report includes both morphological and genetic data which undoubtedly show that the new species is a new naked freshwater foraminifer and the second species of the clade K. © 2014 Elsevier GmbH. All rights reserved. Key words: 18S rDNA; Foraminifera; phylogeny; protist; Reticulomyxa.

Introduction Foraminifera (d’Orbigny, 1826) are single cell protists that often form a multilocular test and share a common specific net-like organization of pseudopodia (Bowser and Travis 2002; Lee et al. 2002). Classifications were primarily based on the morphological description of the test (Cavalier-Smith 1993) and differentiated between multilocular (Polythalamea Ehrenberg, 1838), unilocular (Monothalamia Haeckel, 1862) and testless (Athalamia Haeckel, 1862) forms, while recent classifications were rather based on the test structure (Sen

1

Corresponding author; fax +33 140 793 499 e-mail [email protected] (M. Dellinger).

http://dx.doi.org/10.1016/j.protis.2014.03.007 1434-4610/© 2014 Elsevier GmbH. All rights reserved.

Gupta 1999 and references therein), excluding testless organisms. All organisms with this particular pseudopodial organization were grouped into a single phylum (Granuloreticulosa Lee, 1990), that was divided into testless (Athalamea) and testate (Foraminifera Lee, 1990) forms (Lee et al. 2002). However, molecular data, when available, grouped all these organisms together in a monophyletic rank based on specific insertions in the 18S small subunit ribosomal gene (18S rDNA) (Pawlowski 2000), thus making the phylum Granuloreticulosa synonymous with Foraminifera (Lipps et al. 2011). The most recent protist classification (Adl et al. 2012) divided Foraminifera (d’Orbigny, 1826) into three ranks: (i) one rank (Monothalamids Pawlowski et al. 2003) included all species with a single chamber test (monothalamous), no

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matter whether they have an organic (Allogromiida) or agglutinated (Astrorhizida) wall, or no test wall at all (“naked” species); (ii) two other ranks (Tubothalamea and Globothalamea Pawlowski et al., 2012; Pawlowski et al. 2013) included multi-chambered species. Most of the supposed athalamean species have only been described morphologically, and as a consequence data for molecular phylogeny are missing. They are now considered as “amoebae of uncertain affinities” (Patterson et al. 2002) or insertae sedis (Adl et al. 2012). Only Reticulomyxa filosa (Nauss 1949), which was rediscovered in the 80’s by N. Hülsmann (Hülsmann 1984) and M. Schliwa (Schliwa et al. 1984) who established cell cultures, is still alive in several institutions, at least in the University of Bochum (Germany) with a replicate in the Muséum national d’Histoire naturelle (Paris, France). This cell line was undoubtedly shown to be a foraminifer (Pawlowski et al. 1999) as it possesses the typical foraminiferan insertions in the 18S rDNA (Pawlowski 2000) and is grouped with Foraminifera in molecular phylogenies using rDNA, actin and tubulin genes, and in multigene phylogenies (Burki et al. 2010; Sierra et al. 2013). In rDNA phylogenies of Foraminifera, R. filosa is branching within monothalamous (unilocular) testate species, which challenges the division between Athalamea and Monothalamea (Holzmann et al. 2003; Pawlowski et al. 1999; Pawlowski and Holzmann, 2002). Finally, the athalamiids (Reticulomyxa) could evolved from the ancestor common to Acantharea and Foraminifera (Burki et al. 2010) or more likely from testate species that lost their test secondarily (Pawlowski et al. 2003). R. filosa is a naked freshwater species that was initially discovered on decaying leaves in a metropolitan area of New York City (USA) (Nauss 1949), while the current strain comes from a German lake. Other strains or similar reported species (Hülsmann 1984; Koonce and Schliwa 1985) have all been lost before any molecular analysis could be done. On the other hand, all the supposed freshwater foraminiferan species suggested by molecular phylogeny analyses of lake and soil environmental samples lack morphological observations (Habura et al. 2004, 2008; Holzmann and Pawlowski 2002; Holzmann et al. 2003; Lejzerowicz et al. 2010; Pawlowski et al. 2003). The only other freshwater species, Edaphoallogromia australica (Meisterfeld et al. 2001), that has been described using both morphological and molecular data is a testate species and the strain has been lost (Meisterfeld, personal communication). Then, R. filosa is unique in (i) being the only naked species described with both morphological and molecular data and (ii)

being still available as a cell culture. R. filosa has often been used as a cell model for intracellular transport of organelles (Koonce et al. 1986, 1998; Orokos et al. 1997), and more recently, as a reference organism for the genetics and genomics of Foraminifera (Burki et al. 2006; Glöckner et al. 2014). In this paper, we describe a new naked (in the sense of Nauss 1949, i. e. “without wall or other permanently formed enveloping membrane”) freshwater protist related to R. filosa, presently the only valid athalamean species. We show that the morphology of this new species is very similar to R. filosa, but with critical particularities that support the erection of a new genus. We sequenced the 18S rDNA that is of a very large size and used it for phylogenetic analyses that placed the new species into the Foraminifera (d’Orbigny, 1826) as the closest species to R. filosa. After R. filosa, this is the second naked foraminifer whose report is based on both morphological and molecular data.

