Accepted Manuscript Title: Gregarine site-heterogeneous 18S rDNA trees, revision of gregarine higher classification, and the evolutionary diversification of Sporozoa Author: Thomas Cavalier-Smith PII: DOI: Reference:
S0932-4739(14)00053-4 http://dx.doi.org/doi:10.1016/j.ejop.2014.07.002 EJOP 25340
To appear in: Received date: Revised date: Accepted date:
28-5-2014 19-7-2014 21-7-2014
Please cite this article as: Cavalier-Smith, T.,Gregarine site-heterogeneous 18S rDNA trees, revision of gregarine higher classification, and the evolutionary diversification of Sporozoa, European Journal of Protistology (2014), http://dx.doi.org/10.1016/j.ejop.2014.07.002 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
*Manuscript
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Gregarine site-heterogeneous 18S rDNA trees, revision of gregarine higher classification, and the evolutionary diversification of Sporozoa
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Thomas Cavalier-Smith
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Department of Zoology, University of Oxford, South Parks Road, Oxford OX1 3PS, UK
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E-mail address: [email protected] (T. Cavalier-Smith)
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Abstract Gregarine 18S ribosomal DNA trees are hard to resolve because they exhibit the most disparate rates of rDNA evolution of any eukaryote group. As site-heterogeneous treereconstruction algorithms can give more accurate trees, especially for technically unusually
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challenging groups, I present the first site-heterogeneous rDNA trees for 122 gregarines and an extensive set of 452 appropriate outgroups. While some features remain poorly resolved, these trees
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fit morphological diversity better than most previous, evolutionarily less realistic, maximum likelihood trees. Gregarines are probably polyphyletic, with some ‘eugregarines’ and all
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‘neogregarines’ (both abandoned as taxa) being more closely related to Cryptosporidia and
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Rhytidocystidae than to archigregarines. I establish a new subclass Orthogregarinia (new orders Vermigregarida, Arthrogregarida) for gregarines most closely related to Cryptosporidium and group
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Orthogregarinia, Cryptosporidiidae, and Rhytidocystidae as revised class Gregarinomorphea. Archigregarines are excluded from Gregarinomorphea and grouped with new orders Velocida
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(Urosporoidea superfam. n. and Veloxidium) and Stenophorida as a new sporozoan class Paragregarea. Platyproteum and Filipodium never group with Orthogregarinia or Paragregarea and
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are sufficiently different morphologically to merit a new order Squirmida. I revise gregarine higherlevel classification generally in the light of site-heterogeneous-model trees, discuss their evolution, and also sporozoan cell structure and life-history evolution, correcting widespread misinterpretations.
Keywords: Cryptosporidium classification; Orthogregarinia; Paragregarinea; Squirmidea; Stenophora; Terragregarina.
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Introduction Gregarine phylogeny is currently unclear, partly because of extreme variation (approximately 15-fold) in rate of evolution of their 18S rDNA. Despite that it is well known that existing higher gregarine classification based on 60-year-old evolutionary ideas (Grassé 1953) is
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phylogenetically unsound and has become purely a matter of convenience (Rueckert et al. 2011a). Therefore, having assembled the most extensive gregarine alignment to date in the accompanying
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paper for identifying a gregarine sequence contaminating a heterolobosean culture (Cavalier-Smith
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2014), I have carried out the first comprehensive phylogenetic analysis of gregarines and their alveolate outgroups using a site-heterogeneous model of base substitution (the CAT-GTR-
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GAMMA model) implemented in PhyloBayes (Lartillot and Philippe 2004). This model is evolutionarily more realistic than homogeneous models used for previous maximum likelihood
and Philippe 2008).
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(ML) gregarine trees, and copes better with extreme long branches (Lartillot et al. 2007; Lartillot
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It did indeed produce trees that are more congruent with both morphology and parasite-host preferences than previous gregarine trees, though some features remain unresolved. I discuss
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gregarine and myzozoan evolution generally in the light of these site-heterogeneous model trees, which show that classical gregarines comprise two distinct probably not directly related major clades, here made subclass Orthogregarinia and class Paragregarea, and that a third clade comprising Platyproteum and Filipodium (Rueckert and Leander 2009) is probably unrelated to either but groups weakly with either Perkinsozoa within Dinozoa or Apicomonadea and thus may have evolved independently of Sporozoa. I here establish a new order Squirmida for these two genera, which differ greatly from classical gregarines, and substantially improve gregarine higherlevel classification, dividing them into five orders not three as did Grassé (1953); only his order Archigregarinida is retained, the distinction between eugregarines and neogregarines being arteficial. A long-overdue change is establishing a new order for Cryptosporidium, here formally transferred from Coccidea to Gregarinomorphea as new subclass Cryptogregaria in the light of
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4 much recent morphological and sequence tree evidence (Valigurová et al. 2007). Overall Gregarinomorphea is now divided into three new subclasses and four orders (three new) and two new suborders; all 21 newly established suprafamilial taxa for broadly ‘gregarine’ taxa correspond with major clades on the site-heterogeneous tree (over two thirds strongly supported) and reflect
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reasonable morphological and host range distinctions. I also discuss the probable polyphyletic origins of gregarines in relation to the huge increases in cell and genome size of gut parasites
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compared with small cell size and genomes of intracellular parasites (Cavalier-Smith 1978, 1980,
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1982) and differences in multiple fission, cilia and centrioles amongst the sporozoan classes.
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Material and Methods
Data selection and maximum likelihood (ML) analysis of the 652-eukaryote sample were as
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in the accompanying paper (Cavalier-Smith 2014). Analysis for two smaller taxon samples of 196 or 276 alveolates and 1577 well aligned nucleotides used both this and the site-heterogeneous CAT-
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GTR-GAMMA model of PhyloBayes v. 3.3 (Lartillot and Philippe 2004) with two chains for many generations beyond the point of plateauing and 25% of trees removed as burnin. A third taxon
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sample of 1614 well aligned nucleotide positions for 137 Myzozoa only with all long-branch gregarines and coccidiomorphs excluded was analysed by the same site-heterogeneous model to obtain the most accurate myzozoan tree minimally perturbed by long-branches and distant outgroups; the two chains for this 137-taxon analysis and the 196-taxon analysis each converged well to a single topology, but the 276-alveolate analysis gave alternative slightly different trees for two weakly supported branch positions.
Results and Discussion Bayesian analysis using the site-heterogeneous CAT-GTR-GAMMA substitution model is evolutionarily more realistic than those available for ML (Lartillot and Philippe 2008) and often gives more reliable trees, especially for difficult-to-place taxa such as those with idiosyncratic long Page 4 of 60
5 branches or with very short internal stems (Brown et al. 2013; Roure et al. 2013). Though in these respects gregarines are one of the most phylogenetically challenging of all protist groups these superior site-heterogeneous methods have not been previously applied to them. To make discussion
Contrasts between homogeneous and heterogeneous rDNA trees
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of the trees more comprehensible the new classification is first summarised in Table 1.
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The eukaryote-wide Figure 1 like all other recent maximum likelihood trees groups marine eugregarines and the insect-host Gregarinoidea with archigregarines. One marine group (Velocida,
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comprising Veloxidium and Urosporoidea) is actually within archigregarines, thus partially
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supporting the classical view of Grassé (1953) that archigregarines are ancestral to eugregarines. But, contrary to that idea, most terrestrial gregarines (i.e. Stylocephaloidea/Actinocephaloidea; a
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well-supported (76%) clade here called ‘core orthogregarines’) weakly group with Rhytidocystis, and this clade with Cryptosporidium and not with archigregarines. Archigregarines are paraphyletic,
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some grouping weakly with Velocida and others still more weakly with a heterogeneous longbranch clade comprising Gregarinoidea and marine gregarines (Fig. 1). The basal branching order
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of all these clades has no significant support, and I argue that the phenotypically mixed long-branch grouping with only 1% support is probably a long-branch artefact - it is not found in any siteheterogeneous trees irrespective of their taxon sampling. The PhyloBayes CAT-GTR-GAMMA tree in Figure 2 focuses on Myzozoa only; to clarify relationships of the three main gregarine-like clades to other alveolates it omits the three longest-branch gregarine clades (Trichotokara and superfamilies Gregarinoidea and Porosporoidea). Like Fig. 1 it shows core orthogregarines (Stylocephaloidea/Actinocephaloidea) as a robust clade strongly supported as sister to Cryptosporidium, not to Rhytidocystidae as insignificantly by ML; rhytidocystids are strongly sisters of Cryptosporidium/core orthogregarines, the three groups together forming a robust gregarinomorph clade that excludes both archigregarines and Squirmidea (Platyproteum plus
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6 Filipodium). This Figure 2 site-heterogeneous tree strongly indicates that archigregarines are not sisters to Orthogregarinia, implying that classical gregarines are polyphyletic. As long supposed, Archigregarinida are paraphyletic in Fig. 1 (ML) and in all CAT-GTRGAMMA trees, even if all long-branch gregarine clades and alveolate outgroups are included (Fig.
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3). Archigregarines are clearly ancestral to the robust clade Velocida, comprising a relatively small subset of aseptate gregarines (its subclade Urosporoidea former eugregarines). Figs 2 and 3 also
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show for the first time that the millipede-inhabiting eugregarine Stenophora, not included in recent trees for marine gregarines, is sister to classical archigregarines plus Urosporoidea, not to the
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insect-infecting core orthogregarines; in that respect ML and CAT trees agree. Though Stenophora
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weakly groups with Gregarinoidea on Fig. 1, I give more weight to its grouping with archigregarines and Velocida (together here called core paragregarines) on the probably more
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accurate CAT trees, so core paragregarines and the new order Stenophorida are here grouped as a new sporozoan class Paragregarea. Figs 2-3 suggest more weakly that Paragregarea may not even
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be sister to Gregarinomorphea, but possibly sister to Gregarinomorphea plus Coccidiomorphea (Fig. 2) or to Coccidiomorphea alone (Fig. 3); if either is correct, gregarines are polyphyletic. In all three
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trees the free-living predatory, myzocytotic, apicomonad flagellates (here shown as including the algae Chromera and Vitrella as clearly separate lineages) are the earliest diverging Apicomplexa, sisters to Sporozoa. Both CAT trees (Figs 2-3) agree with the ML tree (Fig. 1) in showing more clearly than before that the recently discovered Squirmida are probably not gregarines or even sporozoa; their precise position however is inconsistent, ML placing them weakly as sisters of all Dinozoa, whereas Fig. 2 puts them within Dinozoa and Perkinsozoa weakly as sister to the also parasitic Perkinsea and Fig. 3 places them within the apicomplexan Apicomonadea. Site-heterogeneous Figure 3 substantially rearranges the long-branch gregarine clades compared with present and past ML trees and past homogeneous Bayesian trees in a way that makes much more sense in terms of both host specificity and morphology than other gregarine trees; and reassuringly it is also largely congruent with the site-heterogeneous Figure 2 that
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7 excluded long-branch gregarines and ciliates, showing that the well-supported basal branching order of Figure 2 is not distorted when long-branch taxa are added, even though their support values unsurprisingly drop. This strongly suggests that less evolutionary realistic site-homogeneous substitution models do generally misplace many long-branch gregarines through artefactual mutual
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attraction, making past trees misleading. Figure 3 includes the gregarines Pyxinia (omitted in Figs 1 and 2 because of its exceptionally long branch for terragregarines), showing it to be an
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actinocephaloid, and like Fig. 2 Apicystis (not in Fig. 1). The CAT trees also include environmental sequence BOLA566 (Dawson and Pace 2002) to check whether it is a gregarine; it clearly is, being
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consistently (never strongly) a distant sister to Stenophora – its inclusion in Figs 2-3 is another
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reason for thinking they probably more accurately place Stenophora than did Fig. 1.
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Are long-branch gregarines orthogregarines or paragregarines?
Figure 3 places the long-branch insect host Gregarinoidea within core orthogregarines as
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sister to superfamily Actinocephaloidea (0.46 PP) to form a clade entirely of gregarines that infect terrestrial arthropods or oligochaetes; core orthogregarines and Gregarinoidea are here united as a
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new taxon Terragregarina, comprising all terrestrial gregarines but Stenophora. Fig. 3 also weakly groups the extremely long-branch crustacean gregarines (Porosporoidea) with Stylocephaloidea (0.68 PP). Terragregarines and Porosporoidea together form a weakly supported (0.34) clade, here made a new order Arthrogregarinida within Orthogregarinia (an extremely weakly supported clade in Fig. 3), assuming that all share an arthropod-gut-dwelling common ancestor. Despite its weak support the arthrogregarine grouping is evolutionarily more reasonable than the even weaker exclusion of Gregarinoidea from Orthogregarinia and their probably artefactual grouping with other long-branch gregarines in the ML tree (Fig. 1). Terragregarines probably ancestrally infected insects and/or arachnids and later one branch colonized earthworms. Fig. 2 also weakly (PP 0.76) groups the very-long branch polychaete-host Trichotokara with the polychaete host Paralecudina and numerous environmental sequences, a clade with low (49%)
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8 support in ML (Fig. 1), now family Trichotokaridae. Trichotokaridae is sister to the polychaete host Polyplicariidae and further environmental sequences; despite weak support (0.62) this grouping of two families of species from marine polychaete hosts (plus one from a foraminiferan host) makes good evolutionary sense, so I make this whole clade a new order Vermigregarida, grouped with
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Arthrogregarinida as subclass Orthogregarinia. In the 196-alveolate tree (Fig. 3), Trichotokaridae support rose from 49 to 71% and 0.72; with ML they moved to become very weakly sister to the
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Selenidium bocardellae clade and with CAT sister to the unidentified environmental clade.
A CAT tree with 83 coccidiomorph sequences (Fig. 4), not just seven as in Fig. 3, also put
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Porosporoidea within Orthogregarinia, but as sister to core orthogregarines only. However, the Fig.
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4 analysis chains did not converge though each plateaued (maxdiff 0.376); the other chain for the same taxa showed only two differences: Porosporoidea was weakly (0.37) sister to Coccidea, not
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core orthogregarines, and Squirmidea sister to all Apicomplexa (not just Apicomonadea as in Fig. 4 or within Apicomonadea as in Fig. 3). Positions of Gregarinoidea and Trichotokara are also
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unstable to outgroup taxon sampling. Fig. 4 (both chains) put Gregarinoidea with the millipede-host Stenophora within archigregarines, i.e. within paragregarines not orthogregarines. In Fig. 4
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Trichotokaridae broke up into two: the relatively short-branch Paralecudina clade remained within the deepest branch of Orthogregarinia, but the extremely long-branch Trichotokara grouped instead with the somewhat long Oxyrrhis branch within dinoflagellates – almost certainly a long-branch artefact as it was not seen in any other trees and makes no evolutionary sense. Exclusion of Gregarinoidea from Orthogregarinia in Fig. 4 may be a long-branch artefact possibly caused by including so many long-branch coccidiomorphs (unlike Fig. 3). Figs 3 and 4 both imply that the crustacean-host Porosporoidea are more closely related to Terragregarina (grouped with them as Arthrogregarinida) than is the polychaete-host Trichotokaridae.
Detailed Orthogregarine Phylogeny
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9 All my trees (and many previously published ones) indicate that Orthogregarinia, especially core orthogregarines, are more closely related to Cryptosporidium and Rhytidocystidae than to archigregarines. In Fig. 3 Pyxinia groups with Hoplorhynchus and the ATCC 50646 clade with weak support (35% BS; 0.28 posterior probability) and Apicystis is sister to Mattesia with 100% or
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0.99 support (also 0.99% support in Fig. 4, both chains). In Fig. 4 the Pyxinia/Hoplorhynchus/ ATCC 50646 cluster also includes two additional environmental DNA sequences (EF100358/63);
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in Fig. 3 they are sister to Hoplorhynchus plus ATCC 50646 (0.44 support), but with ML moved just below the Hoplorhynchus/Monocystis last common ancestor. Addition of Pyxinia and Apicystis
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does not change the positions of the main gregarine clades on ML trees, though Apicomonadea
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joined Squirmidea as a second sister group to Dinozoa (low support). Within Terragregarina it induced some rearrangement, the Psychodiella/Ascogregarina and the
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Mattesia/Apicystis/Pseudomonocytis subclades being more strongly supported and now sisters (54%, 0.82 support). Moreover bootstrap support for some terragregarine backbone bipartitions
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dropped in Fig. 3: the highest ones (90, 87, 98, 79) became 74, 87, 84, and 55 respectively (Bayes: 0.84, 0.99, 0.99, -). Presumably the drop for the first bipartition is caused by adding Pyxinia, whose
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branch length is approximately twice that of ATCC 50646, such long branches being noted for a tendency to move somewhat too low in trees, whereas that for the fourth reflects the Fig. 3 evidence that Stylocephalidae are not sisters of Actinocephaloidea alone but of Actinocephaloidea plus Gregarinoidea. In marked contrast to the sparse distance and parsimony trees of Alarcón et al. (2011) and the ML, Bayesian and parsimony trees of Clopton (2009), which grouped Pyxinia strongly with Stenophora, I found no evidence for placing it there or with any marine gregarines. As Stenophora is a diplopod parasite and Pyxinia and Hoplorhynchus are insect (beetle or dragonfly) parasites (prior to Clopton (2009) both in Actinocephalidae), and major clades of gregarines show strong host group specificity (Rueckert et al., 2013) – even stronger than before in my heterogeneous trees, it is likely that the earlier grouping of Pyxinia with Stenophora on earlier less well sampled trees (only 51 or 27 gregarines respectively; 122 here) was a long-branch artefact.
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10 Transfer of Pyxinia from Actinocephalidae to Stenophoroidea (Clopton 2009) was probably therefore incorrect and is rejected here (Table 1). Its transfer then to Monoductidae may also be incorrect as two of the three genera of that family (Monoductus, Stenoductus) are millipede parasites like Stenophora, so their placement in Stenophoroidea may be correct; if so, their similar
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oocyst dehiscence, extruding cells in chains, may be convergent with Pyxinia – however until sequences are available we cannot rule out the less likely possibility that they also belong in
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Terragregarina and are related to Pyxinia. Within Gregarinoidea, Blabericolidae nest within
Gregarinidae in CAT trees (Fig. 3 and 4), but Gregarinidae are weakly holophyletic by ML (Fig. 1).
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Arthrogregarida from arthropods and oligochaetes are weakly sister to Vermigregarida from
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polychaetes, which also embraces an unidentified clade that includes a parasite or contaminant of the foraminiferan Ammonia (still wrongly annotated in GenBank as a foraminiferan though I
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discovered that error 19 years ago, and several publications have corrected it, notably Pawlowski et al. (1996)). The Polyplicariidae subclade of Vermigregarida was identified by Wakeman and
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Leander (2013) after I completed my analyses, so I was unable to include their three Polyplicarium sequences, which all grouped with clone CCA38 rather than CCI31. As Polyplicarium are
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symbionts of capitellid polychaetes, the grouping here of CCA38/CCI31 with the polychaete symbiont Trichotokaridae supports the new order Vermigregarida and suggests that like archigregarines they were ancestrally polychaete symbionts. Wakeman and Leander (2013) did not include either Trichotokaridae or the Ammonia symbiont subclade so did not reveal this relationship; their ML tree did however have insignificantly supported terragregarine and orthogregarine clades and grouped Squirmidea with Dinozoa but had no basal resolution, partly because only 1007 nucleotides and 79 Myzozoa were included.