Results Localization and Frequency One cell was observed in 2005 in a 50 mL sample of water and sediments from a home-made garden pond filled with rain water (pH neutral). This cell could not be maintained in primary culture for more than a few weeks. It is only in October 2008 that a second cell (Fig. 1A) was found in a 0.5 L sample, despite several sampling operations in the same pond every year. We managed to subculture this second cell, thus giving rise to a cell line (Fig. 1B) still maintained in the free-living protist collection of the Muséum national d’Histoire naturelle (Paris, France) with the reference number CEU-MNHN-050.

Cell Culture and Growth Requirements Cells were cultivated with Volvic® spring water as medium, at 21 ◦ C, under low light with normal day/night alternation. No attempts have been made to cultivate it in axenic or monoxenic conditions. Various food sources were tried, most of them inducing encystment or cell death after a few days. No growth was observed with environmental bacteria grown on autoclaved grains of rice or wheat, wheat germs or LB medium or with the eukaryotic preys Chlorella vulgaris or Chilomonas (Cryptomonas) paramecium. Cells were maintained for a few weeks with Cryptomonas

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Figure 1. Cell shape and pseudopodial network. A: bright field microscopic image of the first cell observed in the primo-culture (pseudopodia not visible). B: phase-contrast microscopic image of a cultured cell showing an elongated cell body with two main pseudopodia and some smaller ones. C: bright field microscopic image of a part of a R. filosa cell. D: image of a dried fixed cell obtained by stitching several dark-field (inverted colors) microscopic images showing the network of pseudopodia radiating from the cell body. Scale bars: 500 ␮m.

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sp. or even longer with Chlamydomonas reinhardtii, but long-term healthy cultures were only obtained with Chlorogonium elongatum. Prey was usually added once a week for maintenance and twice a week to increase cell density. Frequent medium changes were usual, but not always required, as some cultures could be unexpectedly maintained as they were for several months. Deteriorating cultures decreased in cell density, had small or rounded floating cells and were usually treated by medium change and frequent subculturing. Over five years of continuous culture, the strain went through several crises evidenced by failing subcultures and decreased feeding activity. We could not find proof that these crises were caused by contaminations with bacteria or unknown small eukaryotes and they therefore still remain unexplained. Healthy cells always adhered to the dish and could not be easily picked up without damage. Then, subculturing was most often done with floating cells that always emerged as a result of cell migration or after a medium change. Subcultured cells usually adhered slightly to the plastic surface in about one hour, but could stay globular for several days until they migrated to a new position. Cell density was usually as low as 5-15 cells (up to 100) per 90 mm Petri dish.

Morphology Cells had a central cell body from which numerous pseudopodia emerged (Fig. 1B, D). The cell body could vary from a small, flat or rounded shape (down to 0.1 mm diameter) to a large, irregular and elongated one (up to 2.5 x 0.3 mm), but the common shape was massive (undivided), unique and elongated (Fig. 1B, D) thus undoubtedly differing from the common shape of R. filosa (Fig. 1C). The cell body could sometimes be branched or with holes, but was never extended and reticulated like R. filosa. The cytoplasm of starved cells appeared to be colorless or grey-brownish, with numerous granules, while in fed cells its color could vary from green to brown due to prey ingestion and digestion. The cytoplasm always presented several adjacent opposite conspicuous flows. Feulgen staining revealed hundreds of small nuclei (2-4 ␮m) equally spread over the central cell body (Fig. 2A, B). In a typical large cell (1300 ␮m x 75 ␮m), it was inferred from the analysis of 12% of the cell that the total number of nuclei was about 1800. The pseudopodia that emerged from any part of the cell were of two types: a few large ones (1-3 per cell) (“veins” as in Nauss 1949 and Hülsmann 1984) that reached a length of several mm (up to 30); and numerous

small ones that formed an interconnected network (anastomosing connections) around the cell body and the large pseudopodia. All pseudopodia showed a bidirectional movement of attached granules and particles (granuloreticulopodia). Contractile vacuoles were present in both cytoplasm and pseudopodia. Cells could encyst (Fig. 2C) but no other forms, such as flagellate stages or any gamete-like bodies, have been observed.

Cell Biology Cells fed by capturing prey with their small pseudopodia and transporting it through the large veins into the cell body. The ingested prey was always included into vesicles, partly digested as shown by their color which turned from green to brown, and then exported out of the cell and released. Although the culture could be maintained for a long time with Chlorogonium elongatum, the natural prey is still not known. The other vacuoles were numerous contractile vacuoles that grew and fused together before the fusion with the plasma membrane occurred, leading to water ejection that left small resting vesicles. The cycle time was around one minute. Cells did not seem to crawl as amoebae do, but regularly migrated to a new position by transferring the whole cytoplasm through a large vein, including organelles and nuclei, that slowly flowed out of the initial cell body into the new one. Cell division occurred when the vein divided or when several veins (usually two, sometimes three) were involved in the migration (Fig. 2E). New cells temporarily spread in a star-like shape (Fig. 2F), then returned to the usual massive elongated one. Cells were usually single, but nearby cells could partly fuse their pseudopodial network and two migrating cells could fuse into a single cell. When over-fed or in altered media, cells often rounded up and randomly made single cysts covered with a larger thin second envelope (Fig. 2C) and containing numerous nuclei (Fig. 2D). Very big cells could sometimes form several cysts in a cluster. Cysts were most often orange-colored when the cells were fed with C. elongatum.

Molecular Phylogeny The gene of the 18S rRNA was completely amplified in five overlapping fragments between external primers sA10 (Pawlowski et al. 1999) and Medlin B (Medlin et al. 1988) and using internal primers. The number of nucleotides in some polyA or polyT series was sometimes difficult to establish (end sequence positions at 243, 432, 707, 1376, 1449,

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Figure 2. Nuclei, cysts and cell migration. Nuclei (A, B). A: phase-contrast microscopic image of a fixed cell stained with the Feulgen reaction. B: bright field microscopic image of the region of interest (white box) in figure A showing numerous stained-nuclei. Cysts (C, D). C: phase-contrast microscopic image of living cysts showing the thin second envelope. D: bright field microscopic image of a cyst stained with the Feulgen reaction showing numerous nuclei. Cell migration (E, F). E: bright field microscopic image of a cell migrating and segmenting into two daughter cells (pseudopodia not visible). F: phase-contrast microscopic image of a cell showing an extended cell body when migrating. Arrows indicate the direction of cellular migrations. Scale bars A-D, F: 100 ␮m, E 500 ␮m.

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1824, 1919, 1935, 1990, 2094, 2212, 2338, 2370, 2543, 3129, 4763). Most often, we chose the majority value, and for four uncertain series (243, 1449, 1824, 3129) we chose the mean value. There was usually a difference of one (up to three) between the observed number of bases and the mean value. It was not clear whether these base number variations derived from polymerase “slippage” during the PCR reaction or from polymorphism of the gene copies (Pawlowski 2000). The consensus sequence was thus established to be 4863 nt long, with a maximum range variation of -22/+15 nt. Taking into account the usual position of the external primers sA10 and Medlin B in the 18S rDNA of other species, we estimated that the complete sequence should be close to 4900 nt. No group I intron was found using the Rfam database, so we assumed that the rRNA has the same size. To date, this sequence is one of the longest known 18S rDNA sequences (Torres-Machorro et al. 2010; Xie et al. 2011). The closest sequence found by a blast search of the whole gene was the 18S rDNA of R. filosa, and the first hundred sequences were all Foraminifera species. Within these sequences, one environmental sequence (AJ318009, misspelled AJ3180009 in Holzmann et al. 2003) from Lake Neuchâtel (Switzerland) was reported to be close to R. filosa. This short sequence (310 nt, 11 uncertain bases) is very similar to its corresponding part in the new species described herein (309 nt identical or compatible). The sequence of the new species had many insertions, compared with the sequence of Tetrahymena thermophila (1753 nucleotides for X56165). These insertions were similar to those found in other foraminifers such as R. filosa (AJ132367), Cribrothalammina alba (AJ318225) and Allogromia sp. (X86093) (data not shown). The length of the sequence (4863 nt) was 45% longer than the one of R. filosa (3347 nt), mainly due to larger insertions in the eukaryotic variable regions V2 (which reached 770 nt vs 262 and 90 in R. filosa and T. thermophila, respectively) and V4 (1155 vs 391 and 212), and in the helix 15 (344 vs 196 and 26) (using regions and helices as defined in Lee and Gutell 2012 and in Rabl et al. 2011). The G+C content was 27.18%, which is lower than the values for R. filosa (32.58 %) (Pawlowski et al. 1999) and other foraminifers (27.7 to 44%) (Pawlowski 2000). Most of the A+T nucleotides were grouped in A+T rich regions, 77% being in A+T series of at least 5 nucleotides (54% and 29% for R. filosa and T. thermophila, respectively). Phylogenetic analyses of the new species using the 3’ end sequence

of morphologically defined clades of Foraminifera (Pawlowski et al. 2002) placed this new species as a sister species of R. filosa in the clade K (Fig. 3). This grouping is supported by high bootstrap values of 98, 96 and 99%, corresponding to neighbor-joining, maximum parsimony and maximum likelihood, respectively. As in Lejzerowicz et al. (2010), the clade K is included in the Group 2 of soil foraminifers without any other freshwater environmental sequences.