Paragregarine Phylogeny Archigregarines and their relatives (probably derived Velocida, and Stenophora plus BOLA 566) form a clade (Paragregarea: 0.23 support Fig. 3) grouping not with orthogregarines but with
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11 Coccidia in Figs 3 and 4 or as sister to Coccidiomorphea plus Gregarinomorphea in Figure 2. BOLA 566 was moderately supported as sister to Stenophora in the best CAT trees (Figs. 2, 3), not sister to Selenidium terebellae as weakly in Rueckert and Leander (2010) or contradictorily to rhytidocystids in Leander and Ramey (2006), neither of which included Stenophora. The weakly-
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supported branching order of Selenidium terebellae and the Selenidium pisinnus/orientale clade in Fig. 1 of the accompanying paper (Cavalier-Smith 2014) was the same in Fig. 4 (CAT) but Fig. 3
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suggests instead that Selenidium pisinnus/orientale may be more closely related to the
Veloxidium/Urosporoidea clade than to the longer branch Selenidium serpulae clade. Though all my
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trees place Velocida within Selenidium, as do most but not all published trees, statistical support for
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this is so low that one cannot rule out the alternative possibility that Velocida are sisters to rather than derived from Archigregarinida.
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My trees confirm that two environmental DNAs from anoxic habitats once claimed to represent novel kingdoms (CCA5; LEMD119: Dawson and Pace 2002; Stoeck and Epstein 2003)
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are also gregarines as I showed previously using only 1044 nucleotide positions and many fewer gregarines (Cavalier-Smith 2004a). Berney et al. (2004) identified some of these as gregarines;
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more precise positions of many of them within the gregarine radiation were identified by Rueckert et al. (2011a), and largely confirmed here.
Phyletic distinctiveness of Squirmidea
Strikingly, Squirmidea (whose branch lengths are much shorter than for many gregarine clades, and are therefore unlikely to be grossly misplaced) never group with other gregarines or any Sporozoa on any of my trees or on any published trees with enough outgroups to judge adequately their affinities. On the ML tree (Fig. 1) they are weakly sister to Dinozoa; on the myzozoan CAT tree omitting long-branch taxa (Fig. 2), perhaps the most reliable indicator of their affinities, they are within Dinozoa, sister to the parasitic Perkinsea. CAT trees including long-branch gregarines place them contradictorily: weakly sister to Apicomonadea (Fig. 4) or all Apicomplexa (second
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12 chain for Fig. 4 taxa) or within Apicomonadea as sister to the Chromera/Voromonas/’Colpodella’ clade (Fig. 3: 0.3 support). Such clear, consistent exclusion of Squirmidea from Sporozoa in all trees could not have been detected on published ML gregarine-rich trees as non-sporozoan apicomplexa were always excluded; as the first taxonomically sparse ML tree insignificantly
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grouped them with long-branch archigregarines, they have been assumed to be gregarines even though there is no evidence for a sporozoan style apical complex (and morphological reasons to
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doubt that – see below) or any positive morphological evidence for inclusion in archigregarines.
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Myzozoan Phylogeny
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On Figure 3 Dinozoa are extremely weakly sisters to Sporozoa only, not sisters to all Apicomplexa, the non-sporozoan Apicomplexa being markedly paraphyletic, with Colpodella
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(=Alphamonas) edax (formerly grouped with Voromonas in Voromonadida: Cavalier-Smith and Chao 2004) plus an environmental DNA being sister to all other Myzozoa. Though basal branching
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of Myzozoa is too weakly supported for confidence in all details of this topology, the marked differences in several features of my heterogeneous trees indicate that it is unwise to take the deep
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gregarine branching order on homogeneous model trees too seriously, as the longest branches tend to group together as in Fig 1, I argue artefactually, which they certainly do not with the more realistic site-heterogeneous model. Use of heterogeneous models is advisable for future gregarine studies given the huge diversity in evolutionary rates amongst lineages. However, the remaining but relatively much smaller differences with taxon-sampling in Figs 3 and 4 emphasize the difficulty of resolving deep gregarine and myzozoan phylogeny from 18S rDNA alone and show that even CAT trees cannot iron out all inconsistencies among taxon samples for this evolutionarily exceedingly unclock-like molecule. Nonetheless, it is useful to compare these results with those of Fig. 1 to see which groupings are stable and which are not. Fig. 4 showed the tree from the chain that gave results most similar to Fig. 2; the internal phylogeny of all clades collapsed on Figs 3 and 4 was precisely the same as on Fig. 2, so they were internally stable.
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13 It is evident that both Vitrella and Chromera are simply photosynthetic apicomonads and do not merit a separate phylum; photosynthesis was clearly lost at least three times within Apicomplexa. I regard Chromerida as an order within Apicomonadea that on all my trees is sister to Voromonas plus species presently called ‘Colpodella’, but which for reasons to be explained
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elsewhere are generically distinct; by contrast Vitrella is more closely related to Colpodella (=Alphamonas) edax, which I now consider a true Colpodella. In Figs 2 and 4 the Colpodella
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(=Alphamonas) edax/Vitrella apicomonad subclade) was holophyletic (not paraphyletic like in Figs 1, 3) and the five hypersaline Lake Tyrell sequences formed a sister clade to marine Vitrella (not to
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the C. edax freshwater clade as in Heidelberg et al. (2013) fig. 5), this clade in turn being sister to
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the freshwater clade including C. edax. However Apicomonadea were paraphyletic with the Voromonas/Chromera subclade branching weakly more deeply than the C. edax/Vitrella subclade.
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Paraphyly of apicomonads was also shown by a chloroplast gene tree but the other way round (Janouškovec et al., 2012). If either is true, sporozoa definitely evolved from apicomonad ancestors.
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However branches within apicomonads are so close that multiple transcriptomes or genomes from both the Vitrella/C. edax and Voromonas/Chromera subgroups are necessary to resolve their
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topology and thus clarify the origins of sporozoa from such predatory flagellates.
Implications of Improved Gregarine Phylogeny for Classification The shallow nesting of neogregarines (e.g. Syncystis, Mattesia) within terrestrial gregarines in Figure 1 of the accompanying paper (Cavalier-Smith 2014) and on Figure 2 supports Grassé’s idea that neogregarines evolved from eugregarines by evolving merogony (Grassé 1953). However, their nesting shallowly in three different places amongst both ‘septate’ and aseptate eugregarines convincingly shows that neogregarines are multiply polyphyletic. Likewise the fact that on Figs 1-4 the classically eugregarine Velocida form a derived clade within archigregarines, that Stenophora is sister to archigregarines plus Velocida, and that other eugregarines are widely dispersed as the paraphyletic basal group of Terragregarina, or still more basally, suggests that classical
Page 13 of 60
14 eugregarines are also polyphyletic. Thus the even greater taxon and intramolecular sampling here strongly supports the recent conclusion that retention of the classical subdivision of gregarines into three orders (Archigregarinida, Eugregarinida, Neogregarinida: Adl et al. 2012; Grassé 1953) is ‘based more on convenience than phylogenetic relationships’ (Rueckert et al. 2011a). Sufficient
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information now exists to revise gregarine higher classification radically along sounder phylogenetic lines (Table 1), abandoning the polyphyletic neogregarines and eugregarines as taxa.
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Archigregarinida are probably paraphyletic, but retained as a morphologically well-defined group. Other classical gregarines make up a single new subclass Orthogregarinia comprising both aseptate
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and ‘septate’ gregarines subdivided into two new orders: Vermigregarida for two clades of aseptate
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polychaete gregarines, and Arthrogregarida for the remainder, predominantly inhabiting arthropods, rarely oligochaetes or molluscs, and largely ‘septate’. Table 1 includes in less detail other major
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taxa of the apicomplexan superclass Sporozoa as here revised.
A key feature of the new classification is the transfer of Cryptosporidium from Coccidia to
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class Gregarinomorphea, as a new order and subclass. The merits of such transfer were argued in detail by Barta and Thompson (2006) and need not be repeated. It is fully supported by multigene
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trees based on 10,753 amino acids that group Cryptosporidium with the actinocephalid terragregarine Ascogregarina with strong support (Templeton et al. 2010) and by the remarkable similarity of host-attachment structures in both groups (Valigurová et al. 2007). A second nongregarine group, previously treated as order Agammococcidiida that includes Rhytidocystidae, is clearly more closely related to both Cryptosporidium and orthogregarines than to archigregarines on all my trees and all sufficiently well sampled published trees; here I establish a separate subclass Histogregaria for this order and place it within Gregarinomorphea. As Archigregarinida never grouped with either Cryptosporidium or core orthogregarines (superfamilies Stylocephaloidea and Actinocephaloidea), which always group with Cryptosporidium to the exclusion of Archigregarinida, the conclusion that classical gregarines excluding Cryptosporidium and rhytidocystids are polyphyletic is now inescapable. Therefore a new sporozoan class Paragregarea
Page 14 of 60
15 is established to group archigregarines with those eugregarines that sequence trees consistently exclude (albeit usually with low support) from the Gregarinomorphea clade. Archigregarinida are marine non-septate, highly contractile, non-gliding gregarines that probably have apicoplasts, as organelles indistinguishable from apicoplasts are clearly visible in
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Selenidium hollandei and pendula (membranous vesicles in Figs 28-30, 39 of Schrével 1971). I postulate that apicoplasts were lost in the last common ancestor of Cryptosporidium and
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orthogregarines (arrow on Fig. 2), being apparently present in Rhytidocystis polygordiae and
sthenelais (Leander and Ramey 2006; Porchet-Hennere 1972). The only non-archigregarines
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consistently branching within archigregarines in all my trees are Velocida. Adl et al. (2012) put
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Veloxidium in archigregarines, but put Difficilina, Lecudina, Lankesteria, Pterospora and Lithocystis contradictorily in Eugregarinida. As all my and all published trees place the latter five
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genera (collectively a robust clade, superfamily Urosporoidea) together with Veloxidium with 100% support, it is cladistically wrong to include only Veloxidium within archigregarines. I therefore
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establish order Velocida for this extremely robust Veloxidium/urosporoidean clade. For a recently recognized clade of non-septate polychaete gregarines, e.g. Trichotokara,
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Paralecudina (Rueckert and Leander 2010), I establish a new family Trichotokaridae on account of their isolated phylogenetic position. Positions of the three longest branch taxa (Trichotokaridae, Porospororoidea, Gregarinoidea) are especially hard to establish on 18S rDNA trees probably because they have long unbroken stems, as is that of Stenophora with no close relatives breaking its much shorter branch. Though they all group together with ML (Fig. 1), this is probably a longbranch artefact as they occupy four separate positions on the heterogeneous model tree of Fig. 3, where all three longest branches group separately within the short-branch orthogregarines and Stenophora with, but not within, Archigregarinida. Though weakly supported (here 49%; 43% in (Rueckert and Leander 2010); 59% in (Rueckert et al. 2013), Trichotokaridae are a clade on all recent trees except Figure 4 where Trichotokara oddly moved outside Apicomplexa to join the long-branch Oxyrrhis. But the position of Trichotokaridae varies: in ML trees it was originally
Page 15 of 60
16 weakly sister to Urosporoidea (Rueckert and Leander 2010), to Urosporoidea plus Porosporoidea (Rueckert et al., 2011a); these contradictory positions or its placement in a mixed long-branch clade (Fig. 1) lack statistical support or biological rationale. By contrast the moderately supported grouping of the polychaete gregarines Polyplicariidae and Trichotokaridae on the evolutionarily
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more realistic heterogeneous tree (Fig. 3), irrespective of whether they have long or short branches, makes evolutionary sense, so I establish order Vermigregarida for this clade. Placing the three
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longest branch clades in distinct superfamilies or orders seems well justified. Their assignment to higher groups is more debatable, but done consistently with morphology and host preference and
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what I judge the best trees, but because of the low bootstrap support needs testing by sequencing at
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least another molecule like Hsp90 less subject to long-branch problems, and ideally by transcriptome analysis and multigene trees. I hope this revised classification stimulates such tests.
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The major long-branch clade comprising insect gregarines, for which I adopt the superfamily name Gregarinoidea used by Chakravarty (1960), consists of the invariably robustly
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related families Gregarinidae (ancestral) and Blabericolidae (derived). In the ML tree that did not include BOLA 566, this groups weakly with the millepede gregarine Stenophora. However CAT-
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model Figs 2-4 slightly more strongly group Stenophora with environmental sequence BOLA566, this clade appearing very weakly sister to Archigregarinida. The third subclade (crustacean gregarines: Porosporoidea) is sister to the short-branch terrestrial gregarines (Actinocephaloidea plus Stylocephaloidea) on both well-converged heterogeneous trees (Figs 2, 3). Gregarinoidea corresponds with terrestrial subclade II of Rueckert et al. (2011a) and Actinocephaloidea with their terrestrial subclade I. However, because Gregarinoidea grouped inside Actinocephaloidea on a later ML tree (Rueckert et al. 2013) they altered the labelling to include a subset of their subclade I in subclade II. However subclade II sensu Rueckert et al. (2013) is evidently a phylogenetic artefact of an ML tree with inadequate taxon (76 gregarines) and sequence (988 nucleotides not 1540 or 1577 as here) sampling. A later basally unresolved 1085-nucleotide tree did not put Gregarinoidea within Actinocephaloidea, and the authors reverted to the clade I and II labelling of Rueckert et al. (2011a)
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17 (Wakeman and Leander 2013b). None of my much better sampled trees place Gregarinoidea within Actinocephaloidea. Heterogeneous-model Figure 3 places them as sister to Actinocephaloidea, without Stenophora, with low (0.46) support. My trees have high 97% BS (Fig. 1) and 0.97 PP (Figs 2-3) support for holophyly of Actinocephaloidea (terrestrial gregarines sensu Rueckert et al.
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(2011) and Wakeman and Leander (2013b)), apparently a very robust clade that includes the new ATCC/stained glass window environmental clade (Cavalier-Smith 2014). The non-converging
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CAT-tree (Fig. 4) grouped Gregarinoidea with Stenophora and BOLA 566 very weakly within Archigregarinida, like ML, both I suspect long-branch artefacts. Stenophora and Pyxinia were
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omitted from all recent trees focusing on marine gregarines only.
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To exclude potentially misleading artefacts caused by long-branch gregarines Fig. 2 used 137 Myzozoa alone, chosen from the Fig. 4 taxa by excluding the longest-branch gregarines and
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long-branch Aggregata and Plasmodium, but supplemented by additional environmental apicomonad sequences (including five from Heidelberg et al. (2013) who obtained 1406
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apicomonad sequences from hypersaline Lake Tyrrell) and two extra Oxyrrhis sequences to break up their long branch. Archigregarines were represented only by the shortest branch Selenidium
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species. It also excluded a few of the fastest evolving sites that were used for Figs 3 and 4 trees to focus more on deep-branch topology. Fig. 2 converged well (maxdiff 0.128) and still placed gregarines and Squirmidea in three separate clades: with CAT Orthogregarinia was sister to Cryptosporidium (0.92 support), rather than to Rhytidocystidae as in other trees; and Squirmidea were not even in Apicomplexa, but weakly sister to Perkinsea (0.47 support, but much higher 0.85 support for including them in Dinozoa, not Sporozoa). Support was strong for a Stenophora/Selenidium clade (0.89) (new class Paragregarea) that was sister to Coccidiomorphea plus Gregarinomorphea; for Paragregarea to move to become sister to Orthogregarinia they would have to cross three nodes with support of 0.65, 0.92, and 0.92. Thus the heterogeneous tree with artefact-inducing long-branches excluded fairly strongly supports the conclusion that gregarines as previously understood are polyphyletic and evolved from smaller celled ancestors three times
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18 independently, once within Dinozoa and twice within Sporozoa. Virtually all other published trees have shown the same triphyletic ‘gregarine’ topology, but as they all included so many long-branch taxa and did not use heterogeneous trees or nearly so extensive outgroups, that topology was much more weakly supported and the conclusion of triple polyphyly much less convincing than it now is. I conclude that classical aseptate archigregarines are probably specifically related to but not
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ancestral to the ‘septate’ Stenophorida (not included in most published trees) but are ancestral to the
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also aseptate Velocida; they are apparently not ancestral to or sisters of orthogregarines.
One limitation of present knowledge of gregarine phylogeny is the absence of sequence data
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or ultrastructure for the non-septate blastogregarine Siedleckia (only four species), which unlike
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true gregarines show no evidence of szygy. Grassé (1953) considered them phylogenetically distinct from but distantly related to gregarines. However Levine et al. (1980) included them as a suborder
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within eugregarines primarily because of the absence of merogony, but that is not a sound reason as it does not distinguish them from Cryptosporidium or Histogregaria. As their anisogamy suggests a
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possible closer relationship with Cryptosporidium I prefer to follow Grassé (1953) and retain them as a separate order, as they might be a very early diverging group not within any known rDNA
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clade. Though their general body form and life style within polychaete guts is reminiscent of aseptate gregarines, the latter are probably polyphyletic, so I put blastogregarines incertae sedis in Sporozoa. Unsurprisingly, a preliminary report indicates S. nematoides branching with gregarines or Coccidia depending on taxon sampling and method (Diakin et al. 2012).
New myzozoan class Squirmidea
Previously Filipodium was placed in Lecudinidae, then transferred to Selenidiidae because it robustly grouped with Platyproteum vivax (formerly considered a Selenidium) as a clade here called squirmids, which once weakly grouped with S. terebellae on an ML tree within Selenidium (Rueckert and Leander 2009). Less poorly sampled ML trees (Rueckert and Leander 2010; Rueckert et al. 2011a,b, 2013; Wakeman and Leander 2013a, b) all agree with my even better
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19 sampled trees in placing squirmids more deeply than archigregarines, orthogregarines, Cryptosporidium, and coccidiomorphs (with both ML and heterogeneous CAT-model trees); the also weakly sampled ML tree of Rueckert et al. (2010) excluded Filipodium but grouped Platyproteum with a miscellaneous set of long-branch gregarines as sister to S. serpulae (but not in
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the corresponding Bayesian tree). There is no convincing sequence tree evidence for placing squirmids within Archigregarinida, and no cellular reason other than their pendular movement that
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could easily be convergent.
Indeed there is no evidence from ultrastructure concerning their apical complex – whether
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they have even have one or not, and if they do whether it is more like that of Apicomonadea than
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Sporozoa. Thus we do not know that they are even Sporozoa, as now defined as all Apicomplexa that descended from the first one that evolved a true symmetric conoid (Cavalier-Smith and Chao
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2004). Figs 1-4 all make it likely (but not certain) that they are not gregarines or even Sporozoa. It might be suggested that the consistent non-grouping of Squirmida with other gregarines here and in
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every other well-sampled tree is a long-branch artefact caused by the extremely diverse evolutionary rates within marine gregarines, and the absence of close relatives to Squirmida.
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However, their branch is much shorter than many branches that the CAT model places sensibly within archigregarines and orthogregarines and distinctly shorter than the long-branch Myrionecta branch that it correctly places within Ciliophora (Fig. 3; unlike ML which often misleadingly excludes it from Ciliophora leading to past treatment of its environmental sequences CCW75 and CCW100 and DH145-EKD11 as novel kingdoms (Dawson and Pace 2002) or phyla (LópezGarcía et al. 2001; Richards et al. 2005), until their ciliate nature was discovered (Bass et al. 2009)). We should therefore be open to the likelihood that squirmids are a truly independent myzozoan lineage that evolved large-celled gut parasites. Data from other gene sequences are vital to test that, but even now they are sufficiently distinctive genetically and morphologically to merit separation from all other apicomplexans as new order Squirmida.
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20 The limited thin section ultrastructure for Filipodium makes it possible that they are related to archigregarines as pellicle structure is similar except for the presence of hair-like projections. However the similarities consist simply of a wavy or shallowly folded surface (Hoshide and Todd 1996; Schrével 1971), which could easily have evolved independently, as all trees suggest. In
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contrast to some lecudinid archigregarines (e.g. Lankesteria) and some actinocephaloid orthogregarines, which independently lost longitudinal epicytic folds, there is no reason to postulate
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that squirmids had an ancestor with such folds.
Adl et al. (2012) retained Filipodium and Platyproteum in Archigregarinorida as did
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Schrével and Desportes (2013) ‘temporarily’ within Selenidiidae in Archigregarinorida. However,
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there is no sound phylogenetic evidence for that. Though the apical end of the squirmidean trophont is sometimes called a mucron, this term is best avoided as it begs the question of homology with
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that of gregarines for which there is no evidence, and strong contraindications. Its unusual asymmetry suggests that it will not be ultrastructurally the same as in archigregarines and
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orthogregarines, and that trophonts may lack conoids. Though that does not prove they are absent in all life cycle stages, as conoids can be lost secondarily in some developmental stages of some
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gregarines, the simplest interpretation of all my Bayesian trees (branching with Perkinsea Fig. 2; within Apicomonadea Fig. 3; as sister to Apicomonadea or Apicomplexa Fig. 4) and their rostral asymmetry is that Squirmidea have an asymmetric pseudoconoid like Apicomonadea and Dinozoa (see Okamoto and Keeling, 2014). I therefore establish a new class Squirmidea to contain Squirmida. I exclude Squirmidea from Sporozoa and tentatively place it with Perkinsea in Perkinsozoa, now a dinozoan superclass (Cavalier-Smith 2013b):
Class Squirmidea (in superclass Perkinsozoa Norén and Moestrup in Norén et al. (1999) stat. n. Cavalier-Smith 2013a) Squirmidea cl. n. Diagnosis: lumenal parasites of sipunculid intestines, some with intracellular phase; trophonts flattened throughout (Platyproteidae) or just at apical end (Filipodium); epicytic
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21 folds if present shallower than in Gregarinomorphea and underlain by longitudinal microtubules without complex fibrillar skeleton; unlike orthogregarines and Urosporoidea without gliding motility; trophonts with extensible asymmetric rostrum, not a symmetric apical mucron as in many gregarines. Merogony, sexual reproduction, presence or absence of apical complex, cilia, and
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rhoptries unknown. Etymol: squirm E., referring to their squirming motility. Sole order Squirmida
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ord. n. Diagnosis: as for class Squirmidea.
Filipodiidae fam. n. Diagnosis: highly contractile (bending, twisting) trophonts covered in hair-like
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projections (cytopili) supported by tubular extensions of the cortical alveoli, but without a
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microtubular or fibrillar axoneme; in some species at least cytopili arranged in longitudinal rows within every third or fifth longitudinal groove; cell apex flattened, often laterally expandable as a
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blunt cornered triangle wider than body. Type and sole genus Filipodium Hukue, 1939. Platyproteidae fam. n. Diagnosis: trophozoite with temporary fine transverse striations, associated
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with peristaltic motility, but no hair-like projections or longitudinal folds; bending and peristaltic shape changes; an extensible asymmetric flattened rostrum; with lumenal and intracellular stages;
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no evidence for merogony. Type and sole genus Platyproteum Rueckert and Leander, 2009.
Revised Gregarine Taxonomy
Classically gregarines were called Gregarinae (Haeckel 1866; Pritchard 1861), Grégarines (Claparède 1861), Gregarinidae (Lankester 1863) or Gregarinida (Pritchard, 1861) treated as a class in the animal kingdom even before Coccidea were recognized or Sporozoa invented to embrace both, though sometimes only treated as an order within Rhizopoda (Saville Kent 1880-1882). Other spellings are subclass Gregarinidea (Lankester 1885), Gregarinomorpha (Grassé 1953), or Gregarinia (Levine et al. 1980) or Gregarinasina (Perkins 2002 dated 2000) if ranked as subclass or Gregarinea when a class (Cavalier-Smith 1993a; Cavalier-Smith and Chao 2004). I here adopt the broader class name Gregarinomorphea and use the vernacular gregarinomorphs to refer collectively
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22 to all its members (i.e. gregarines, Cryptosporidium, and Histogregaria), and recommend that ‘gregarines’ is continued as in traditional usage for classical Gregarinea alone (i.e. orthogregarines and paragregarines), allowing their contrast collectively with Cryptosporidium, Histogregaria, and Squirmidea. As Gregarinomorphea include both many classical gregarines and two orders that are not gregarines, its suffix -morph usefully emphasises that it includes both gregarines and non-
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gregarine organisms (as in ‘nucleomorph’ which means related to, but not the same as, a nucleus);
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for the same reason I adopt Doflein’s class name Coccidiomorphea to include both classical
coccidia and organisms like Hematozoa and nephromycids that are not coccidia, but related to or
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derived from them. Diagnoses of new taxa follow:
Superclass Sporozoa Leuckart, 1879
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Gregarinomorphea Grassé, 1953
Histogregaria subcl. n. Diagnosis: apicoplast present; gametogony and merogony both absent;
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inhabit invertebrate tissues, not gut lumen. Etymology: histos Gk tissue, because of habitat. Sole order Histogregarida nom. nov. pro order Agamococcidiorida Levine, 1979; substitution because
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Levine’s name misleadingly implies they are coccidia. Cryptogregaria subcl. n. Diagnosis: epicellular parasites of vertebrate epithelia; initial round trophont becomes spherical whilst being surrounded by host lamellar extension of plasma membrane; after rhoptry and microneme discharge a single-membrane anterior vacuole develops that fuses with host plasma membrane after trophozoite plasma membrane fuses with that of host; the anterior vacuole membrane on the trophozoite cytosol side folds extensively to construct a feeder organelle with no homologues in Coccidomorphea, but some similarities to certain gregarines. Anisogamous; microgametes non-ciliate. Oocytes have 4 naked sporozoites without sporocysts. Unlike coccidiomorphs, Histogregaria, and archigregarines, lack apicoplast. Etymology: crypto Gk hidden: refers to hidden, epicellular location of the trophozoite (i.e. not in
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23 gut or coelomic lumen as in most mature gregarines) and to sole genus Cryptosporidium. Sole order Cryptogregarida ord. n. Diagnosis: as for subclass Cryptogregaria.
Orthogregarinia subcl. n. Diagnosis: Trophozoites ambicellular, i.e. partially embedded in host
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cell, partially extracellular (Cavalier-Smith 2004b). Aseptate or ‘septate’ gregarines with mucron but no apicoplasts; invertebrate hosts (arthropods or annelids); gamonts of similar size. Unlike
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archigregarines or Platyroteum, usually not strongly flattened. Well-developed longitudinal epicytic folds in gut lumenal forms; motile by gliding, without dramatic contractions seen in
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Archigregarinida, Veloxidium, and squirmids. Etymol: ortho Gk straight, correct, true, as includes
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the standard originally described gregarines (Gregarina Dufour, 1828).
Vermigregarida ord. n. Diagnosis: Trophozoites aseptate, with densely packed, deep epicytic
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folds; mucron often elongate, cylindrical and hairy, sometimes squat; typically in gut lumen of polychaete hosts; szygy lateral or end-to-end; gamonts without mucron; gametes, sporozoites,
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oocysts or merogony not described; inhabit polychaete guts or rarely foraminifera. Etymol: vermis L. worm emphasises that known hosts are generally polychaete worms.
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Trichotokaridae fam. n. Diagnosis: Polychaete-infecting gregarines, commonly with elongate hairy mucron. Type genus Trichotokara (Rueckert and Leander 2010). Other genus Paralecudina Rueckert et al., 2013.
Polyplicariidae fam. n. Diagnosis: Polychaete-infecting gregarines, with blunt non-hairy mucron, and high density of longitudinal epicytic folds (4-5/m). Type and sole genus Polyplicarium (Wakeman and Leander 2013a).
Arthrogregarida ord. n. Diagnosis: ancestrally ‘septate’ gregarines living in gut lumen of arthropod hosts, plus derived parasites of other body cavities or tissues in insects and oligochaete annnelids, and some simplified aseptate forms. Mucron ranges from short to elongated as an epimerite. Ancestrally without merogony but some morphologically reduced genera secondarily
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24 evolved it. Szygy usually late and caudofrontal, sometimes frontal. Male gamont often uniciliate, never biciliate. Etymol: arthro Gk joint as all hosts are jointed/segmented animals; also refers to the trophonts’ joint-like appearance in the ‘septate’ majority. Porosporina subord. n. Diagnosis: ‘septate’ or aseptate lumenal parasites of crustacean (rarely
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mollusc) intestines; syzgy early, usually caudofrontal, sometimes involving more than two cells; merogony absent. Etymol: Named after type genus Porospora Schneider, 1875 of the oldest
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included family. Contrary to Rueckert et al. (2011a) the sole superfamily cannot be called
Cephaloidophoroidea if it includes Porosporidae, because superfamily names must be based on the
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oldest included family and attributed to the authors of that family not the person(s) raising it to that
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rank (article 23.3 of the International Code of Zoological Nomenclature, applicable to Sporozoa). Terragregarina subord. n. Diagnosis: ancestrally ‘septate’ insect, arachnid or oligochaete parasites
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plus some secondarily simplified non-sepate forms; merogony ancestrally and commonly absent but present in some insect parasites (former neogregarines). Etymol: terra L. earth, emphasizes that
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hosts are terrestrial (or as nymphs in freshwater) not marine. Superfamily Actinocephaloidea Léger, 1892 stat. n. Diagnosis: szygy frontal, late; insect,
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arachnid, myriapod hosts. Phylogenetically defined as the clade containing all gregarines more closely related to Monocystis, Ascogregarina, and Mattesia than to Stylocephalus or Gregarina.
Class Paragregarea
Paragregarea cl. n. Diagnosis: aseptate marine gut-dwelling or coelomic gregarines; annelid, nemertean, mollusc, deuterostome hosts; trophozoite with longitudinal or transverse folds or knoblike projections and surface crenulations; often with marked bending or thrashing motility; mucron short, pointed or blunt. Etymol: para Gk but, against, contrary to; emphasizes they are probably not directly related to orthogregarines. Velocida ord. n. Diagnosis: Aseptate. Trophont either contractile with bending and twisting motility, but neither gliding nor with longitudinal folds, or else non-contractile with longitudinal
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25 folds and gliding motility; mucron round or nipple-like. Etymol: velox, velocidis L. rapid because of rapid twisting of Veloxidium and gliding of Urosporoidea. Velocid- is the combining stem form that grammatically ought to have been used for Veloxidium. Urosporoidea Léger, 1892 superfam. n. Diagnosis: trophont aseptate, with longitudinal folds,
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knobs, and gliding motility; unlike Veloxidium, no bending or twisting motility; mucron round. Veloxidioidea superf. n. Diagnosis: as for sole family: Veloxidiidae fam. n. Diagnosis: trophont
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contractile with bending and twisting motility; lacking gliding motility or longitudinal folds. Type and sole genus Veloxidium Wakeman and Leander, 2012.
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Evolution and Taxonomy of Coccidiomorphea
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Stenophoridae Léger and Dubosq, 1904 (e.g. Stenophora).
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Stenophorida ord. n. Diagnosis: myriapod hosts; ‘septate’. Etymology: Named after sole family
Removal of Cryptosporidium makes Coccidiomorphea holophyletic on sequence trees.
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Recently it was shown that the sea-squirt symbionts Nephromyces (Saffo 1981) and Cardiosporidium (order Nephromycida (Cavalier-Smith 1993b)) belong in Sporozoa and are
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phylogenetically related to piroplasms (Saffo et al. 2010). Fig. 2 strongly confirms that but contradicts their non-CAT ML and Bayesian trees in showing coccidians as paraphyletic, with the nephromycid/piroplasm clade as sister to Eimeriida only not to all Coccidia. However, Saffo et al. (2010) did not include Haemosporida, so could not exclude an alternative relationship with them. Fig. 4 therefore included Haemosporida and other previously excluded coccidiomorphs to clarify their position. Nephromycids remain sister to piroplasms, with slightly reduced but still reasonable support and their joint clade remains sister to Eimerida only but with sharply reduced support. Haemosporida, though currently grouped with piroplasms as class Hematozoa, group with weak but significant support with adelinid coccidians not with piroplasms (Fig. 4). This makes it possible the grouping Haemosporida and piroplasms as Aconoidasida (Mehlhorn et al. 1980) or Hematozoa (Vivier 1982) was artificial and shared features are convergent. On present evidence retaining a
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26 separate class uniting piroplasms only with Haemosporidia is unjustified. Multigene trees strongly group piroplasms as sister to Eimeriida, but as transcriptomes are unavailable for Adeliida do not reveal whether they are nested within Coccidia or their sisters. Until they are it may be premature to abolish Hematozoa, retained here as a subclass of Coccidiomorphea to which Nephromycida is added (Table 1); the original rationale for Aconoidasida embracing piroplasms and Haemosporidia
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(Mehlhorn et al. 1980) however is defunct, as some ookinetes in Haemosporidia have conoids
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(Paterson and Desser 1989). A new superorder is needed for piroplasms plus nephromycids:
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Aconoidia superord. n. Intracellular parasites of ascidian haemocytes with posterior biciliate phase
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or non-ciliate parasites of mammalian blood cells with microtubule-supported projections from gametes; conoid absent. Etymol: a- without + conoid; Aconoidasida is a misleading name for
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Hematozoa as Haemosporidia often have conoids (Paterson and Desser 1989), never seen in piroplasms and nephromycids (Ciancio et al. 2008).
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Aggregata, initially considered a gregarine, has very long branches on 18S rDNA trees so was excluded from both coccidians and gregarines and left incertae sedis in Apicomplexa by Adl et
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al. (2012). However, its elongate triciliate microgametes are typical for Coccidia, and Fig. 4 strongly supports its grouping with Hepatozoon within Adeleida, as found by Castellanos-Martínez et al. (2013), so it clearly belongs in that order. Eimeria and Goussia are both found in two highly divergent clades that are genetically more different from each other than is either from Sarcocystidae, partly because rDNA has evolved much faster in one of these clades (Figs 2, 4). Clearly both genera have been overlumped and inaccurately split; Calyptospora forms one of three subclades within the longer branch Eimeriidae clade and probably does not deserve its own family. Important phylogenetically are Coelotrophiida (=Protococcidiida), e.g. the polychaete parasites Coelotropha and Grellia with early intracellular and later coelomic phases, for which I once established a separate class Coelotrophea (Cavalier-Smith 1993a). Sequence trees are needed to test their unity and phylogenetic position: a preliminary report of an ML tree suggests that they
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27 may be nearer to apicomonads than to Sporozoa (Diakin et al. 2012), but sequences are unavailable and could not be included here. However, their biciliate gametes do not resemble apicomonads (Bardele 1966; Porchet-Henneré 1967), and are distinct from those of Coccidia and Haemosporida in not being elongated with strongly reflected backward pointing centrioles; unlike Coccidia the centrioles are well separated and not closely attached behind a pointed
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microtubule-supported perforatorium. Moreover, as their complete conoid is typically
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coccidian-like with two intraconoid microtubules (Bardele 1966; Porchet-Henneré 1971),
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from Coccidia and Hematozoa merits a new third subclass:
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Table 1 conservatively keeps them in Sporozoa and Coccidiomorphea. Microgamete distinctness
Coelotrophia subcl. n. Diagnosis: Gametogony and sporogony extracellular, unlike other
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coccidiomorphs. Biciliate microgametes near-spherical, not elongated as in Coccidia and Haemosporida, not aciliate as in piroplasms; mitochondrion lies between anteriorly indented
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nucleus and the centrioles, not laterally as in Coccidia and Haemosporida. Microtubule-supported microgamete perforatorium absent, unlike Coccidia. Etymol: Named after sole included order
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Coelotrophiida.
Changing views of gregarine and sporozoan evolution The large and predominantly extracellular gregarines were known long before the small-celled intracellular coccidia. Soon after Leuckart (1879) united both groups as Sporozoa, Bütschli (1880-1882) suggested that Sporozoa evolved from flagellates by losing cilia as an adaptation to parasitism. Sporozoa are united primarily by having a non-flagellate cell invasion stage, the sporozoite, with radially symmetric apical cytoskeleton adapted for penetrating the host plasma membrane. Bütschli thought they might have evolved from euglenoids, now clearly not so. More presciently, the discovery of the coccidian-like syndiniid intranuclear parasite Coccidinium by Chatton and Biecheler (1934) led Chatton to suggest that evolution of multiple fission by an
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28 intracellular parasitic dinoflagellate could have led to the origin of Coccidia and Sporozoa generally. Levine favoured perkinsids as the closest flagellates to Sporozoa, and recent thinking has been dominated by his view (Levine, 1985) that gregarines are ancestral to coccidians (e.g. Leander 2008; Perkins et al. 2002 dated 2000; Schrével and Desportes 2013; Vivier and Desportes
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1990) even though I am unaware of any compelling explicit arguments for it. Grassé drew an analogy between bodonids and coccidian male gametes, but was inclined to think similarities
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convergent and regarded gregarines as two clades diverging from a common sporozoan ancestor and did not postulate that one evolved from the other. Vivier and Desportes (1990) wrote
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‘gregarines, which are exclusively parasitic on invertebrates, surely evolved earlier than other
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apicomplexan classes, which parasitise vertebrates.’ However that argument is fallacious because it confuses secondary invasion of vertebrate hosts with the primary origin of each group; coccidia and
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Hematozoa both parasitize invertebrates too so could be just as old as gregarines or even older. Anyway, if we now accept Cryptosporidium as a gregarinomorph then one gregarinomorph lineage
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did invade vertebrate hosts. Gregarines were still called ‘the early Apicomplexa’ by Shrével and Desportes (2013) even though their schematic tree showed gregarines, coccidia, and
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Cryptosporidium branching from the same point. The true flagellate ancestors of Sporozoa became clearer when other French protistologists, Brugerolle and Mignot (1979), showed that a myzocytotic predatory flagellate then called Spiromonas perforans (transferred by Patterson and Zölffel (1991) to Colpodella, then by me to Chilovora (Cavalier-Smith and Chao 2004)) had ultrastructure suggesting an intermediate position between dinoflagellates and Sporozoa, and was most unlike the bodonid Dimastigella described under the name Spiromonas angusta (MacDonald and Darbyshire 1977). Discovery of another myzocytotic predator (Foissner and Foissner 1984) led Krylov and Mylnikov (1986) to make a subclass Spiromonadomorphina and order Spiromonadida within Sporozoa for these free-living predators and Cavalier-Smith (1991) to place Colpodella within Apicomplexa and formulate the concept of alveolates as ancestrally photosynthetic eukaryotes with cortical alveoli and tubular
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29 mitochondrial cristae, and formally establish infrakingdom Alveolata to embrace ciliates, dinoflagellates, Apicomplexa, and protalveolates (Colponema). Earlier the haploid dinoflagellates and Sporozoa were formally grouped as Miozoa (Cavalier-Smith 1987), now a phylum with subphyla Protalveolata (sole class Colponemea) and Myzozoa (Cavalier-Smith 2013a,b).
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Spiromonadida was renamed Colpodellida by Cavalier-Smith (1993) (as Spiromonas is confusing and probably invalid) and the class Apicomonadea outside Sporozoa (but within Apicomplexa) was
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established for colpodellids and perkinsids, it being proposed that Sporozoa and dinoflagellates evolved independently from such myzocytotic apicomonads. When the first sequence tree to
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include a colpodellid grouped it, but not Perkinsus, with Sporozoa (Cavalier-Smith 2000) I
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explicitly proposed free-living Colpodella-like myzocytotic predatory flagellates as the ancestors of Sporozoa. After more colpodellid and perkinsid sequences became available Perkinsus was
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removed from apicomonads when it become clearer that Perkinsea was the deepest branch within Dinozoa and Apicomonadea the deepest within Apicomplexa (Cavalier-Smith and Chao 2004).