Taxonomic Description The new classification of Eukaryota (Adl et al. 2012) was used to place the new described species into the rank of Monothalamids (Pawlowski et al. 2003). We do not recognize the family Reticulomyxidae (Lee et al. 2002) that was defined based only on the species Reticulomyxa filosa, nor did we place the new species into the genus Reticulomyxa (Nauss 1949) because of the morphological difference between the two species and the reports of several strains of R. filosa that might be different species. We do not recognize the genus Wobo (Hülsmann 2006) and the species Wobo gigas (Hülsmann 2006), despite the record in the Zoological Records, because they are not valid in the sense of the articles 8.6, 9.8 and 16.4 of the International Code of Zoological Nomenclature. We do recognize that N. Hülsmann reported a similar but bigger species that could be of the same genus at the Thirty-Seventh Annual Meeting of the Society of Protozoologists (University of Georgia, Athens, Georgia 19-24 August 1984) (Hülsmann 1984). However, this species was never named and described in a peer-reviewed journal. Consequently, the new genus of Haplomyxa was erected to place the new species, based mainly on the massive shape of the cell body, the genus of Reticulomyxa being restricted to naked freshwater foraminifers with an extended reticulated cell body. Classification following Adl et al. (2012) [Eukarya: SAR: Rhizaria: Retaria: Foraminifera: Monothalamids] Haplomyxa Dellinger gen. nov. Diagnosis: naked multinucleated cell of various shape, cell body from globular to flat extended shape, usually massive and elongated, may be with few branches and/or holes but not extensively and constantly reticulated, cytoplasm with several alternatively opposite flows, producing a few large and extended pseudopodia (“veins”) and a network of interconnected filopodia (reticulopodia as defined

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Figure 3. Molecular phylogenetic analyses. Maximum-likelihood phylogenetic tree of defined (A to M) or environmental freshwater (N) clades and environmental soil groups (1 to 4) of Foraminifera showing the placement of Haplomyxa saranae into the clade K. The tree was rooted with clades I+M as suggested in Pawlowski et al. (2013). Bootstrap values of clades and groups (in the order of neighbor-joining/maximum-parsimony/maximumlikelihood) indicated when reached at least 60%. Dot marks indicate sequences from freshwater and soil environments.

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in Adl et al. 2012) all forming from any part of the cell and bearing granules and attached particles circulating bidirectionally (granuloreticulopodia), numerous cytoplasmic granules and contractile vacuoles, migration and division by translation within large pseudopodia, making covered cysts. Freshwater.Type species: Haplomyxa saranae Dellinger. Etymology: from the Greek haplo (single, simple) and myxa (slime), to report on its massive shape, different from the reticulated shape of Reticulomyxa. Other species: at least two other undescribed species reported by N. Hülsmann might be of the same genus (Hülsmann 1984; Kaufmann D, Wylezich C, Hülsmann N (2006) Deutschen Gesellschaft für Protozoologie: Protist Diversity: Past, Present and Future 8-11 March 2006, Liebenwalde, Germany, Abstract KV4) and one species only known by an environmental rDNA sequence reported from Lake Neuchâtel (Switzerland) (Holzmann et al. 2003). Haplomyxa saranae Dellinger sp. nov. Diagnosis: cell body from small rounded (0.1 mm) to large elongated shape (up to 3.5 x 0.3 mm), usually flat but may become globular in altered media or when forming cysts, cytoplasm colorless or lightly grey-brownish, may appear dark green or brown due to algal preys, binary -sometimes ternary- division by cytoplasm migration through large pseudopods, cyst usually single or few in a cluster, feeding mainly on microalgae. Hapantotype: two slides of cells stained with aniline blue and two others with the Feulgen coloration; mounted in Canada balsam and deposited in the slide collection of the Muséum national d’Histoire naturelle (Paris, France) with the reference numbers ZS112 to ZS115 respectively. Isotype: cell culture deposited in the living protist collection (Collection d’Eucaryotes Unicellulaires) of the Muséum national d’Histoire naturelle (Paris, France) with the reference number CEU-MNHN-050. Type locality: freshwater pond in Saran, 45770 France, sample collected by L. Perrouault in October 2008. Isolator: M. Dellinger. Distribution: only isolated from the type locality, similar species in Lake Neuchâtel (Switzerland) observed by molecular phylogeny analyses of environmental samples (Holzmann et al. 2003). Culture: CEU-MNHN-050 cell line maintained in Volvic® water and fed with Chlorogonium elongatum. Type 18S rDNA sequence: HE965431. Etymology: from the locality of Saran (France) where it was discovered.