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Now sequences of both rDNA genes for Colponema place it as a sister clade to Myzozoa (Janouškovec et al. 2013) in accord with that classification; I mistrust their concatenated 6-gene
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tree that put it weakly as sister to all other alveolates, as the individual trees for the four proteins (the same used in the accompanying paper: Cavalier-Smith 2014) were confused and none of the six genes individually placed it so distantly.
The first explicit suggestions of how sporozoa evolved from an apicomonad emphasized that most gregarines are not truly extracellular but ambicellular (Cavalier-Smith 2004b; CavalierSmith and Chao 2004). I argued that the conoid, composed of conically arranged spiral tubulin fibres arranged like a compressed spring (Hu et al. 2002), was an innovation for cell penetration, and that the extracellular condition of bigger gregarines was developmentally and evolutionarily secondary. At least in Toxoplasma conoidal tubulin fibres are not microtubules but sharply curved protein sheets, comma-like in transverse section (Hu et al. 2002). Now that my CAT trees make it likely that orthogregarines and paragregarines evolved independently from early non-gregarine
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30 Sporozoa we can see that they constitute two somewhat distinct organismal types. Trophozoites of orthogregarines may start off as intracellular parasites but those inhabiting intestines spend most of their growth phase as ambicellular. From their broad distribution on the trees such ambicellular gut parasites are clearly the ancestral state for orthogregarines, so this lifestyle imposed the initial
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selective forces on their origin, outlined previously (Cavalier-Smith 2004b; Cavalier-Smith and Chao 2004). Orthogregarines secondarily invaded other habitats, notably the coelom of oligochaetes
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and then lost ambicellularity and became secondarily purely extracellular trophozoites. By contrast archigregarines are not ambicellular but always initially intracellular and in some but not all cases
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become truly extracellular by extrusion into the gut lumen before growth finishes. Some Selenidium
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even undergo multiple fission within the intestinal epithelium.
This basic difference in lifestyle between archigregarines and orthogregarines is consistent
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with their having evolved large trophozoites independently. The fact that only orthogregarines are truly ambicellular explains why only they have evolved true epimerites that anchor the partially
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embedded cell tip into the epithelium and are autotomised when growth finishes, making them truly extracellular only when differentiating into gamonts. The constricted condition (traditionally
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misleadingly called ‘septate’; they have a constriction not a septum) is also restricted to ambicellular arthropod-infecting orthogregarines and Stenophora, the constriction being arguably an adaptation to facilitate later autotomy and deutomerite release. The facts that ‘septate’ orthogregarines do not all cluster together on Fig. 3 supports Grassé’s (1953) judgement that septation has only weak systematic value; yet even though he rejected order Septata (Lankester 1885), Levine (1885) resurrected it as suborder Septatorina), whence it has lingered on as suborder Septatorina in the latest revisions of both of The Illustrated Guide to Protozoa (Clopton 2002 dated 2000) and more surprisingly of Grassé’s own Treatise chapter (Schrével and Desportes, 2013). It is now evident that Septatorina are polyphyletic, most comprising the ancestrally ambicellular orthogregarines that inhabit arthropod guts (a paraphyletic subgroup of septates) here placed in the new order Arthrogregarinida that also includes non-septate gregarines that clearly
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31 evolved from them (e.g. Monocystis) by switching life style to extracellularity. Stenophorids apparently evolved septation independently through convergent adoption of the ambicellular lifestyle in arthropod (millipede) guts (though protein sequence data are sorely needed to check they are really sisters to core paragregarines; relationship instead with septate Gregarinomorphea is not
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strongly excluded); however unlike ‘septate’ orthogregarines their mucron is retractible (Grassé 1953) and thus not a true caducous epimerite. Aseptatorina Chakravarty, 1960 (= Acephalina von
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Kölliker, 1848; Haplocyta Lankester, 1885) by contrast is a polyphyletic mixture of archigregarines and velocids which are never ‘septate’ and derived orthogregarines which lost the constriction
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secondarily.
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Grassé (1953) argued that the division of gregarines into schizogregarines with multiple fission unrelated to sex (merogony) and eugregarines without merogony by Léger (1900) was no
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less artificial; he thought that some gregarines had evolved merogony secondarily (his order Neogregarinida), whereas others had it primitively (his Archigregarinida). Sequence phylogeny
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shows he was right about the secondary nature of neogregarines, but that merogony evolved several times independently in different groups of orthogregarines (Fig. 3). Thus neogregarines are
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polyphyletic and the distinction evolutionarily too trivial to be used at ordinal level. Therefore the new phylogenetic classification (Table 1) abandons Neogregarinida, Eugregarinida, Aseptatorina, and Septata/Septatorina (all polyphyletic). Grassé’s ideas about archigregarines and merogony remain untested as there are no sequences for merogonic archigregarines (Merogregarinidae, Exoschizonidae), only for Selenidium of Selenidiidae, a family that Levine transferred from archigregarines to eugregarines (Levine 1971). Contrary to Levine, recent discussions of gregarine sequence phylogeny have all referred to Selenidium as an archigregarine (Rueckert et al. 2010; Rueckert and Leander 2009), followed here as I regard the presence or absence of merogony as evolutionary relatively trivial changes compared with cell morphology. Like Grassé (1953) but unlike recent Levine-influenced compilations (Perkins et al. 2002 dated 2000; Schrével and Desportes 2013; Vivier and Desportes 1990) I include Selenidiidae within archigregarines. It is
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32 noteworthy that Grassé (1953 p. 348) remarked that the most archaic gregarines are Selenidiidae, Urosporidae, and Lecudinidae – essentially Archigregarinida plus Urosporoidea (Table 1), i.e. all paragregarines then known. Sequence trees now confirm that he rightly singled out these groups as differing from other gregarines, as all three form a completely non-septate paragregarine clade, but
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suggest that he was wrong in assuming gregarine monophyly. Merogonic archigregarines might possibly be primitive for Paragregarea, but there is no reason to think they are ancestral to
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orthogregarines. The intracellular nature of merogonic multiple fission in Merogregarina reflects the fact that unlike the larger celled non-merogonic archigregarines trophozoites remain
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intracellular. Further growth prior to syngamy that forces older trophozoites of non-merogonic
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archigregarines to become extracellular is arguably a derived character for Sporozoa. However, that does not mean that non-merogonic archigregarines or orthogregarines
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evolved from merogonic ancestors by losing merogony. More likely both evolved divergently from non-merogonic ancestors independently of several lineages that independently evolved merogony.
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Cell biologically, multiple fission, whether preceding syngamy (gametogony), following it to produce sporozoites (sporogony) or in between periods of vegetative growth, is the same process.
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Multiple fission cell cycles are widespread in protists, especially in parasites and in free-living unicellular algae like Chlamydomonas (Craigie and Cavalier-Smith 1982) and must therefore be easy to evolve. They can evolve from normal binary fission cell cycles merely by imposing a reversible block to division, temporarily uncoupling normal size-dependent division control so cells become manyfold (not just two-fold) larger than daughter cells before dividing (Cavalier-Smith 1980). A round of multiple fission can be introduced at any point in a life cycle merely by relieving that block by some developmental signal – a simple switch, nothing complicated like evolving a conoid or myzocytosis. Clearly multiple fission had already evolved in the apicomonad ancestor prior to the origin of Sporozoa by evolving the conoid and non-ciliate sporozoite. Apicomonads grow manyfold whilst feeding and then the large cell encysts and undergoes multiple fission to make new flagellates; an adaptation to maximising food uptake and the number of daughters during
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33 a single feeding bout, as the rareness of food encounters makes it important to get as many offspring as possible from each. Commonly a division cyst yields four daughters. Almost nothing is known about apicomonad sex, so we cannot say whether their fission is gametogony, merogony or sporogony or more than one. Anteriorly connected quadriciliate cells of Algovora (=Colpodella) pugnax were suggested to represent syngamy (Simpson and Patterson
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1996), which may be correct if mitotic division occurs only within a cyst; it is less likely that it is a
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freak misdivided cell (common in flagellates) as one would expect division to begin anteriorly and sister hemicells to be posteriorly linked. However, this species could actually be a dinozoan not an
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apicomonad (Cavalier-Smith and Chao 2004), making its relevance to sporozoan origins unclear.
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Nonetheless it is likely that some apicomonads still are sexual; the one ancestral to sporozoa must have been and would probably have had both post-feeding and post-syngamy multiple fissions,
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probably into four daughters, and thus have two rather than three multiple fission stages in its life cycle. Almost certainly it would have been isogamous and copulating freely in water not within a
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cyst; thus anisogamy and gamont szygy were sporozoan innovations. I suggest that both evolved in the ancestral sporozoan. Contrary to what is sometimes stated, enclosure of gamonts within a cyst is
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not a unique gregarine feature but also occurs in some adelinid coccidians, even though the cyst is tenuous. Therefore it was probably lost by Eimeriida and Hematozoa.
Ciliary evolution, cell and genome size, and the polyphyletic origins of gregarines Sequence trees strongly indicate that gregarines are polyphyletic, not paraphyletic as commonly assumed since Levine (1985). Three arguments from cell evolution also make it improbable that gregarines are ancestral to coccidiomorphs. First consider ciliary evolution. Myzozoan cilia are distinctly different from the ancestral alveolate state represented by Colponema with normal triplet centrioles and two 9 + 2 cilia with equal sized central pair microtubules. In virtually all Myzozoa, unlike any other eukaryotes, one central pair microtubule is narrower than the other; though most clearly obvious and only explicitly previously noticed in Perkinsea
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34 (Leander and Hoppenrath 2008) it is also true of dinoflagellates, apicomomonads and Coccidiomorphea. A few images suggest equal sized central microtubules (e.g. Speer and Danforth 1976 Fig. 7) suggesting an occasional reversal to the ancestral state, presumably by losing or modifying some special protein(s) needed for making the narrow microtubule. With the
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notable exception of Aggregata, which has normal triplets (Heller 1970), Apicomplexa have generally reduced their ciliated centrioles largely to doublets (true of apicomonads and Sporozoa:
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Brugerolle and Mignot 1979; Colley 1967); only at the extreme base of the Chromera centriole can one clearly see triplets (Oborník et al. 2011). Sporozoa further modified their mitotic
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centrioles only by blocking centriolar development to yield short nine-singlet procentrioles capable
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of further duplication when ciliary development was suppressed during the evolutionary origin of sporozoites with complete conoids. Orthogregarines apparently lost the centre pair of their cilia and
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have 9 + 0, 6 + 0, or 3 +0 cilia, with only slightly reduced 8-doublet centrioles (Schrével and Besse 1975; Schrével and Desportes 2013). The 9 + 2 cilia of Coccidiomorphea could not have evolved
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from them. Gregarines have either no cilia or a single posterior cilium only in male gametes, so the
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ancestors of both orthogregarines and paragregarines lost the posterior cilium and its associated centriolar roots. Thus neither gregarine group can be ancestral to Coccidiomorphea, most of which have two cilia; Piroplasmida lost cilia altogether, but their nephromycid sisters have two posterior 9 + 2 cilia and apparently a two-microtubule anterior ciliary root and posterior directed multiplemicrotubule root as well as a dorsal microtubule fan in their microgametes (Cianco et al. 2008). Eimeriid and adelinid coccidian microgametes have two posterior-pointing cilia arranged as in apicomonads with an amorphous connector between them, but their cell apex is modified; nonetheless the dense apical perforatorium is some species clearly includes a short anterior fan of microtubules. Some have a third cilium, probably a multiply derived state. Several Eimeria and the adelinid Aggregata have a posterior microtubular root emanating between the two primary centrioles, but this root has several times been misinterpreted as a third or fourth recurrent
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35 intracellular ciliary axoneme (e.g. Scholtyseck et al. 1972). Eimeria perforans unusually has a third cilium (Scholtyseck 1965); Colley (1967) thought Eimeria nieschulzii also has a third intermediate ciliary stub, which may be correct even though some of the structures he labeled basal bodies are probably the amorphous centriolar connector not a centriole; however the structures labeled third flagellum might instead represent the posterior root of five singlet microtubules plus
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dense fibrillar material found there in Eimeria truncata (Gajadhar and Stockdale 1986) or 8-10
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microtubules in Eimeria magna (Speer and Danforth 1976) or four microtubules in Sarcocystis (Vetterling et al. 1973). These singlet microtubules cannot be relics of a hypothetical third cilium
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as Vetterling and others assumed. They are associated with dense material and probably are
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homologous with the most prominent right posterior root of apicomonads (R1) whose two centrioles are connected and oriented in the same way, and therefore provide evidence that Coccidia
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evolved from ancestors having two connected centrioles and a prominent posterior microtubular root. Haemosporida have either two posterior cilia (Haemoproteus: Bradbury and Trager 1968)
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with highly attenuated/modified centrioles or one (Plasmodium) with no trace of microtubular roots. It seems clear that the ancestral coccidiomorph had two cilia later reduced in one lineage to one, in
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another to zero, and increased in two others to three. In addition to a centriolar connector it would have had a slender anterior ciliary root (arguably R3), perhaps of 2 or a fan of about 8 microtubules, and a posterior root with several microtubules. Therefore Coccidiomorphea could not have evolved from either group of gregarines, which have a single cilium, which as primary differentiations after a non-ciliate stage would be developmentally homologous with the anterior cilium of biciliate eukaryotes even though they may point and swim posteriorly (see Cavalier-Smith 2013a for this argument) and should therefore lack homologues of the posterior roots associated with the older cilium. According to this argument, Coccidiomorphea ancestrally retained ciliary and centriolar transformation and features specific to the posterior cilium and its roots, but gregarines probably lost such features independently in Orthogregarinia and Paragregarea. Unfortunately ciliary
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36 ultrastructure is unknown for Paragregarea, most of which lack cilia, so we cannot say whether it is more like that of Orthogregarinia or Coccidiomorphea. A second argument relates to cell and genome size, a key difference between gregarines and Coccidiomorphea. As previously emphasized, coccidian cells are much smaller than the ancestral
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apicomonads and have correspondingly miniaturized their genomes, whereas most gregarines are far larger than their host cells and have greatly magnified their cells, nuclei and genome sizes
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compared with apicomonads or coocidia (Cavalier-Smith 1985 p. 123); thus their chromosomes and genomes are generally thousands of times more massive than those of Coccidia as first emphasized
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by Goldschmidt (1955) which led him to doubt that DNA can be the genetic material, because he
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did not envisage that in eukaryotes it actually has both genic functions and separate structural functions that make the amount of non-genic DNA highly dependent on cell size (Cavalier-Smith
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1978, 1982, 1985, 2005). These correlations reflect the universal eukaryotic scaling laws between cell, nuclear, and genome size, and have been explained by the theory of optimal cytonuclear ratio
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coupled with the skeletal function of all nuclear DNA whether it is genic or not (Cavalier-Smith 2005). The immense size of gregarines was favoured by their gut luminal life style, which offers an
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essentially unlimited food source allowing much more protracted cell growth than is possible for a predator of single cells, whereas the tiny size of coccidiomorphs is required for rapid intracellular reproduction within a similarly spatially restricted food source. The large-celled ambicellular or maturely extracellular habit would lead to an inexorable increase in cell size and in associated genome size through thousands of independent duplications of non-genic nucleoskeletal DNA. I argue that this would sooner rather than later become irreversible, because gregarine cells would become too large to shift lifestyle to purely intracellular location; large reductions in genome size would be impossible in a single step, as that would require thousands of concerted deletions, so gregarines would be trapped in the largely extracellular trophic condition and could not secondarily become minute intracellular parasites like coccidia; eventually they would pass a point where they could not even fit into single animal cells.
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37 The ciliary evolution and cell and genome size arguments both reinforce Grassé’s view that gregarines and coccidians are independent lines. It is historically incorrect to attribute to him the recent dominant Levinean view that coccidia evolved from gregarines. It is now clear that both coccidia and gregarines in the classical sense are polyphyletic. That is hardly surprising because the
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small cell intracellular versus large cell extracellular dichotomy is very simple, involving transitions crossed many times during protist evolution, so one should not expect organisms sharing these
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features to be phylogenetically coherent unless they share unique morphological features, which is not the case within Sporozoa. Relying too much on them led to the small-celled intracellular
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Cryptosporidium being wrongly classified with coccidians rather than orthogregarines. Interestingly
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unlike Coccidia, but like Piroplasmida and some but not all Haemosporida, it has lost the conoid (Matsubayashi et al. 2008). Loss of the apicoplast in the common ancestor of Cryptosporidium
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and orthogregarines is a third reason why this gregarinomorph subclade cannot have been ancestral
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to Coccidiomorphea, many of which have apicoplasts.
Multiple fission in apicomplexan evolution
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The third traditional dichotomy between coccidia and gregarines lies in gametogony. In all Sporozoa gametes are anisogamous, unlike virtually all other protists except a few volvocalean green algae, whose life cycle involves multicellular coenobia so they are really multicellular, not true unicellular protists. In gregarines male and female gamonts divide a similar number of times, whereas in classical coccidia female gamonts do not divide. This dichotomy also has a simple adaptive explanation that allows it to evolve easily more than once. I argued above that szygy evolved in the ancestor of all sporozoa and was independently secondarily lost by Eimeriida and Hematozoa. The enclosure of gamonts by a cyst (unique in protists) inevitably led to anisogamy with respect to ciliary development because as syngamy occurred within a cyst it would be wasteful for both gametes to develop cilia and better for one to focus its resources on food storage for zygote development; only one needed to make cilia to find the other. The presence of the cyst however
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38 radically changed selective forces relating to cell size anisogamy (see Bulmer and Parker 2002; Parker 1978). In that conventional argument a major driver of cell miniaturization to make sperm is the benefits of fertilizing ever more genetically different eggs. But by being locked up within the cyst wall with access to only one female genotype this driver for evolving many more divisions into
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ever smaller sperm is absent in gregarines, so there is no benefit of having more multiple fissions than the female. Because of that constraint gregarines kept the ancestral condition of an equal
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number of fissions in both sexes. Those Coccidiomorphea that lost the zygy cyst probably did so because, unlike in the large gut where there can be many sexual partners, a strictly intracellular life
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style (specially in small host cells) greatly reduces the chances of finding a genetically different
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partner in that cell, so multiple divisions have to precede partner association (making merogony almost a sine-qua non for an intracellular parasite) and at least one partner must be much more
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mobile and seek the other across tissues.
Moreover, as soon as szygy was lost for that reason, male gametes suddenly were for the
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first time exposed to the selective force favouring numerous tiny sperm, and thus evolved more divisions than the female; conversely this tipping of the balance imposed a new force on the female
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to reduce the number of divisions eventually to zero to maximise egg size (Bulmer and Parker 2002; Parker 1978). Intracellularity of Cryptosporidium exposed it to the same selective forces as in most coccidiomorphs, so it independently evolved female gamonts that do not divide before fertilization. As apicomonads probably have equal biciliate gametes little different from vegetative cells, both the gregarine and the coocidia-like, gametogony patterns are derived compared with the ancestral condition. But both almost certainly diverged independently polyphyletically from a less sexually dimorphic ancestral sporozoan, which was probably an intracellular parasite like coccidia rather than a purely extracellular one. An intracellular ancestry is more likely because in both orthogregarines and Paragregarea development starts intracellularly and because, as I explain in a separate paper, full penetration into host cells prior to intracellular development provides a marked contrast to the extracellular nature of apicomonads and thus a novel selective force that favoured the
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39 origin of symmetrical conoids, two preconoidal rings, and microtubule nucleating polar ring from the asymmetric pseudoconoid and pellicles of apicomonads. Thus the contrasting patterns of sexual dimorphism in Sporozoa are also consistent with gregarine polyphyly, as they are simply explicable
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as multiple, mutationally and developmentally easy, responses to similar selective forces.
Derived gliding motility
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It is notable that actin-myosin based gliding motility of most orthogregarines is similar to the coccidiomorph cell entry machinery (Valigurová et al. 2013); coccidiomorphs use actin-based
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gliding past the host/parasite cell junction ring to enter host cells. I suggest that this gliding
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machinery evolved in a common ancestor of Gregarinomorphea and Coccidiomorphea, possibly but not necessarily after it diverged from Paragregarea. The absence of translational gliding and
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presence of pendular or threshing movements in archigregaines and Veloxidium is a marked difference consistent with an early divergence of Paragregarea from a putative
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gregarinomorph/coccidiomorph clade, as suggested by the CAT tree of Fig. 2. Also suggestive of such a clade is the fact that their conoids have two intraconoid microtubules, never more. Though
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evidence concerning paragregarine intraconoid microtubules is scrappy and inconclusive, Lankesteria has at least one (Garnham et al. 1971 Fig. 3; possibly two if the two densities between the two right hand rhoptries on the transversely sectioned conoid in Fig. 1 are microtubules). One image for the archigregarine Selenidium hollandei (Schrével 1971 Fig. 24) suggests that there may be three, like the anterior rostral root of Chilovora perforans (Brugerolle and Mignot 1979). If that difference were to be confirmed it could be explained as early divergence from an apicomonad ciliary root incorporated into the conoid lumen when it closed (Cavalier-Smith unpublished). Gliding motility probably evolved independently in orthogregarines and Urosporoidea. Though orthogregarines and Paragregarea most likely evolved independently from the earliest ancestral sporozoans, non-gliding Squirmidea are probably not even Sporozoa. Squirmids probably lack not only conoids (which can be secondarily lost in both coccidiomorphs and
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40 gregarinomorphs) but even a symmetrical apical complex found even in all secondarily aconoidal sporozoans – even Cryptosporidium which has largely lost pellicular microtubules retains a radially symmetrical and extra thick polar ring; Cryptosporidium also retained the two gregarinomorph intraconoidal microtubules with an historical central asymmetry, as in all sporozoan apical
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complexes. Even Plasmodium, where some species have no conoid or intraconoid microtubules (despite their radially symmetric apical protrusion being called the conoid region or sometimes
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carelessly even conoid (Ladda et al. 2001)), retains two preconoidal rings (Hepler et al. 1966) - at least one species kept an asymmetrical dense fibre connecting its degenerate apical complex to the
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centriole, which is likely to have descended from an apicomonad anterior root, an idea reinforced
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by the discovery of the highly conserved corticate anterior striated root protein SF-assemblin in the
Conclusions
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probably homologous centriolar/conoidal connector of Toxoplasma (Francia et al. 2012).
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1. The improved CAT-GTR-GAMMA phylogenies provide a firmer myzozoan tree onto which cellular and host-interaction differences across Sporozoa map in a much more
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evolutionarily comprehensible way than before. 2. Gregarines are polyphyletic. Orthogregarines are most closely related to Cryptosporidium and then rhytidocystids (here placed in three separate subclasses of Gregarinomorphea) than to Paragregarinea (Archigregarinida, and new orders Velocida and Stenophorida). Platyproteum and Filipodium are probably not gregarines (i.e. neither orthogregarines nor paragregarines) nor Sporozoa and placed in a new class Squirmidea. 3. The three former gregarine groups represent similar convergent adaptation towards terminally extracellular giant trophic cells within intestines, but show significant differences in detail reflecting their probably independent origins from broadly similar myzocytotic ancestors. Environmental DNA BOLA566 is probably from a stenophorid paragregarine.
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41 4. Many improvements to sporozoan classification have been made, especially at highest ranks and within orthogregarines. Most notably, Eugregarinida have been split into four orders (orthogregarine Vermigregarida and Arthrogregarida; paragregarine Velocida and Stenophorida) and Neogregarinida merged into Arthrogregarinida.
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5. I explained the key selective forces that led to the differentiation of sporozoan cell structure and life-histories in an often convergent manner that was previously taxonomically
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confusing.
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Appendix A. Supplementary data
Supplementary data associated with this article can be found, in the online version, at
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http://dx.doi.org/...
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Perkins, F.O., Barta, J.R., Clopton, R.E., Pierce, M.A., Upton, S.J., 2002 dated 2000. Phylum
M
Apicomplexa Levine 1970, in: Lee, J.J., Leedale, G., Bradbury, P. (Eds.), An Illustrated Guide to the Protozoa, 2nd Ed.Vol. 1. Society of Protozoologists, Lawrence, Kansas, pp. 190-
d
369.
Porchet-Henneré, E., 1967. Preliminary observations on the fine structure of the male gamete of
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Coccidia Coelotropha durchoni (coccidiomorphous sporozoa). Compt. Rend. Hebd. Sean. Acad. Sci. D. 264, 2130-2133.
Porchet-Henneré, E., 1971. La fécondation et la sporogenèse chez la coccidie Coelotropha durchoni. Étude en microscopie photonique et électronique. Z. Parasitenk. 37, 94-125. Pritchard, A., 1861. A History of the Infusoria, Including the Desmidiaceae and Diatomaceae, British and Foreign. 4th Ed. Whittaker, London. Richards, T.A., Vepritskiy, A.A., Gouliamova, D.E., Nierzwicki-Bauer, S.A., 2005. The molecular diversity of freshwater picoeukaryotes from an oligotrophic lake reveals diverse, distinctive and globally dispersed lineages. Environ. Microbiol. 7, 1413-1425. Roure, B., Baurain, D., Philippe, H., 2013. Impact of missing data on phylogenies inferred from empirical phylogenomic data sets. Mol. Biol. Evol. 30, 197-214.
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50 Rueckert, S., Chantangsi, C., Leander, B.S., 2010. Molecular systematics of marine gregarines (Apicomplexa) from North-eastern Pacific polychaetes and nemerteans, with descriptions of three novel species: Lecudina phyllochaetopteri sp. nov., Difficilina tubulani sp. nov. and Difficilina paranemertis sp. nov. Int. J. Syst. Evol. Microbiol. 60, 2681-2690.
ip t
Rueckert, S., Leander, B.S., 2009. Molecular phylogeny and surface morphology of marine archigregarines (Apicomplexa), Selenidium spp., Filipodium phascolosomae n. sp., and
cr
Platyproteum n. g. and comb. from North-Eastern Pacific peanut worms (Sipuncula). J. Eukaryot. Microbiol. 56, 428-439.
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Rueckert, S., Leander, B.S., 2010. Description of Trichotokara nothriae n. gen. et sp.
Onuphidae). J. Invert. Pathol. 104, 172-179.
an
(Apicomplexa, Lecudinidae)--an intestinal gregarine of Nothria conchylega (Polychaeta,
M
Rueckert, S., Simdyanov, T.G., Aleoshin, V.V., Leander, B.S., 2011a. Identification of a divergent environmental DNA sequence clade using the phylogeny of gregarine parasites
d
(Apicomplexa) from crustacean hosts. PLoS One 6, e18163. Rueckert, S., Villette, P.M., Leander, B.S., 2011b. Species boundaries in gregarine apicomplexan
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parasites: a case study-comparison of morphometric and molecular variability in Lecudina cf. tuzetae (Eugregarinorida, Lecudinidae). J. Eukaryot. Microbiol. 58, 275-283. Rueckert, S., Wakeman, K.C., Leander, B.S., 2013. Discovery of a diverse clade of gregarine apicomplexans (Apicomplexa: Eugregarinorida) from Pacific eunicid and onuphid polychaetes, including descriptions of Paralecudina n. gen., Trichotokara japonica n. sp., and T. eunicae n. sp. J. Eukaryot. Microbiol. 60, 121-136. Saffo, M.B., 1981. The enigmatic protist Nephromyces. BioSystems 14, 487-490. Saffo, M.B., McCoy, A.M., Rieken, C., Slamovits, C.H., 2010. Nephromyces, a beneficial apicomplexan symbiont in marine animals. Proc. Natl Acad. Sci. USA 107, 16190-16195. Saville Kent, W., 1880-1882. A manual of the Infusoria. Bogue, London.
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51 Scholtyseck, E., 1965. The microgamete development of Eimeria perforans. Z. Zellforsch. Mikrosk. Anat. 66, 625-642. Scholtyseck, E., Mehlhorn, H., Hammond, D.M., 1972. Electron microscope studies of microgametogenesis in Coccidia and related groups. Zeitschr. Parasitenk. 38, 95-131.
ip t
Schrével, J., 1971. Observations biologiques et ultrastructurales sur les Selenidiidae at leurs conséquences sur la systématique des Grégarinomorphes. J. Protozool. 18, 448-470.
cr
Schrevel, J., Besse, C., 1975. Un type flagellaire functionnel de base 6 + 0. J. Cell Biol. 66, 492507.
us
Schrével, J., Desportes, I., 2013. Chapter 1 Introduction: Gregarines among Apicomplexa, Treatise
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on Zoology - Anatomy, Taxonomy, Biology: The Gregarines Vol. 1, pp. 7-. Simpson, A.G.B., Patterson, D.J., 1996. Ultrastructure and identification of the predatory flagellate
M
Colpodella pugnax Cienkowski (Apicomplexa) with a description of Colpodella turpis n. sp. and a review of the genus. Syst. Parasitol. 33, 187-198.
d
Speer, C.A., Danforth, H.D., 1976. Fine-structural aspects of microgametogenesis of Eimeria magna in rabbits and in kidney cell cultures. J. Protozool. 23, 109-115.
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Stoeck, T., Epstein, S., 2003. Novel eukaryotic lineages inferred from small-subunit rRNA analyses of oxygen-depleted marine environments. Appl. Environ. Microbiol. 69, 2657-2663. Templeton, T.J., Enomoto, S., Chen, W.J., Huang, C.G., Lancto, C.A., Abrahamsen, M.S., Zhu, G., 2010. A genome-sequence survey for Ascogregarina taiwanensis supports evolutionary affiliation but metabolic diversity between a gregarine and Cryptosporidium. Mol. Biol. Evol. 27, 235-248.
Valigurová, A., Hofmannová, L., Koudela, B., Vávra, J., 2007. An ultrastructural comparison of the attachment sites between Gregarina steini and Cryptosporidium muris. J. Eukaryot. Microbiol. 54, 495-510. Valigurová, A., Vaškovicová, N., Musilová, N., Schrével, J., 2013. The enigma of eugregarine epicytic folds: where gliding motility originates? Front. Zool. 10, 57.
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52 Vetterling, J.M., Pacheco, N.D., Fayer, R., 1973. Fine structure of gametogony and oocyst formation in Sarcocystis sp. in cell culture. J. Protozool. 20, 613-621. Vivier, E., 1982. Reflexions et suggestions à propos de la systématique des Sporozoaires: création d'une classe des Hematozoa. Protistologica 18, 449-453.
ip t
Vivier, E., Desportes, I., 1990. Phylum Apicomplexa, in: Margulis, L., Corliss, J.O., Melkonian, M., Chapman, D.J. (Eds.), Handbook of Protoctista. Jones & Bartlett, Boston, pp. 549-573.
cr
Wakeman, K.C., Leander, B.S., 2013a. Identity of environmental DNA sequences using
from capitellid polychaetes. Mar. Biodiv. 43, 133-147.
us
descriptions of four novel marine gregarine parasites, Polyplicarium n. gen. (Apicomplexa),
an
Wakeman, K.C., Leander, B.S., 2013b. Molecular phylogeny of marine gregarine parasites (Apicomplexa) from tube-forming polychaetes (Sabellariidae, Cirratulidae and Serpulidae),
M
including descriptions of two new species of Selenidium. J. Eukaryot. Microbiol. 60, 514-525. Yabuki, A., Toyofuku, T., Takishita, K., 2014. Lateral transfer of eukaryotic ribosomal RNA
doi:10.1038/ismej.2013.252
d
genes: an emerging concern for molecular ecology of microbial eukaryotes. ISME J. 1-4
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1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65
Page 52 of 60
53 Figure legends
Fig. 1. GTR gamma maximum likelihood tree for 18S rDNA of 652 eukaryotes using 1540 nucleotide positions, showing gregarine branches collapsed on Fig. 1. Numbers on branches
ip t
show bootstrap support (400 resamplings) for almost all bipartitions (those ≥ 85% in bold); black blobs indicate 100% support. To compress the tree to a single page some speciose clades
cr
are collapsed, the number of taxa in each being indicated to the right of the clade name.
Percolozoa are shown non-collapsed and Dinozoa and Euglenozoa less collapsed in Fig 2.
us
Fig. 2. PhyloBayes CAT-GTR-GAMMA tree for 18S rDNA of 137 Myzozoa only using 1614
an
nucleotide positions, excluding long-branch gregarines. Support values for this consensus tree from two well converged chains (maxdiff 0.128) are posterior probabilities; black blobs indicate
M
1.0 PP. Exclusion of longer-branch gregarines coupled with the evolutionarily more realistic site-heterogeneous CAT model makes it even clearer than in Fig. 1 that Squirmidea formerly
d
treated as gregarines do not branch within Sporozoa but with Dinozoa and that archigregarines are less closely related to orthogregarines than is Cryptosporidium. The arrow marks the
Ac ce pt e
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65
putative time of loss of the apicoplast in a common ancestor of Cryptosporidium and orthogregarines.
Fig. 3. PhyloBayes CAT-GTR-GAMMA tree for 18S rDNA of 196 alveolates only using 1577 well-aligned nucleotide positions, including long-branch gregarines. Support values are posterior probabilities; black blobs indicate 1.0 PP. Sum of two well-converged chains (maxdiff 0.101316). Though an ML tree was also run for this alignment, for clarity its support values are not shown, but some key ones are mentioned in the text. Fig. 4. PhyloBayes CAT-GTR-GAMMA tree for 18S rDNA of 276 alveolates only using 1577 well-aligned nucleotide positions. Support values are posterior probabilities; black blobs indicate 1.0 PP. This tree is for one of two chains that did not converge (maxdiff 0.6801); the other chain differed in only two respects as discussed in the text.
Page 53 of 60
54 Table 1. Revised classification of parvphylum Sporozoa Leuckart, 1879 and its 4 classes and 13 orders Class 1. Paragregarea cl. n. (mainly marine; annelid, nemertean, myriapod, mollusc, deuterostome hosts) (three orders, two new) Order 1. Archigregarinida* Grassé, 1953 (largely marine and aseptate; longitudinal ridges; contractile, non-gliding)
Levine, 1971) (Selenidioides, Meroselenidium, Merogregarina)
ip t
Family 1. Merogregarinidae Porter, 1908 (invalid junior synonym Selenidioididae
Family 3. Exoschizonidae Levine, 1971 (Exoschizon)
us
Order 2. Velocida ord. n. (aseptate; motility varied)
cr
Family 2. Selenidiidae* Brasil, 1907 (e.g. Selenidium*, Selenocystis)
Superfamily 1. Veloxidioidea superfam. n. (contractile, non-gliding; no longitudinal folds)
an
Family Veloxidiidae fam. n. (Veloxidium)
Superfamily 2. Urosporoidea Léger, 1892 superfam. n. (formerly eugregarines;
M
gliding, longitudinal folds)
Family 1. Lecudinidae Kamm, 1922 (e.g. Lecudina, Lankesteria, Difficilina) Family 2. Urosporidae Léger, 1892 (e.g. Urospora, Pterospora, Lithocystis)
d
Order 3. Stenophorida ord. n. (millipede hosts; ‘septate’) Family Stenophoridae Léger and Dubosq, 1904 (e.g. Stenophora)
Ac ce pt e
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65
Class 2. Gregarinomorphea Grassé, 1953 em. (four orders; three new) Subclass 1. Histogregaria subcl. n.
Order Histogregarida nom. n. pro Agamococcidiorida: Levine, 1979 = Agamococcidiida Levine, 1980.
Family 1. Rhytidocystidae Levine, 1979 Family 2. Gemmocystidae Upton and Peters, 1986
Subclass 2. Cryptogregaria subcl. n. Order Cryptogregarida ord. n.
Family Cryptosporidiidae Léger, 1911 Subclass 3. Orthogregarinia subcl. n. Order 1. Vermigregarida ord. n. (mainly polychaete gregarines) Family 1. Trichotokaridae fam. n. (Trichotokara, Paralecudina) Family 2. Polyplicariidae fam. n. (Polyplicarium) Order 2. Arthrogregarida ord. n. (mainly arthropod hosts; often ‘septate’) Page 54 of 60
55 Suborder 1. Porosporina subord. n. (crustacean, seldom mollusc, hosts) Superfamily Porosporoidea Labbé, 1899 (not Porosporicae Chakravarty, 1960; syn. Cephaloidophoroidea Simdyanov & Aleoshin in Rueckert et al., 2011) Family 1. Porosporidae Labbé, 1899 (Porospora, Nematopsis, Pachysporospora, Thiriotia) Family 2. Cephaloidophoridae Kamm, 1922 (e.g. Cephaloidophora)
ip t
Family 3. Uradiophoridae Grassé, 1953 (e.g. Heliophora, Uradiophora) Family 4. Ganymedidae Huxley, 1910 (Ganymedes)
cr
Suborder 2. Terragregarina subord. n. (terrestrial arthropod or oligochaete hosts; cilia without central pair microtubules)
us
Superfamily 1. Stylocephaloidea Léger, 1892 (rank given by Clopton, 2009) Family 1. Stylocephalidae Léger, 1892 (e.g. Stylocephalus, Xiphocephalus, Colepismatophila) has one cilium szygy frontal, late; beetle, Diplura, Thysanura hosts.
an
Superfamily 2. Actinocephaloidea Léger, 1892 superfam.. n. (szygy frontal, late; insect, arachnid, myriapod hosts)
M
Family 1. Actinocephalidae* Léger, 1892 (e.g. Geneiorhynchus, Prismatospora, Hoplorhynchus, Pyxinia)
Paraschneideria)
d
Family 2. Monocystidae Bütschli, 1882 (Monocystis, Ascogregarina,
Family 3. Ophryocystidae Léger and Dubosq, 1908 (Ophryocystis, Apicystis)
Ac ce pt e
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65
Family 4. Syncystidae Schneider, 1886 (Syncystis, Steinina) Family 5. Lipotrophidae Grassé, 1953 (e.g. Mattesia) Family 6. Sphaerocystidae Chakravarty, 1960 (Sphaerinina)
Superfamily 3. Gregarinoidea Labbé, 1899 (not Chakravarty, 1960) szygy early Family 1. Gregarinidae* Labbé, 1899 (e.g. Gregarina, Leidyana, Amoebogregarina) Family 2. Blabericolidae Clopton, 2009 (e.g. Blabericola, Protomagalhaensia)
Terragregarina incertae sedis***: Dactylophoridae Léger, 1892 (centipede hosts)
Class 3. Coccidiomorphea Doflein, 1901 (five orders) em. Subclass 1. Coccidia Leuckart, 1879 (3 orders: Adeleida Léger, 1911 (with szygy) including families Adeleidae and Aggregatidae; Eimeriida Léger, 1911 (Eimeriidae, Calyptosporidae, Sarcocystidae, Elleipsisomatidae); Ixorheida Levine, 1984 (no gamogony). Subclass 2. Hematozoa Vivier, 1982 (=Aconoidasida Mehlhorn et al., 1980**) Superorder Haemosporidia Danilewsky, 1885; order Haemosporida Danielwsky, 1885
Page 55 of 60
56 Superorder Aconoidia superord. n. orders Nephromycida Cavalier-Smith, 1993 (ascidian symbionts: Nephromyces, Cardiosporidium); Piroplasmida Wenyon, 1926) Subclass 3. Coelotrophia subcl. n. Order Coelotrophiida Vivier, 1982 (=Protococcidiida Kheisin, 1956) (5 families, e.g. Coelotropha)
Sporozoa incertae sedis: Order Blastogregarinida Chatton and Villeneuve, 1936 (no folds)
ip t
Family Siedleckiidae Chatton and Villeneuve, 1936 (Siedleckia) For all taxa I use the uniform zoological endings recommended by Pearse (1936) for phyla (-
cr
a) classes (-ea), subclasses (-ia) orders (-ida), and suborders (-ina), which have become
standard for Protozoa and phagotrophic or parasitic non-algal chromists, not the longer
us
replacement forms suggested by Levine (1959) for all zoological nomenclature but not adopted by anyone else except anomalously for Sporozoa. I suggest that Levine’s idiosyncratic respellings of older names (e.g. Piroplasmorida for Piroplasmida) be now abandoned for
an
Sporozoa also. Families listed only for gregarines. * paraphyletic short 2-microtubule conoid.