Discussion The present study morphologically and phylogenetically identifies H. saranae as a new species of the clade K of Foraminifera which, until now, only contained R. filosa (Hülsmann 1984; Nauss 1949; Pawlowski et al. 2002; Schliwa et al. 1984). Both species are freshwater foraminifers that share many features, but also differ in several morphological, biological and genetic characteristics. The morphology of both species was found to be very similar: a large (up to a few mm) naked (i. e. testless) cell, which is usually flat and elongated, multinucleated, and always surrounded by a network of pseudopodia. Pseudopodia of H. saranae were of granuloreticulopodial type, like those of R. filosa and other foraminifers. The main morphological difference between the two species was found to lie in the shape of their cell body, which is usually massive for H. saranae, while it is reticulated for R. filosa (Fig. 1C). No obvious crawling was observed in either species, but there was a similar migration through large pseudopodia which could result in cell partition. Indeed, both species share a particular mode of division that does not involve a classical cytokinesis, but rather the separation of daughter cells through several main pseudopodia. Cell partition in H. saranae was usually in 2-3 parts, whereas in R. filosa, it is in 3 parts for the American strain (Nauss 1949) and 3-7 parts for the German strain (Hülsmann 1984). Allogromia has been reported to use several modes of division, including binary separation and schizogony (McEnery and Lee 1976). In some reported cases, pseudopodia are also involved in the schizogony, the daughter cells being directed out of the parental cell using the pseudopodial network (Bowser et al. 1984). Although naked athalamids and Allogromia use their main pseudopodia in the cell division, it is not established if these fission mechanisms are related. The two species also differ on the number of cysts. Although both species make cysts which are covered with a second thin layer, H. saranae usually made single cysts, while R. filosa always makes large clusters of cysts. All known freshwater foraminifers, whether of clade K (R. filosa and H. saranae) or of clade M (E. australica), have contractile vacuoles which are osmoregulatory organelles mainly found in freshwater protists. It is not known whether all such contractile vacuoles are similar, but they might have a common origin because they are present in most protist lineages. In that sense, the contractile vacuole would be an ancestral character

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lost secondarily in marine species. Indeed, in some marine protists such as ciliates, the contractile vacuole remains (Kitching 1938), however, they have not been reported in most marine protists such as foraminifers (Patterson 1980). Foraminifera and the marines groups Acantharea and Polycystinea form a monophyletic group (Sierra et al. 2013). The absence of contractile vacuoles in all the marine species and the presence of similar ones in the freshwater species of different foraminiferal clades suggest that the contractile vacuole system has been conserved, at least up to the root of the Foraminifera. To better determine the relationship between H. saranae and R. filosa, we sequenced the gene of the 18S rRNA. This gene was found to be the longest foraminiferal 18S rDNA ever sequenced (4863 nt) and was assumed to be free of intron I, as estimated with the Rfam database. This sequence is much longer than the one of R. filosa (3347 nt) (Pawlowski et al. 1999), with a lower G+C percentage (27.18% vs 32.58%). Expansion of the rDNA is the rule in Foraminifera, which have unusually long sequences (2200 to 5500 nt) (Pawlowski 2000, unpubl. data). Only twelve 18S rDNA sequences in GenBank are longer than the one of H. saranae, but eleven of them include several introns I, so that their rRNA has the usual size for eukaryotic 18S rRNA (1600-2200 nt). Finally, the longest sequence without reported introns (6373 nt, full size estimated as 6500) is from the Euglena sanguinea strain SAG1224-30 (Karnkowska-Ishikawa et al. 2013). Expansion of the rDNA is also common in Euglenozoa in both osmotrophic (Busse and Preisfeld 2002) and phototrophic (KarnkowskaIshikawa et al. 2013) species but depends on the species. Moreover, for the three strains that have been retained as E. sanguinea, the 18S rDNA sequences differ from 4336 nt to 4544 up to 6273 nt for helix 8-48 (Karnkowska-Ishikawa et al. 2013). As expected, the sequence of H. saranae has numerous insertions in the conserved eukaryotic and variable regions of the gene, some of them being specific to Foraminifera. The increase in the rDNA 18S sequence in H. saranae and R. filosa is mainly due to a gene expansion in the variable regions V2 and V4, which are also involved in the gene expansion in E. sanguinea. It has been suggested that variable regions could be used to barcode Foraminifera (Pawlowski and Lecroq 2010), based on the differences between species, but little is known on the intraspecific variations of these variable regions (Pillet et al. 2012). In this study, our molecular phylogeny analyses used previously aligned conserved regions of the