M
** name unsuitable as conoid present on one haemosporidan ookinete and apparently has
d
***Many other mainly insect-host, probably terragregarine, families are not listed because no sequences are available and it is therefore unclear to which superfamily they belong.
Ac ce pt e
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65
Page 56 of 60
Figure 1
6
87
10
20 Apicomonadea 8 93 Coccidea/Hematozoa 7 4 87 Cryptosporidium Rhytidocystis cyamus GQ149767
17 33
Sporozoa
9
31
49
22 23 42
C3_E012 AY046842 C2_E026 AY046816 BOLA212 AF372767 BOLA458 AF372771 environmental clone BOLA327 AF372770 Paralecudina polymorpha AY196706
40
31 2
26
3
85
BAQA65 AF372825
Blabericola migrator FJ459754
97 Blabericola haasi FJ459753 Protomagalhaensia granulosae FJ459757 Protomagalhaensia wolfi FJ459758 Blabericolidae 99 99 Gregarina basiconstrictonea FJ459740
Gregarinoidea
1
85 37
4
24
67
Veloxidium leptosynaptae JN857966
Filipodium phascolosomae FJ832163 Platyproteum vivax AY196708
63 Dinozoa 36
Chilodonella uncinata
2
0
Diphylleia rotans AF420478 triciliatum ContigKST Diphylleida 16 RigifilaCollodictyon ramosa AB686266 Rigifilida Micronuclearia podoventralis anoxic marine sediment DNA CCA32 AY179990 NOT dinoflagellate Telonema subtilis RCC3587 Telonema antarcticum AJ564773 Telonemea 24 54 Ministeria vibrans AF271998 63 29 Capsaspora owczarzaki 59 Monosiga ovata AF271999 Monosiga brevicollis 80 28 Salpingoeca infusionum Amoebidium parasiticum Y19155 29 Ichthyophonus hoferi 43 Corallochytrium limacisporum Fonticula alba FJ816018 72 36 RT5iin16 AY082991 Cristidiscoidea RT5iin14 AY082985 42 Nuclearia simplex 4 Rozellidea 52 36 Aphelidida Rozellida 4 35 47 60 25 FUNGI 13 85 Manchomonas bermudensis Apusomonas proboscidea 40 microbial mat cold seep DNA RM1-SGM31 AB505488 62 30 marine marine mIirobial mat cold seep DNA RM2-SGM40 AB505548 marine microbial mat cold seep DNA RM2-SGM42 AB505550 33 Thecamonas trahens EU542595 12 AT4-11 AF530526 Thecamonas sp. CCAP 9795 contaminant 28 marine microbial mat cold seep DNA RM2-SGM44 AB505552 12 25 90 marine microbial mat cold seep DNA RM1-SGM33 AB505490 95 marine microbial mat cold seep DNA RM1-SGM37 AB505494 marine microbial mat cold seep DNA DSGM-75 AB275075 marine microbial mat cold seep DNA RM1-SGM35 AB505492 marine microbial mat cold seep DNA RM2-SGM45 AB505553 lake sediment NAMAKO-2 DNA AB252742 90 RM2-SGM71 AB505579 RM1-SGM28 AB505485 95
Varisulca Sulcozoa
kingdom ANIMALIA 7
Ac ce pt e 0
10
2
2
21
3
95
0 7 12
53
28
77
34
38
91
79
26
99
77 49
D3P05C09 EF100280 BOLA366 AF372746
70
D4P07D12 EF100407 SA2_2C8 EF527166 BOLA187 AF372745
Subulatomonas tetraspora HQ342676
RM1-SGM29 AB505486 NAMAKO-1 AB252741 RM2-SGM39 AB505547
7
Choanozoa podiates Apusomonadida
Amoebozoa 12 clade
Breviatea
Breviata anathema WIM30 AM114802 Mantamonas plastica Mantamonadida Mantamonas sp.2 Africa 74 Planomonadida 5 Glaucophyta 3
91
6
19
86
picobiliphyte 3 Cryptista Viridiplantae 12
77
28
cr
Hacrobia p. p. (kingdom CHROMISTA)
submarine fumarole anoxic sediment DNA TAGIRI-28 AB1914 V. costatus contaminant EF483923 Microhelida Microheliella maris AF534711
d
3
Dinozoa
M
0
Urosporoidea Velocida
Ciliophora
AF300282
Heliozoa 7
96
'core paragregarines'
53 Heterokonta + Rhizaria 48
Haptophyta 5 Palpitomonas bilix AB508339 Palpitia
16
Veloxidiidae Squirmidea
Mesodinium/Myrionecta 12
Blepharisma americanum BJA16SLRNA Loxodes striatus AM946031 Remanella sp. R101 AM409181 Trachelocerca ditis GQ167153 Tracheloraphis sp. TCIRGSSA AF149979 Paramecium tetraurelia Halteria grandinella AF194410 Pseudourostyla cristata DQ019318
AT4-68 AF530543
19
us
chelyosomae EU670240 92Lankesteria 67 Lankesteria ascidiae JX187607 78 CCI7 AY179977 70 Lecudina tuzetae AF457128 70 53 Lankesteria abbotti DQ093796 99 Pterospora schizosoma DQ093793 Pterospora floridiensis DQ093794 Urosporidae Lithocystis sp. DQ093795 96 99 environmental clone DSGM-6 AB275006
94 26 33
Archigregarinida
an
20
(CHROMISTA)
Vermigregarida
environmental clone SS1E0145 EU050982 environmental clone NAMAKO-25 AB252765 Blabericola cubensis FJ459751
Selenidium idanthyrsae JN857967
37
Trichotokaridae
Selenidium cf. mesnili JN857968 Selenidiidae 86 AY196709 Selenidium serpulae DQ683562 Selenidium terebellae 27 Selenidium pisinnus FJ832162 Selenidium orientale FJ832161 Difficilina paranemertis FJ832159 Difficilina tubulani FJ832160 Lecudina longissima FJ832157 Lecudina phyllochaetopteri FJ832156 Lankesteria cystodytae EU670241 Lecudinidae
52 23
Trichotokara eunicae@JX426618 Trichotokara nothriae GU592817 Trichotokara japonica JX426617
Gregarina cuneata FJ459744 LEMD145 AF372805 LEMD003 AF372797 Gregarina cloptoni FJ459742 Gregarina niphandrodes FJ459747 55 gregarines insect gregarines Gregarina polymorpha FJ459748 81 Gregarina diabrotica FJ459745 80 Gregarina coronata FJ459743 Gregarinidae 19 34 Leidyana erratica FJ459752 Gregarina blattarum FJ459741 42 Gregarina chortiocetes NA18SRRB 99 Gregarina caledia GNA18SRRA Gregarina tropica FJ459749 Amoebogregarina nigra FJ459737 49 Gregarina kingi FJ459746 Stenophorida 64 Uncultured DNA AB275069 98 diplopod gregarinesStenophora robusta FJ459760 Thiriotia pugettiae HQ876006 Porosporidae Ganymedes themistos FJ976721 Ganymedidae Heliospora caprellae HQ876007 Uradiophoridae BOLA48 AF372821 Porosporoidea crustacean gregarines 97 DH148-EKD18 AF290084 85 CS_R003 AY046643 Cepahaloidophoridae 98 Cephaloidophora cf. communis HQ876008 63 Selenidium boccardellae JN857969
28
1
Harosa
81
22
4
Alveolata
short-branch gregarinesTerragregarina pro parte
DSGM_8 AB275008
3
Myzozoa
'core orthogregarines' BS 79%
Rhytidocystidae 49 Stylocepahaloidea + Actinocephaloidea49 parasite of Ammonia beccarii ABU07937 unidentified clade BOLA267 AF372774 clone CCI31 AY179976 clone CCA38 AY179975 Polyplicariidae
Tridacna hemolymph parasite AB000912
ip t
Apicomplexa
Sulcozoa
Glissodiscea
kingdom PLANTAE
clade
83 Rhodophyta 4
Anaeromonadea 10
Metamonada
Malawimonas jakobiformis Malawimonadea Malawimonas californiana Andalucia godoyi AND19 Andalucia godoyi AND28 Andalucia incarcerata Jakobea Guyamas hydrothermal marine sediment DNA CS_E022 AY046649 5 Histionina saltmarsh pool water DNA CCW8 AY180011 Tsukubamonas globosa AB576851
77 Euglenoida/ Postgaardea
Excavata Percolozoa
Eozoa 45
121 Kinetoplastea/Diplonemea Euglenozoa
0.4
Page 57 of 60
Figure 2
Tridacna haemolymph parasite AB000912 Rhytidocystis polygordiae DQ2739887 Rhytidocystis cyamus GQ149767
Cryptosporidium fragilis
Stylocephalus giganteus FJ459761 Colepismatophila watsonae FJ459738
0.38 0.75 0.56
Eimeria leucisci
DNA D5P09F10 EF100363 DNA D5P09C08 EF100358 Steinina (=Gregarina) ctenocephali GU320208 Monocystis agilis AF457127 Ophryocystis electroscirrha FJ865354
Actinocephaloidea
Goussia szekelyi Goussia koertingi Goussia pannonica Goussia janae Goussia desseri Goussia kessleri Goussia chalupski Goussia sp. Goussia balatonica Calyptospora spinosa Calyptospora serrasalmi Calyptospora funduli
ip t
Gregarinomorphea
Eimeriida
Frenkelia glareoli Hyaloklossia lieberkuhni Toxoplasma gondii Besnoitia oryctofelis Nephromyces sp. MO-3F HM469391 Nephromyces sp. MO-5F HM469379 Nephromyces sp. MO-4G HM469380 Nephromyces sp. MO-6H HM469378 Nephromyces sp. MO-4B HM469377 .75 Nephromyces sp. MO-6H HM469376 Nephromyces sp. MO-4F HM469375 Nephromyces sp. MR-1N HM469383 Nephromyces sp. MM-3N HM469382 Nephromyces sp. MC-2N HM469384 intrahemocytic apicomplexan from Halocynthia EF558768 Cardiosporidium cionae Babesia microti Babesia conradae Cytoauxzoon felis Theileria parva Babesia odocoilei Babesia gibsoni Babesia orientalis Babesia bovis Klossia helicina Hepatozoon catesbianae Hepatozoon canis Adelina grylli 0.99 Adelina bambarooniae Vitrella brassicaformis DQ174731 environmental DNA LT85_AS KC487674 environmental DNA LT85_K21 KC487845 environmental DNA LT35_H13 KC486567 environmental DNA LT35_ET3 KC486520 0.98 environmental DNA KC487652 environmental DNA GU067926 0.99 Colpodella (= Alphamonas) edax AY234843 BOLA914 AF372772 BOLA553 AF372785 .87
0.54 0.36 0.98
.93
0.98
Coccidia p. p.
Coccidiomorphea
Nephromycida Aconoidia
M
0.82
cr
0.99 Eimeria variabilis Eimeria anguillae 0.84 Goussia neglecta Eimeria arnyi 0.69 Eimeria myoxi Eimeria mitis 0.65 0.95 Cyclospora papionis Cyclospora colobi Cyclospora cayetanensis 0.93 0.99 Sarcocystis oviformis Sarcocystis sinensis Sarcocystis hirsuta 0.96 Sarcocystis gallotiae
d
0.99 0.95 0.98 0.54
Ac ce pt e
Adeleida
Piroplasmida
Sporozoa
Coccidia p. p.
Vitrella/Colpodella subclade
Apicomonadea
Chromerida Voromonas pontica AY078092
0.48
Chromera velia DQ174731
0.44 0.64
'Colpodella' sp. HEP GQ4111073 'Colpodella' sp. ATCC5059 AY142075 'Colpodella' tetrahymenae AF330214 BOLA176 AF372786 Psammosa pacifica JN873311 Psammosa atlantica JN873310 .54 Kofoidinium sp. FG256 GU355681 Syndinium sp. ex Corycaeus DO146406 .12 Hematodinium sp. MF-2000 AF286023 Amoebophrya sp. ex Prorocentrum micans AY208893 0.99 Amoebophrya sp. ex Ceratium tripos AY208892 Amoebophrya sp. ex Karlodinium 0.53 Amoebophrya sp. .57.58 0.69 Amoebophrya sp. AF069516 Heterocapsa triquetra GU594638 .47 Cochliodinium polykrikoides Karenia brevis Abedinium dasypus .21 Alexandrium tamarense AB088333 .41 Prorocentrum gracile .27 Polarella glacialis Haplozoon praxillellae .98 Haplozoon axiothellae
0.99
0.85
0.97
0.83 0.97 0.99
0.29
PROSOPE.E3_95m.109 EU793257 marine DNA 65 DQ916410 uncultured Dubosquella 3905 JF791022 Ichthyodinium chabelardi FJ440625 Kofoidinium cf. pavillardii GU355680 Spatulodinium pseudonoctiluca GU355685
Noctiluca scintillans AF022200
Dinozoa
Syndinea p. p. Dinoflagellata Oxyrrhida 0.99
Syndinea p. p.