3 -end sequence of the 18S rDNA (Habura et al. 2004) that represent the main morphologicallydefined clades of the Allogromiids (Pawlowski et al. 2002). Total rDNA cannot be used in such analyses due to the lack of the 5 -end in most of the published Monothalamids’ sequences. Phylogenetic analyses with neighbor joining, maximum parsimony and maximum likelihood all returned the same result with high bootstap values, namely that H. saranae is the closest species to R. filosa. Molecular phylogenetic analyses and morphological comparisons are consistent with the finding that H. saranae is the second identified species in the clade K of Allogromiid Foraminifera. The clade K groups only into the second group of soil foraminifers and not with other soil or freshwater groups (Fig. 3). Despite several environmental samplings of freshwaters and soils in Europe and in America (Habura et al. 2004, 2008; Holzmann and Pawlowski 2002; Holzmann et al. 2003; Lejzerowicz et al. 2010; Pawlowski et al. 2003), only one short (310 nt) environmental sequence from Lake Neuchâtel (Switzerland) was previously reported to be close to R. filosa (Holzmann et al. 2003), although this was not shown in any phylogenetic analysis. This sequence is almost similar to its corresponding part in H. saranae, which suggests that the species from Lake Neuchâtel is very close to H. saranae, and should be placed into the same genus, if not into the same species. Another sequence obtained from the same environmental sample (sample 51, Lake Neuchâtel, Switzerland) was grouped with several North American sequences of another clade (clade N in Holzmann et al. 2003), whereas sequences obtained from Lake Geneva (Switzerland) belonged to a third, different clade (clade L). This suggests a rare, even sparse, distribution of clade K species in these freshwater environments. The phylogenetic analysis of Monothalamids using the conserved regions of the 3 -end of the 18S rDNA was not improved by the addition of a second species of the clade K and is still unresolved between the various clades. This could be due to undersampling of taxa, but also to the high evolution rate of this gene in this rank. Then, the use of other genes with a slower evolution rate could be more appropriate to define the root of the Foraminifera. R. filosa has been used as a cell model in cellular dynamics and foraminiferan genetics, but the fast evolving rate of some of its genes may be a source of biased results (Burki et al. 2010). The discovery of a second species within the clade K not only provides another cell model of a naked foraminifer, but may also prove

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helpful in obtaining more accurate phylogenies of the Monothalamids. Although both species live in freshwater, their natural environment is rather unclear as they were only isolated once in anthropogenically influenced sites. Indeed, R. filosa was isolated first in a metropolitan area (New York City, USA) (Nauss 1949), while similar strains were further isolated from the tropical fish pond of a botanical garden (Bochum University, Germany) (Hülsmann 1984) and from a fresh water aquarium (Berkeley, USA) (Schliwa et al. 1984). All these strains were lost before any molecular analysis could be done, so that their relationship is still uncertain. The remaining strain, which was definitively proved to be a foraminifer by several phylogenetic analyses (Pawlowski et al. 1999), was isolated from a German lake (Glöckner et al. 2014). Similarly, H. saranae was isolated from a home-made garden pond, so that it cannot be excluded that it was introduced with non-local plants. Although, two other species which looked like Haplomyxa have been briefly reported at conferences (Hülsmann 1984; Kaufmann D, Wylezich C, Hülsmann N (2006) Deutschen Gesellschaft für Protozoologie Protist Diversity: Past, Present and Future 8-11 March 2006 Liebenwalde, Germany Abstract KV4), and one environmental species has been detected in a Swiss lake (Holzmann et al. 2003), they were all likely from anthropogenically influenced ponds. Finally, more sampling would be necessary to clarify the natural environment of these species. It is still unclear whether such species are rare in their natural environments, which have to be defined, or if they were infrequently reported because of inadequate procedures. There have been reports on several known or possible freshwater Monothalamids (i. e. Edaphoallogromia, Lieberkhunia, Apogromia, environmental soil and freshwater forms) in addition to the naked R. filosa and H. saranae. Morphologies and 18S rDNA sequences, when known, differ between these species, which suggests that this group is not monophyletic and that adaptation to freshwater occurred several times independently. Our knowledge of Monothalamids, and especially of freshwater species, is still poor and should benefit from the isolation and culture of more species. Although H. saranae was not easily isolated and maintained in culture, we believe that better sampling with adequate food supply, such as C. elongatum in place of bacteria, would allow the (re)discovery of other species of the newly erected genus Haplomyxa.