Oxyrrhis marina AF280077 Oxyrrhis marina AB033717 Oxyrrhis marina AF482425
Dubosquellidae
Noctilucea Thalassomyces fagei AY340590
IN242 NA2_2D10 EF526760 Parvilucifera procentrum FJ424512 Parvilucifera sinerae EU502912 Parvilucifera infectans AF133909 Perkinsus atlanticus AF509333 marine hydrothermal vent DNA AT4-98 AF530536
0.47 .49 0.99 0.38 0.47 0.32
Apicomplexa
Voromonadida
Peridinea = Dinophyceae
.18 .12
Orthogregarinia
Paraschneideria metamorphosa DQ462456 Ascogregarina culicis Pseudomonocystis lepidota PNT18SRRNA Mattesia geminata AY334568 Apicystis bombi FN546182
0.86 0.94
0.65
0.45
Histogregaria
Cryptogregarida Cryptosporidium serpentis Cryptogregaria baileyi 0.970.95 Cryptosporidium Xiphocephalus ellisi FJ459762 Xiphocephalus triplogemmatus FJ459763 Stylocephaloidea
0.99
0.27
Rhytidocystidae
Cryptosporidium parvum
0.91
0.99
Stenophorida
Stenophora robusta FJ459760
us
0.92
Paragregarea
Archigregarinida
Selenidium terebellae AY196709 Selenidium pisinnus FJ832162 Selenidium orientale FJ832161 BOLA566 AF372760
an
0.51 0.89 0.69 0.92
0.94 0.3
Ellobiopsida Ellobiopsis chattonii FJ593706
Perkinsea Squirmidea
Perkinsozoa Platyproteum vivax AY196708 Filipodium phascolosomae
Page 58 of 60
Figure 3 Colpodella (=Alphamonas) edax AY234843
Vitrella/Colpodella clade Voromonadidida Voromonas pontica AY078092 Chromera/Voromonas clade Chromerida Chromera velia DQ174731 Platyproteum vivax AY196708
environmental DNA GU0679
'Colpodella'-like 3
Squirmidea 0.99 Rhytidocystidae Histogregaria 0.98
Filipodium phascolosomae FJ832163
Cryptosporidium 4
Cryptosporidiidae
Tridacna haemolymph parasite AB000912 Rhytidocystis polygordiae DQ2739887
Rhytidocystis cyamus GQ149767
0.99
0.5
clone CCI31 AY179976 clone CCA38 AY179975
0.48
0.23
Polyplicariidae
parasite of Ammonia beccarii U07937
BOLA267 AF372774
0.58
0.62
0.67 0.68
0.51 0.19
0.34
0.99 0.86
0.91
0.99
environmental clone BOLA327 AF372770
Stylocephalidae ES
Blabericola haasi FJ459753
Blabericolidae
Amoebogregarina nigra FJ459737 Leidyana erratica FJ459752
0.71
Gregarinidae
Actinocephalidae ES
Pyxinia crystalligera FJ459759 cooling tower water DNA CTW-5c JF774861
stained glass window DNA clone 1195 DQ451601
0.99
Hoplorhynchus acanthatholius FJ459750
Prismatospora evansi FJ459756 Monocystidae Monocystis agilis AF457127 saltmarsh sediment DNA alveolate clone CCA5 AY179988
strain ATCC 50646 GQ377652
EA
Actinocephalidae ES
us
0.97
Gregarinoidea
Gregarina tropica FJ459749
DNA D5P09F10 EF100363 DNA D5P09C08 EF100358
0.29 0.47 0.4
Stylocephaloidea
Protomagalhaensia wolfi FJ459758 Protomagalhaensia granulosae FJ459757 Blabericola migrator FJ459754
Geneiorhynchus manifestus J459739
0.95
0.45
Porosporoidea 8 crustacean gregarines
Blabericola cubensis FJ459751 Gregarina blattarum FJ459741 0.99 Gregarina chortiocetes NA18SRRB Gregarina caledia GNA18SRRA Gregarina kingi FJ459746 0.85 Gregarina diabrotica FJ459745 0.88 Gregarina coronata FJ459743 Gregarina polymorpha FJ459748 Gregarina niphandrodes FJ459747 0.32 0.4 Gregarina cloptoni FJ459742 LEMD145 AF372805 LEMD003 AF372797 Gregarina cuneata FJ459744 0.99 0.86 0.89 0.43 Gregarina basiconstrictonea FJ459740
0.33
13 soil DNA clones (Lesaulnier et al. 2008) wrongly annotated as Eimeriidae + Lake AF37277 & EU910605 or Cryptosporidiidae Syncystidae N Syncystis mirabilis DQ176427 Steinina ctenocephali GU320208 Actinocephalidae ES 0.98 0.99 Psychodiella chagasi FJ865354 0.85 Ophryocystis electroscirrha FJ865354 Ophryocystidae N 'Lecudinidae' Paraschneideria metamorphosa DQ462456 Sphaerocystidae EA 0.91 0.98 Ascogregarina 4 0.99 0.83 Pseudomonocystis lepidota PNT18SRRNA E Mattesia geminata AY334568 0.8 Lipotrophidae N 0.99 Apicystis bombi FN546182 Stenophora robusta FJ459760 BOLA566 AF372760 0.45 0.99 Selenidium idanthyrsae JN857967 0.23 0.99 Selenidium serpulae DQ683562
0.99
an
gregarine env. DNACHI_S2_24 AY821921
0.14
0.84 Selenidium cf. mesnili JN857968 Selenidium boccardellae JN857969 Selenidium pisinnus FJ832162 Selenidium orientale FJ832161 Selenidium terebellae AY196709 Veloxidium leptosynaptae JN857966 0.67 environmental clone DSGM-6 AB275006 0.59 Difficilina tubulani FJ832160 Difficilina paranemertis FJ832159 Pterospora schizosoma DQ093793 0.88 Pterospora floridiensis DQ093794 Lithocystis sp. DQ093795 Lecudina longissima FJ832157 0.98 Lecudina phyllochaetopteri FJ832156 0.27 Lankesteria cystodytae EU670241 0.78 Lankesteria chelyosomae EU670240 0.99 Lankesteria ascidiae JX187607 Lankesteria abbotti DQ093796 0.99 Coccidiomorphea 7 Lecudina tuzetae AF457128 0.99 0.86 CCI7 AY179977
Selenidiidae
0.77
Kofoidinium sp. FG256 GU355681
AF022200 0.7 Noctiluca scintillans Thalassomyces fagei AY340590 0.45 0.24 Ellobiopsis chattonii FJ593706 Ellobiopsida 0.99 Oxyrrhida 0.99 Psammosa pacifica JN873311 Oxyrrhis marina AF482425 Psammosa atlantica JN873310 0.36 0.11 Amoebophryidae 5 0.84 Dubosquellidae 4 0.96 0.48 Syndinea Syndinium sp. ex Corycaeus DO146406 Hematodinium sp. MF-2000 AF286023 0.63 0.82 0.97 Peridinea 9
0.96 0.6 0.99
Stenophorida
Archigregarinida
Urosporidae
Lecudinidae
Urosporoidea
Velocida Paragregarea
Dinozoa
Syndiniidae
0.99 Mesodinium pupula N412739
Mesodinium pulex QD-I DQ845294 Mesodinium pulex DQ411865 Mesodinium pulex JN412740
CCW75 AY180032 0.99 Myrionecta rubra AB364286 Mesodinium rubrum JN412736 rubrum JN412738 0.87 Mesodinium Ciliophora Myrionecta rubra AY587129 Mesodinium rubrum CCMP2563 JN412738 0.89 AF290065 AY180041 0.88 DH145-EKD11 Mesodinium chamaeleon JN084213 0.99 CCW100 AY180041
Karyorelictea 4 other Intramacronucleata 4
Blepharisma americanum BJA16SLRNA
0.98
Terragregarina
Noctilucea
Ac ce pt e
0.99 Spatulodinium pseudonoctiluca GU355685
Actinocephaloidea 0.97
Lecudinidae
d
Perkinsea 7
BAQA65 AF372825
A r t h r o g r e g a r i d a
Veloxidiidae Veloxidioidea
M
0.23
Kofoidinium cf. pavillardii GU355680
Trichotokara nothriae GU592817 Trichotokara eunicae@JX426618 Trichotokara japonica JX426617
Trichotokaridae Vermigregarida
BOLA458_AF372771 BOLA212 AF372767
Stylocephalus giganteus FJ459761 Colepismatophila watsonae FJ459738 0.99
0.75 0.77
0.46
0.35
environmental cloneNAMAKO-25 AB252765 C3_E012 AY046842 C2_E026 AY046816 Paralecudina polymorpha AY196706
0.6
Xiphocephalus triplogemmatus FJ459763 Xiphocephalus ellisi FJ459762
0.51
0.43
unidentified clade
environmental clone SS1E0145 EU050982
0.99
0.5 0.68
Orthogregarinia
DSGM_8 AB275008
0.76
0.1
0.44
Apicomonadea
Vitrella brassicaformis DQ174731
ip t
0.99
cr
0.51 0.75 0.3
0.4
Page 59 of 60
Figure 4 0.46 Voromonas pontica AY078092 Colpodella-like 3 0.99 Voromonadida 0.76 Chromera velia DQ174731 Chromerida Apicomonadea 0.21 Vitrella brassicaformis DQ174731 0.57 environmental DNA GU0679 Vitrella/Colpodella clade (=Alphamonas) edax AY234843 0.21 Colpodella 0.99 Platyproteum vivax AY196708 Filipodium phascolosomae FJ832163 0.77 0.95 Cryptosporidium 4 Cryptosporidia Squirmidea Gregarinomorphea Rhytidocystidae Agamococcidia 0.15 0.99 Polyplicariidae + unidentified subclade 0.57 DSGM_8 AB275008 Porosporoidea 8 crustacean gregarines 0.1 Paralecudina clade .62 0.89 0.65 Stylocephaloidea 4 0.1
Apicomplexa
Geneiorhynchus manifestus J459739
.93 0.41 DNA D5P09F10 EF100363
GQ377652 0.99 stained glass window DNA clone 1195 DQ451601 cooling tower water DNA CTW-5c JF774861
0.97 0.83 0.27 0.5
other Actinocephaloidea
Selenidium idanthyrsae JN857967 Selenidium serpulae DQ683562 Selenidium cf. mesnili JN857968 Selenidium boccardellae JN857969
0.76
BOLA566 AF372760
0.35
.83
Velocida
cr Eimeriidae
Eimeriida
0.7 Calyptospora spinosa FJ904637 Calyptospora serrasalmi FJ904641 0.79 Calyptospora funduli GU479670 Sarcocystis 6 0.65
Frenkelia glareoli AF009245
Sarcocystidae
Nephromyces sp. MO-6H HM469378
.99 Nephromyces sp. MO-4B HM469377 .99.99.99 Nephromyces sp. MO-6C HM469376
Halocynthia intrahemocytic apicomplexan EF558768
d
0.62 .99 Cardiosporidium cionae EU052685
.94
Coccidiomorphea
Nephromycida
Nephromyces sp. MO-4F HM469375 Nephromyces sp. MR-1N HM469383 Nephromyces sp. MM-3N HM469382 .96 Nephromyces sp. MC2-2N HM469384
.99 .61
0.73
Gregarinoidea
(Gregarinomorphea)
lieberkuehni AF298623 .76 Hyaloklossia Toxoplasma gondii X75453 .65 Besnoitia oryctofelisi GU479632 .96 Nephromyces sp. MO-3F HM469381 .99 Nephromyces sp. MO-5H HM469379 .98 .99 Nephromyces sp. MO-4G HM469380
.99
Paragregarea
Stenophorida
Stenophora robusta FJ459760
Goussia szekelyi GU479656 Goussia koertingi GU479647 Goussia pannonica GU479642 Goussia janae GU479644 Eimeria leucisci GU479649 Goussia desseri GU479665 Goussia kessleri GU479645 0.9 Goussia chalupskyi GU479653 Goussia sp. BMR-2011d GU479643 0.89 Goussia balatonica GU479650 Eimeria variabilis GU479674 Eimeria anguillae GU479633 Goussia FJ009242 0.99 Eimeria neglecta arnyi AY613853 0.88 Eimeria myoxi JF304148 Cyclospora papionis AF111187 Cyclospora colobi AF111186 0.91 Eimeria mitis EMU40262 Cyclospora cayetanensis AF111183
0.07
Archigregarinida
an
0.41
0.73
0.4
Selenidium pisinnus FJ832162
orientale FJ832161 0.6 SelenidiumSelenidium terebellae AY196709
M
0.28 0.21 0.99 0.82 0.97
0.97
.86
0.94
Actinocephaloidea
Hoplorhynchus acanthatholius FJ459750
0.98
0.19
Pyxinia crystalligera FJ459759
strain ATCC 50646
DNA D5P09C08 EF100358
ip t
0.48
us
0.34
Piroplasmida 8
Plasmodium reichenowi Z25819 Plasmodium gallinaceum Plasmodium malariae PFARGBAB Plasmodium ovale PFASSUR 0.44 Plasmodium gonderi AB287270 0.93 0.75 Plasmodium fieldi AB287282 cynomolgi PFARRNASSA 0.86 .32Plasmodium Plasmodium vivax VU07367 0.86 Plasmodium inui EU400396 Plasmodium fragile asexually expressed PFARRSS 0.88 0.7 Plasmodium vivax El Salvador sporozoite PVU07368 0.8 Plasmodium knowlesi PKU83876 Plasmodium coatneyi EU400393 Plasmodium chabaudi DQ241815 Plasmodium berghei PFARGSS
Ac ce pt e
0.39
Hematozoa
Haemosporida
0.57 0.98
0.95
Hepatocystis 6
Adelina dimidiata DQ096835 Klossia helicina HQ224956 0.35 Hepatozoon canis DQ111754 Hepatozoon catesbianae AF130361 0.59 Aggregata octopiana KC188342 0.56 0.84 Aggregata octopiana DQ096837 Aggregata eberthi KC188343 0.5 Aggregata eberthi DQ096838 Adelina grylli DQ096836 .99 Adelina bambarooniae AF494059
0.71
.39
anoxic marine DNA EF526760
Perkinsida 5 0.84
Mid-Atlantic Ridge hydrothermal sediment marine DNA AF530534 0.69
Ellobiopsis chattonii FJ593706 Spatulodinium pseudonoctiluca GU355685
0.99 Noctiluca scintillans AF022200 0.37
Kofoidinium sp. FG256 GU355681
Noctilucida
Kofoidinium cf. pavillardii GU355680 .23 0.08 Amoebophryidae 5 0.82 Dubosquellidae 4 0.93 .55 Syndinium sp. ex Corycaeus DO146406 .97 0.53
Adeleida Aggregatidae Adeleidae
Perkinsea 7
Dinozoa
Oxyrrhis marina AF482425 Oxyrrhida
pacifica JN873311 0.56 0.99 Psammosa Psammosa Psammosa atlantica JN873310 Thalassomyces fagei AY340590 0.12
0.36 .7
Adeleidae
Hematodinium sp. MF-2000 AF286023
4
Syndiniidae
17
Coccidiomorphea
Trichotokara 3 Gregarinomorphea
Ellobiopsida
Oxyrrhea
Dinozoa
Syndinea
Peridinea 9
Ciliophora 0.5
Page 60 of 60
S0932-4739(14)00053-4 http://dx.doi.org/doi:10.1016/j.ejop.2014.07.002 EJOP 25340
To appear in: Received date: Revised date: Accepted date:
28-5-2014 19-7-2014 21-7-2014
Please cite this article as: Cavalier-Smith, T.,Gregarine site-heterogeneous 18S rDNA trees, revision of gregarine higher classification, and the evolutionary diversification of Sporozoa, European Journal of Protistology (2014), http://dx.doi.org/10.1016/j.ejop.2014.07.002 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
*Manuscript
1
Gregarine site-heterogeneous 18S rDNA trees, revision of gregarine higher classification, and the evolutionary diversification of Sporozoa
ip t
Thomas Cavalier-Smith
d
M
an
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Department of Zoology, University of Oxford, South Parks Road, Oxford OX1 3PS, UK
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____________________
E-mail address: [email protected] (T. Cavalier-Smith)
Page 1 of 60
2
Abstract Gregarine 18S ribosomal DNA trees are hard to resolve because they exhibit the most disparate rates of rDNA evolution of any eukaryote group. As site-heterogeneous treereconstruction algorithms can give more accurate trees, especially for technically unusually
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challenging groups, I present the first site-heterogeneous rDNA trees for 122 gregarines and an extensive set of 452 appropriate outgroups. While some features remain poorly resolved, these trees
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fit morphological diversity better than most previous, evolutionarily less realistic, maximum likelihood trees. Gregarines are probably polyphyletic, with some ‘eugregarines’ and all
us
‘neogregarines’ (both abandoned as taxa) being more closely related to Cryptosporidia and
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Rhytidocystidae than to archigregarines. I establish a new subclass Orthogregarinia (new orders Vermigregarida, Arthrogregarida) for gregarines most closely related to Cryptosporidium and group
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Orthogregarinia, Cryptosporidiidae, and Rhytidocystidae as revised class Gregarinomorphea. Archigregarines are excluded from Gregarinomorphea and grouped with new orders Velocida
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(Urosporoidea superfam. n. and Veloxidium) and Stenophorida as a new sporozoan class Paragregarea. Platyproteum and Filipodium never group with Orthogregarinia or Paragregarea and
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are sufficiently different morphologically to merit a new order Squirmida. I revise gregarine higherlevel classification generally in the light of site-heterogeneous-model trees, discuss their evolution, and also sporozoan cell structure and life-history evolution, correcting widespread misinterpretations.
Keywords: Cryptosporidium classification; Orthogregarinia; Paragregarinea; Squirmidea; Stenophora; Terragregarina.
Page 2 of 60
3
Introduction Gregarine phylogeny is currently unclear, partly because of extreme variation (approximately 15-fold) in rate of evolution of their 18S rDNA. Despite that it is well known that existing higher gregarine classification based on 60-year-old evolutionary ideas (Grassé 1953) is
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phylogenetically unsound and has become purely a matter of convenience (Rueckert et al. 2011a). Therefore, having assembled the most extensive gregarine alignment to date in the accompanying
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paper for identifying a gregarine sequence contaminating a heterolobosean culture (Cavalier-Smith
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2014), I have carried out the first comprehensive phylogenetic analysis of gregarines and their alveolate outgroups using a site-heterogeneous model of base substitution (the CAT-GTR-
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GAMMA model) implemented in PhyloBayes (Lartillot and Philippe 2004). This model is evolutionarily more realistic than homogeneous models used for previous maximum likelihood
and Philippe 2008).
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(ML) gregarine trees, and copes better with extreme long branches (Lartillot et al. 2007; Lartillot
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It did indeed produce trees that are more congruent with both morphology and parasite-host preferences than previous gregarine trees, though some features remain unresolved. I discuss
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gregarine and myzozoan evolution generally in the light of these site-heterogeneous model trees, which show that classical gregarines comprise two distinct probably not directly related major clades, here made subclass Orthogregarinia and class Paragregarea, and that a third clade comprising Platyproteum and Filipodium (Rueckert and Leander 2009) is probably unrelated to either but groups weakly with either Perkinsozoa within Dinozoa or Apicomonadea and thus may have evolved independently of Sporozoa. I here establish a new order Squirmida for these two genera, which differ greatly from classical gregarines, and substantially improve gregarine higherlevel classification, dividing them into five orders not three as did Grassé (1953); only his order Archigregarinida is retained, the distinction between eugregarines and neogregarines being arteficial. A long-overdue change is establishing a new order for Cryptosporidium, here formally transferred from Coccidea to Gregarinomorphea as new subclass Cryptogregaria in the light of
Page 3 of 60
4 much recent morphological and sequence tree evidence (Valigurová et al. 2007). Overall Gregarinomorphea is now divided into three new subclasses and four orders (three new) and two new suborders; all 21 newly established suprafamilial taxa for broadly ‘gregarine’ taxa correspond with major clades on the site-heterogeneous tree (over two thirds strongly supported) and reflect
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reasonable morphological and host range distinctions. I also discuss the probable polyphyletic origins of gregarines in relation to the huge increases in cell and genome size of gut parasites
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compared with small cell size and genomes of intracellular parasites (Cavalier-Smith 1978, 1980,
us
1982) and differences in multiple fission, cilia and centrioles amongst the sporozoan classes.
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Material and Methods
Data selection and maximum likelihood (ML) analysis of the 652-eukaryote sample were as
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in the accompanying paper (Cavalier-Smith 2014). Analysis for two smaller taxon samples of 196 or 276 alveolates and 1577 well aligned nucleotides used both this and the site-heterogeneous CAT-
d
GTR-GAMMA model of PhyloBayes v. 3.3 (Lartillot and Philippe 2004) with two chains for many generations beyond the point of plateauing and 25% of trees removed as burnin. A third taxon
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sample of 1614 well aligned nucleotide positions for 137 Myzozoa only with all long-branch gregarines and coccidiomorphs excluded was analysed by the same site-heterogeneous model to obtain the most accurate myzozoan tree minimally perturbed by long-branches and distant outgroups; the two chains for this 137-taxon analysis and the 196-taxon analysis each converged well to a single topology, but the 276-alveolate analysis gave alternative slightly different trees for two weakly supported branch positions.
Results and Discussion Bayesian analysis using the site-heterogeneous CAT-GTR-GAMMA substitution model is evolutionarily more realistic than those available for ML (Lartillot and Philippe 2008) and often gives more reliable trees, especially for difficult-to-place taxa such as those with idiosyncratic long Page 4 of 60
5 branches or with very short internal stems (Brown et al. 2013; Roure et al. 2013). Though in these respects gregarines are one of the most phylogenetically challenging of all protist groups these superior site-heterogeneous methods have not been previously applied to them. To make discussion
Contrasts between homogeneous and heterogeneous rDNA trees
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of the trees more comprehensible the new classification is first summarised in Table 1.
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The eukaryote-wide Figure 1 like all other recent maximum likelihood trees groups marine eugregarines and the insect-host Gregarinoidea with archigregarines. One marine group (Velocida,
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comprising Veloxidium and Urosporoidea) is actually within archigregarines, thus partially
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supporting the classical view of Grassé (1953) that archigregarines are ancestral to eugregarines. But, contrary to that idea, most terrestrial gregarines (i.e. Stylocephaloidea/Actinocephaloidea; a
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well-supported (76%) clade here called ‘core orthogregarines’) weakly group with Rhytidocystis, and this clade with Cryptosporidium and not with archigregarines. Archigregarines are paraphyletic,
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some grouping weakly with Velocida and others still more weakly with a heterogeneous longbranch clade comprising Gregarinoidea and marine gregarines (Fig. 1). The basal branching order
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of all these clades has no significant support, and I argue that the phenotypically mixed long-branch grouping with only 1% support is probably a long-branch artefact - it is not found in any siteheterogeneous trees irrespective of their taxon sampling. The PhyloBayes CAT-GTR-GAMMA tree in Figure 2 focuses on Myzozoa only; to clarify relationships of the three main gregarine-like clades to other alveolates it omits the three longest-branch gregarine clades (Trichotokara and superfamilies Gregarinoidea and Porosporoidea). Like Fig. 1 it shows core orthogregarines (Stylocephaloidea/Actinocephaloidea) as a robust clade strongly supported as sister to Cryptosporidium, not to Rhytidocystidae as insignificantly by ML; rhytidocystids are strongly sisters of Cryptosporidium/core orthogregarines, the three groups together forming a robust gregarinomorph clade that excludes both archigregarines and Squirmidea (Platyproteum plus
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6 Filipodium). This Figure 2 site-heterogeneous tree strongly indicates that archigregarines are not sisters to Orthogregarinia, implying that classical gregarines are polyphyletic. As long supposed, Archigregarinida are paraphyletic in Fig. 1 (ML) and in all CAT-GTRGAMMA trees, even if all long-branch gregarine clades and alveolate outgroups are included (Fig.
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3). Archigregarines are clearly ancestral to the robust clade Velocida, comprising a relatively small subset of aseptate gregarines (its subclade Urosporoidea former eugregarines). Figs 2 and 3 also
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show for the first time that the millipede-inhabiting eugregarine Stenophora, not included in recent trees for marine gregarines, is sister to classical archigregarines plus Urosporoidea, not to the
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insect-infecting core orthogregarines; in that respect ML and CAT trees agree. Though Stenophora
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weakly groups with Gregarinoidea on Fig. 1, I give more weight to its grouping with archigregarines and Velocida (together here called core paragregarines) on the probably more
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accurate CAT trees, so core paragregarines and the new order Stenophorida are here grouped as a new sporozoan class Paragregarea. Figs 2-3 suggest more weakly that Paragregarea may not even
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be sister to Gregarinomorphea, but possibly sister to Gregarinomorphea plus Coccidiomorphea (Fig. 2) or to Coccidiomorphea alone (Fig. 3); if either is correct, gregarines are polyphyletic. In all three
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trees the free-living predatory, myzocytotic, apicomonad flagellates (here shown as including the algae Chromera and Vitrella as clearly separate lineages) are the earliest diverging Apicomplexa, sisters to Sporozoa. Both CAT trees (Figs 2-3) agree with the ML tree (Fig. 1) in showing more clearly than before that the recently discovered Squirmida are probably not gregarines or even sporozoa; their precise position however is inconsistent, ML placing them weakly as sisters of all Dinozoa, whereas Fig. 2 puts them within Dinozoa and Perkinsozoa weakly as sister to the also parasitic Perkinsea and Fig. 3 places them within the apicomplexan Apicomonadea. Site-heterogeneous Figure 3 substantially rearranges the long-branch gregarine clades compared with present and past ML trees and past homogeneous Bayesian trees in a way that makes much more sense in terms of both host specificity and morphology than other gregarine trees; and reassuringly it is also largely congruent with the site-heterogeneous Figure 2 that
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7 excluded long-branch gregarines and ciliates, showing that the well-supported basal branching order of Figure 2 is not distorted when long-branch taxa are added, even though their support values unsurprisingly drop. This strongly suggests that less evolutionary realistic site-homogeneous substitution models do generally misplace many long-branch gregarines through artefactual mutual
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attraction, making past trees misleading. Figure 3 includes the gregarines Pyxinia (omitted in Figs 1 and 2 because of its exceptionally long branch for terragregarines), showing it to be an
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actinocephaloid, and like Fig. 2 Apicystis (not in Fig. 1). The CAT trees also include environmental sequence BOLA566 (Dawson and Pace 2002) to check whether it is a gregarine; it clearly is, being
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consistently (never strongly) a distant sister to Stenophora – its inclusion in Figs 2-3 is another
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reason for thinking they probably more accurately place Stenophora than did Fig. 1.