Methods Cell culture of cell lines used as food source: Chlorogonium elongatum, Chilomonas (Cryptomonas) paramecium and Cryptomonas sp. were obtained from Orsay University’s protist collection (Orsay, France). Chlamydomonas reinhardtii and Chlorella vulgaris were obtained from the collection of cyanobacteria and microalgae (PMC-ALCP) of the Muséum national d’Histoire naturelle (Paris, France). Chlorogonium elongatum, C. reinhardtii and C. vulgaris were cultivated with basal Bold medium as defined in the CCAP (http://www.ccap.ac.uk), C. paramecium with Volvic® and wheat grains, and Cryptomonas sp. with S2T2 medium (Lichtlé 1979). Isolation and maintenance: One cell was found by M. Dellinger in a 0.5 L sample collected by L. Perrouault in October 2008 from a home-made garden pond in Saran (France). The sample was distributed into Petri dishes that were observed during several days with the use of an inverted microscope. The primary culture was maintained at 21 ◦ C under natural day/night light alternation with the addition of Chlorogonium elongatum as a food source. After a few days, the cell divided and a detached cell was isolated and subcultured. This cell line are now maintained in Petri dishes filled with Volvic® spring source (Danone, France). The medium is usually emptied out and replaced every 1-2 week and the cells are fed with Chlorogonium elongatum. Subculturing is done by transferring floating cells into new Petri dishes. The strain (CEU-MNHN-050) is available for researchers, for non-commercial purposes, on request to the collection of free-living protists of the Muséum national d’Histoire naturelle (Paris, France). Fixation and staining: Starved floating cells were dropped onto cleaned slides in Quadriperm culture chambers (VWR, France) filled with Volvic® and let to spread for one day. Cells usually completed the digestion of the prey and made a migration, so no C. elongatum were observed in the cytoplasm. Fixation was done using a protocol adapted from R. filosa (Koonce and Schliwa 1985) and marine Foraminifera (Bowser and Travis 2000; Travis and Allen 1981) protocols used to minimize pseudopodial retraction (“beading-response”). Cells in Volvic® were prefixed with 2.5% PHEM (Pipes HEPES EGTA Medium) (Schliwa and Blerkom 1981), then with 2.5% of the fixation solution (50% PHEM containing 7.4% formol and 2% glutaraldehyde). Cells were fixed for one hour at room temperature with the addition of one volume of the fixation solution containing 5% acetic acid, then washed with PHEM 20% and bidistillated water several times each. To observe pseudopodia, the slides were dried at room temperature and observed with dark field microscopy. For nuclei staining, the cells were first treated to minimize detachments, although the pseudopodia were often altered by the Feulgen reaction. The slides were soaked for 30 min in 1% albumin-glycerol, dried at 4060 ◦ C, post-fixed with ethanol/formol (3/1) and passed through ethanol 95% to ethanol 70%, in which they were stored until stained. Cells were hydrated in ethanol 30%, then in bidistillated water. Free aldehydes were then reduced with 10 g/L NaBH4 in 1% diluted Phosphate Buffered Saline (PBS 1%) for 30-60 min and finally washed several times with PBS 1% and bidistillated water. Nuclei were stained with the Feulgen reaction (as described by Fryd-Versavel in Dragesco and Dragesco 1986), using HCL 6N for 20 min at room temperature to hydrolyze the DNA. Slides were finally passed through alcohol series from ethanol 30% to ethanol 100%, then cleared with xylene and mounted in Canada balsam. Microscopy and imaging: Microscopic images of cells, with normal light (bright field), dark field or phase contrast, were

a New Naked Freshwater Foraminifer 327

Table 1. PCR and sequencing primers Name

Direction

Sequence

MDP34 MDP36 MDP35 MDP33 MDP40 MDP41 s10 MDP32 s43RF s17 s14F1 Medlin B

Forward pcr Reverse pcr Forward pcr Reverse pcr Forward sequencing Reverse sequencing Forward pcr Reverse pcr Forward pcr Reverse pcr Forward pcr Reverse pcr

CTCAAAGATTAAGCCATGCAAGTGGTT TGCGTTCCTTAGAACTAAGAGCGGT TCTGATCCCATAGAAGGAGCACCGTA AGGCATTCAATGCACTTTACAGAGACTATCCGA AACTTCAAGTGGAGGGCAAGTCTGGT AACTACGAGCCTCTTAACCGCAACAATGA CACTGTGAACAAATCAG GCAATTGAGAGCCCATCGTTTGATGTAAG GAGGATCAGATACCCTCG CGGTCACGTTCGTTGC AAGGGCACCACAAGAACGC TGATCCTTCTGCAGGTTCACCTAC

obtained using an Eclipse T200 inverted microscope and a DXM-1200C color CCD camera (Nikon, France). Image treatment and analysis were usually done with the ImageJ software (http://imagej.nih.gov/ij/). The Fiji software (http://fiji.sc/Fiji) was used for image stitching (Preibisch et al. 2009). DNA extraction, PCR amplification and sequencing: Ten to one hundred cells per sample were washed with sterile Volvic® by transfer in Petri dishes, then placed in a sterile tube and centrifuged at 1000 g for 5 min. Most of the supernatant was removed and the sample was usually frozen and stored at -20 ◦ C. DNA extraction was done either on frozen or unfrozen samples using a procedure adapted from the “chelex method” (Regensbogenova et al. 2004). Briefly, cells were treated for 30 min at 60 ◦ C with 25 ␮g/ml Proteinase K in 10% Chelex-100 suspension in 10 mM Tris (pH 8.0) plus 0.1% Triton X-100. The tube sample was then placed in boiling water for 5 min, then centrifuged and the supernatant was collected and stored at -20 ◦ C until used in PCR reactions. The extracted DNA was frequently used as it was, but it was sometimes washed in ethanol 70% after precipitation with sodium acetate/ethanol to improve the PCR reaction. The whole sequence of the 18S rDNA, between a foraminiferal specific primer (Table 1) close to the 5 -end of the sequence (sA10) (Pawlowski et al. 1999; Pawlowski 2000) and the universal eukaryotic reverse primer Medlin B (sB in Pawlowski et al. 1999) at the 3 -end (Medlin et al. 1988), was sequenced in five main overlapping sections. Primers that have been used for Reticulomyxa filosa by Pawlowski’s group (s10, s17, s43RF, s14F1) (Pawlowski et al. 1996, 1999) were used as such or slightly modified (MDP34 i. e. 3 -extented sA10). New primers (MDP) were designed using the ARB software (Ludwig et al. 2004) and the SILVA rRNA database (Pruesse et al. 2007) to select highly conserved regions among Foraminifera. Pairs of primers for each section were: MDP34/MDP36 (1147 nt) for section 1, MDP35/MDP33 (1563 nt) for section 2, s10/MDP32 (679 nt) for section 3, s43RF/s17 (935 nt) for section 4, and s14F1/Medlin B (1096 nt) for section 5. Sections 3-4 and 4-5 were also amplified using the pair of primers s10/s17 and s43RF/Medlin B, respectively. The PCR reaction usually consisted of 35 cycles, with 30 s at 94 ◦ C, 1 min at 50-60 ◦ C (depending on the primer) and 3 min at 72 ◦ C, followed by 7 min at 72 ◦ C. The PCR product was purified with illustra GFXTM PCR DNA and Gel Band Purification Kit (GE Healthcare Life Sciences), cloned into competent bacteria using pGEM-T Easy (Promega) vector and sequenced bidirectionally (Beckman Coulter Genomics) using T7P/SP6 primers. The second

fragment (MDP35/MDP33) was also sequenced with the use of 2 internal primers (MDP40 and MDP41). Each section of the gene was covered at least by three different PCR reactions. Sequence analysis and phylogeny: Eighty one chromatograms were analyzed using the software package Phred-Phrap-consed (Ewing and Green 1998; Ewing et al. 1998; Gordon et al. 1998) and manually refined. The consensus sequence has been deposited in the GenBank/EMBL database under the accession number HE965431. The closest relative sequence to the consensus was searched in GenBank/EMBL’s nucleotide database using blastn. Group I introns were searched in Rfam database 10.1 on the Sanger’s website (http://rfam.sanger.ac.uk). The 3 end sequence of the consensus was manually aligned in Seaview (Gouy et al. 2010) with an alignment (Habura et al. 2004) of defined clades of Foraminifera (Pawlowski et al. 2002). Thirty four sequences of morphologically identified foraminifers were selected and aligned with ten freshwater (Holzmann et al. 2003) and twenty six soil (Lejzerowicz et al. 2010) environmental sequences, and with two additional Monothalamid sequences. Variable regions were removed, leaving five hundred and thirty four sites for analyses. Phylogenetic analyses were performed as described by Habura et al. (2004) using Seaview (i. e. maximum-likelihood with seven rate categories corrected using a F84 model with a gamma-distributed rate of variation, neighbor-joining with K2P model, and maximum-parsimony with bootstrap analysis of 100, 1000 and 500 replicates respectively).

Acknowledgements The authors thank G. Fryd-Versavel for the gift of the Chlorogonium elongatum, Chilomonas (Cryptomonas) paramecium and Cryptomonas sp. strains, C. Yepremian for the gift of the Chlamydomonas reinhardtii and Chlorella vulgaris strains, and L. Raveendran for maintaining these cultures. The authors thank Pr. A. Faissner and U. Theocharidis for the gift of the R. filosa strain, and R. Schulz/Breuker for helpful information on this species. Thanks to B. Ewing, P. Green and D. Gordon for the softwares Phred, Phrap and consed,

328 M. Dellinger et al.

respectively. Thanks to S. Mniai for English corrections. This work was supported by the MNHN Programme PluriFormation (PPF) « Biodiversité et rôle des micro-organismes dans les écosystèmes actuels et passés » in 2009.

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