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Are long-branch gregarines orthogregarines or paragregarines?
Figure 3 places the long-branch insect host Gregarinoidea within core orthogregarines as
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sister to superfamily Actinocephaloidea (0.46 PP) to form a clade entirely of gregarines that infect terrestrial arthropods or oligochaetes; core orthogregarines and Gregarinoidea are here united as a
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new taxon Terragregarina, comprising all terrestrial gregarines but Stenophora. Fig. 3 also weakly groups the extremely long-branch crustacean gregarines (Porosporoidea) with Stylocephaloidea (0.68 PP). Terragregarines and Porosporoidea together form a weakly supported (0.34) clade, here made a new order Arthrogregarinida within Orthogregarinia (an extremely weakly supported clade in Fig. 3), assuming that all share an arthropod-gut-dwelling common ancestor. Despite its weak support the arthrogregarine grouping is evolutionarily more reasonable than the even weaker exclusion of Gregarinoidea from Orthogregarinia and their probably artefactual grouping with other long-branch gregarines in the ML tree (Fig. 1). Terragregarines probably ancestrally infected insects and/or arachnids and later one branch colonized earthworms. Fig. 2 also weakly (PP 0.76) groups the very-long branch polychaete-host Trichotokara with the polychaete host Paralecudina and numerous environmental sequences, a clade with low (49%)
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8 support in ML (Fig. 1), now family Trichotokaridae. Trichotokaridae is sister to the polychaete host Polyplicariidae and further environmental sequences; despite weak support (0.62) this grouping of two families of species from marine polychaete hosts (plus one from a foraminiferan host) makes good evolutionary sense, so I make this whole clade a new order Vermigregarida, grouped with
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Arthrogregarinida as subclass Orthogregarinia. In the 196-alveolate tree (Fig. 3), Trichotokaridae support rose from 49 to 71% and 0.72; with ML they moved to become very weakly sister to the
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Selenidium bocardellae clade and with CAT sister to the unidentified environmental clade.
A CAT tree with 83 coccidiomorph sequences (Fig. 4), not just seven as in Fig. 3, also put
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Porosporoidea within Orthogregarinia, but as sister to core orthogregarines only. However, the Fig.
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4 analysis chains did not converge though each plateaued (maxdiff 0.376); the other chain for the same taxa showed only two differences: Porosporoidea was weakly (0.37) sister to Coccidea, not
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core orthogregarines, and Squirmidea sister to all Apicomplexa (not just Apicomonadea as in Fig. 4 or within Apicomonadea as in Fig. 3). Positions of Gregarinoidea and Trichotokara are also
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unstable to outgroup taxon sampling. Fig. 4 (both chains) put Gregarinoidea with the millipede-host Stenophora within archigregarines, i.e. within paragregarines not orthogregarines. In Fig. 4
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Trichotokaridae broke up into two: the relatively short-branch Paralecudina clade remained within the deepest branch of Orthogregarinia, but the extremely long-branch Trichotokara grouped instead with the somewhat long Oxyrrhis branch within dinoflagellates – almost certainly a long-branch artefact as it was not seen in any other trees and makes no evolutionary sense. Exclusion of Gregarinoidea from Orthogregarinia in Fig. 4 may be a long-branch artefact possibly caused by including so many long-branch coccidiomorphs (unlike Fig. 3). Figs 3 and 4 both imply that the crustacean-host Porosporoidea are more closely related to Terragregarina (grouped with them as Arthrogregarinida) than is the polychaete-host Trichotokaridae.
Detailed Orthogregarine Phylogeny
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9 All my trees (and many previously published ones) indicate that Orthogregarinia, especially core orthogregarines, are more closely related to Cryptosporidium and Rhytidocystidae than to archigregarines. In Fig. 3 Pyxinia groups with Hoplorhynchus and the ATCC 50646 clade with weak support (35% BS; 0.28 posterior probability) and Apicystis is sister to Mattesia with 100% or
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0.99 support (also 0.99% support in Fig. 4, both chains). In Fig. 4 the Pyxinia/Hoplorhynchus/ ATCC 50646 cluster also includes two additional environmental DNA sequences (EF100358/63);
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in Fig. 3 they are sister to Hoplorhynchus plus ATCC 50646 (0.44 support), but with ML moved just below the Hoplorhynchus/Monocystis last common ancestor. Addition of Pyxinia and Apicystis
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does not change the positions of the main gregarine clades on ML trees, though Apicomonadea
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joined Squirmidea as a second sister group to Dinozoa (low support). Within Terragregarina it induced some rearrangement, the Psychodiella/Ascogregarina and the
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Mattesia/Apicystis/Pseudomonocytis subclades being more strongly supported and now sisters (54%, 0.82 support). Moreover bootstrap support for some terragregarine backbone bipartitions
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dropped in Fig. 3: the highest ones (90, 87, 98, 79) became 74, 87, 84, and 55 respectively (Bayes: 0.84, 0.99, 0.99, -). Presumably the drop for the first bipartition is caused by adding Pyxinia, whose
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branch length is approximately twice that of ATCC 50646, such long branches being noted for a tendency to move somewhat too low in trees, whereas that for the fourth reflects the Fig. 3 evidence that Stylocephalidae are not sisters of Actinocephaloidea alone but of Actinocephaloidea plus Gregarinoidea. In marked contrast to the sparse distance and parsimony trees of Alarcón et al. (2011) and the ML, Bayesian and parsimony trees of Clopton (2009), which grouped Pyxinia strongly with Stenophora, I found no evidence for placing it there or with any marine gregarines. As Stenophora is a diplopod parasite and Pyxinia and Hoplorhynchus are insect (beetle or dragonfly) parasites (prior to Clopton (2009) both in Actinocephalidae), and major clades of gregarines show strong host group specificity (Rueckert et al., 2013) – even stronger than before in my heterogeneous trees, it is likely that the earlier grouping of Pyxinia with Stenophora on earlier less well sampled trees (only 51 or 27 gregarines respectively; 122 here) was a long-branch artefact.
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10 Transfer of Pyxinia from Actinocephalidae to Stenophoroidea (Clopton 2009) was probably therefore incorrect and is rejected here (Table 1). Its transfer then to Monoductidae may also be incorrect as two of the three genera of that family (Monoductus, Stenoductus) are millipede parasites like Stenophora, so their placement in Stenophoroidea may be correct; if so, their similar
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oocyst dehiscence, extruding cells in chains, may be convergent with Pyxinia – however until sequences are available we cannot rule out the less likely possibility that they also belong in
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Terragregarina and are related to Pyxinia. Within Gregarinoidea, Blabericolidae nest within
Gregarinidae in CAT trees (Fig. 3 and 4), but Gregarinidae are weakly holophyletic by ML (Fig. 1).
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Arthrogregarida from arthropods and oligochaetes are weakly sister to Vermigregarida from
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polychaetes, which also embraces an unidentified clade that includes a parasite or contaminant of the foraminiferan Ammonia (still wrongly annotated in GenBank as a foraminiferan though I
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discovered that error 19 years ago, and several publications have corrected it, notably Pawlowski et al. (1996)). The Polyplicariidae subclade of Vermigregarida was identified by Wakeman and
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Leander (2013) after I completed my analyses, so I was unable to include their three Polyplicarium sequences, which all grouped with clone CCA38 rather than CCI31. As Polyplicarium are
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symbionts of capitellid polychaetes, the grouping here of CCA38/CCI31 with the polychaete symbiont Trichotokaridae supports the new order Vermigregarida and suggests that like archigregarines they were ancestrally polychaete symbionts. Wakeman and Leander (2013) did not include either Trichotokaridae or the Ammonia symbiont subclade so did not reveal this relationship; their ML tree did however have insignificantly supported terragregarine and orthogregarine clades and grouped Squirmidea with Dinozoa but had no basal resolution, partly because only 1007 nucleotides and 79 Myzozoa were included.
Paragregarine Phylogeny Archigregarines and their relatives (probably derived Velocida, and Stenophora plus BOLA 566) form a clade (Paragregarea: 0.23 support Fig. 3) grouping not with orthogregarines but with
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11 Coccidia in Figs 3 and 4 or as sister to Coccidiomorphea plus Gregarinomorphea in Figure 2. BOLA 566 was moderately supported as sister to Stenophora in the best CAT trees (Figs. 2, 3), not sister to Selenidium terebellae as weakly in Rueckert and Leander (2010) or contradictorily to rhytidocystids in Leander and Ramey (2006), neither of which included Stenophora. The weakly-
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supported branching order of Selenidium terebellae and the Selenidium pisinnus/orientale clade in Fig. 1 of the accompanying paper (Cavalier-Smith 2014) was the same in Fig. 4 (CAT) but Fig. 3
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suggests instead that Selenidium pisinnus/orientale may be more closely related to the
Veloxidium/Urosporoidea clade than to the longer branch Selenidium serpulae clade. Though all my
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trees place Velocida within Selenidium, as do most but not all published trees, statistical support for
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this is so low that one cannot rule out the alternative possibility that Velocida are sisters to rather than derived from Archigregarinida.
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My trees confirm that two environmental DNAs from anoxic habitats once claimed to represent novel kingdoms (CCA5; LEMD119: Dawson and Pace 2002; Stoeck and Epstein 2003)
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are also gregarines as I showed previously using only 1044 nucleotide positions and many fewer gregarines (Cavalier-Smith 2004a). Berney et al. (2004) identified some of these as gregarines;
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more precise positions of many of them within the gregarine radiation were identified by Rueckert et al. (2011a), and largely confirmed here.
Phyletic distinctiveness of Squirmidea
Strikingly, Squirmidea (whose branch lengths are much shorter than for many gregarine clades, and are therefore unlikely to be grossly misplaced) never group with other gregarines or any Sporozoa on any of my trees or on any published trees with enough outgroups to judge adequately their affinities. On the ML tree (Fig. 1) they are weakly sister to Dinozoa; on the myzozoan CAT tree omitting long-branch taxa (Fig. 2), perhaps the most reliable indicator of their affinities, they are within Dinozoa, sister to the parasitic Perkinsea. CAT trees including long-branch gregarines place them contradictorily: weakly sister to Apicomonadea (Fig. 4) or all Apicomplexa (second
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12 chain for Fig. 4 taxa) or within Apicomonadea as sister to the Chromera/Voromonas/’Colpodella’ clade (Fig. 3: 0.3 support). Such clear, consistent exclusion of Squirmidea from Sporozoa in all trees could not have been detected on published ML gregarine-rich trees as non-sporozoan apicomplexa were always excluded; as the first taxonomically sparse ML tree insignificantly
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grouped them with long-branch archigregarines, they have been assumed to be gregarines even though there is no evidence for a sporozoan style apical complex (and morphological reasons to
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doubt that – see below) or any positive morphological evidence for inclusion in archigregarines.
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Myzozoan Phylogeny
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On Figure 3 Dinozoa are extremely weakly sisters to Sporozoa only, not sisters to all Apicomplexa, the non-sporozoan Apicomplexa being markedly paraphyletic, with Colpodella
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(=Alphamonas) edax (formerly grouped with Voromonas in Voromonadida: Cavalier-Smith and Chao 2004) plus an environmental DNA being sister to all other Myzozoa. Though basal branching
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of Myzozoa is too weakly supported for confidence in all details of this topology, the marked differences in several features of my heterogeneous trees indicate that it is unwise to take the deep
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gregarine branching order on homogeneous model trees too seriously, as the longest branches tend to group together as in Fig 1, I argue artefactually, which they certainly do not with the more realistic site-heterogeneous model. Use of heterogeneous models is advisable for future gregarine studies given the huge diversity in evolutionary rates amongst lineages. However, the remaining but relatively much smaller differences with taxon-sampling in Figs 3 and 4 emphasize the difficulty of resolving deep gregarine and myzozoan phylogeny from 18S rDNA alone and show that even CAT trees cannot iron out all inconsistencies among taxon samples for this evolutionarily exceedingly unclock-like molecule. Nonetheless, it is useful to compare these results with those of Fig. 1 to see which groupings are stable and which are not. Fig. 4 showed the tree from the chain that gave results most similar to Fig. 2; the internal phylogeny of all clades collapsed on Figs 3 and 4 was precisely the same as on Fig. 2, so they were internally stable.
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13 It is evident that both Vitrella and Chromera are simply photosynthetic apicomonads and do not merit a separate phylum; photosynthesis was clearly lost at least three times within Apicomplexa. I regard Chromerida as an order within Apicomonadea that on all my trees is sister to Voromonas plus species presently called ‘Colpodella’, but which for reasons to be explained
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elsewhere are generically distinct; by contrast Vitrella is more closely related to Colpodella (=Alphamonas) edax, which I now consider a true Colpodella. In Figs 2 and 4 the Colpodella
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(=Alphamonas) edax/Vitrella apicomonad subclade) was holophyletic (not paraphyletic like in Figs 1, 3) and the five hypersaline Lake Tyrell sequences formed a sister clade to marine Vitrella (not to
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the C. edax freshwater clade as in Heidelberg et al. (2013) fig. 5), this clade in turn being sister to
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the freshwater clade including C. edax. However Apicomonadea were paraphyletic with the Voromonas/Chromera subclade branching weakly more deeply than the C. edax/Vitrella subclade.
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Paraphyly of apicomonads was also shown by a chloroplast gene tree but the other way round (Janouškovec et al., 2012). If either is true, sporozoa definitely evolved from apicomonad ancestors.
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However branches within apicomonads are so close that multiple transcriptomes or genomes from both the Vitrella/C. edax and Voromonas/Chromera subgroups are necessary to resolve their
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topology and thus clarify the origins of sporozoa from such predatory flagellates.
Implications of Improved Gregarine Phylogeny for Classification The shallow nesting of neogregarines (e.g. Syncystis, Mattesia) within terrestrial gregarines in Figure 1 of the accompanying paper (Cavalier-Smith 2014) and on Figure 2 supports Grassé’s idea that neogregarines evolved from eugregarines by evolving merogony (Grassé 1953). However, their nesting shallowly in three different places amongst both ‘septate’ and aseptate eugregarines convincingly shows that neogregarines are multiply polyphyletic. Likewise the fact that on Figs 1-4 the classically eugregarine Velocida form a derived clade within archigregarines, that Stenophora is sister to archigregarines plus Velocida, and that other eugregarines are widely dispersed as the paraphyletic basal group of Terragregarina, or still more basally, suggests that classical
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14 eugregarines are also polyphyletic. Thus the even greater taxon and intramolecular sampling here strongly supports the recent conclusion that retention of the classical subdivision of gregarines into three orders (Archigregarinida, Eugregarinida, Neogregarinida: Adl et al. 2012; Grassé 1953) is ‘based more on convenience than phylogenetic relationships’ (Rueckert et al. 2011a). Sufficient
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information now exists to revise gregarine higher classification radically along sounder phylogenetic lines (Table 1), abandoning the polyphyletic neogregarines and eugregarines as taxa.
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Archigregarinida are probably paraphyletic, but retained as a morphologically well-defined group. Other classical gregarines make up a single new subclass Orthogregarinia comprising both aseptate
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and ‘septate’ gregarines subdivided into two new orders: Vermigregarida for two clades of aseptate
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polychaete gregarines, and Arthrogregarida for the remainder, predominantly inhabiting arthropods, rarely oligochaetes or molluscs, and largely ‘septate’. Table 1 includes in less detail other major
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taxa of the apicomplexan superclass Sporozoa as here revised.
A key feature of the new classification is the transfer of Cryptosporidium from Coccidia to
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class Gregarinomorphea, as a new order and subclass. The merits of such transfer were argued in detail by Barta and Thompson (2006) and need not be repeated. It is fully supported by multigene
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trees based on 10,753 amino acids that group Cryptosporidium with the actinocephalid terragregarine Ascogregarina with strong support (Templeton et al. 2010) and by the remarkable similarity of host-attachment structures in both groups (Valigurová et al. 2007). A second nongregarine group, previously treated as order Agammococcidiida that includes Rhytidocystidae, is clearly more closely related to both Cryptosporidium and orthogregarines than to archigregarines on all my trees and all sufficiently well sampled published trees; here I establish a separate subclass Histogregaria for this order and place it within Gregarinomorphea. As Archigregarinida never grouped with either Cryptosporidium or core orthogregarines (superfamilies Stylocephaloidea and Actinocephaloidea), which always group with Cryptosporidium to the exclusion of Archigregarinida, the conclusion that classical gregarines excluding Cryptosporidium and rhytidocystids are polyphyletic is now inescapable. Therefore a new sporozoan class Paragregarea
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15 is established to group archigregarines with those eugregarines that sequence trees consistently exclude (albeit usually with low support) from the Gregarinomorphea clade. Archigregarinida are marine non-septate, highly contractile, non-gliding gregarines that probably have apicoplasts, as organelles indistinguishable from apicoplasts are clearly visible in
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Selenidium hollandei and pendula (membranous vesicles in Figs 28-30, 39 of Schrével 1971). I postulate that apicoplasts were lost in the last common ancestor of Cryptosporidium and
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orthogregarines (arrow on Fig. 2), being apparently present in Rhytidocystis polygordiae and
sthenelais (Leander and Ramey 2006; Porchet-Hennere 1972). The only non-archigregarines
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consistently branching within archigregarines in all my trees are Velocida. Adl et al. (2012) put
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Veloxidium in archigregarines, but put Difficilina, Lecudina, Lankesteria, Pterospora and Lithocystis contradictorily in Eugregarinida. As all my and all published trees place the latter five
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genera (collectively a robust clade, superfamily Urosporoidea) together with Veloxidium with 100% support, it is cladistically wrong to include only Veloxidium within archigregarines. I therefore
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establish order Velocida for this extremely robust Veloxidium/urosporoidean clade. For a recently recognized clade of non-septate polychaete gregarines, e.g. Trichotokara,
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Paralecudina (Rueckert and Leander 2010), I establish a new family Trichotokaridae on account of their isolated phylogenetic position. Positions of the three longest branch taxa (Trichotokaridae, Porospororoidea, Gregarinoidea) are especially hard to establish on 18S rDNA trees probably because they have long unbroken stems, as is that of Stenophora with no close relatives breaking its much shorter branch. Though they all group together with ML (Fig. 1), this is probably a longbranch artefact as they occupy four separate positions on the heterogeneous model tree of Fig. 3, where all three longest branches group separately within the short-branch orthogregarines and Stenophora with, but not within, Archigregarinida. Though weakly supported (here 49%; 43% in (Rueckert and Leander 2010); 59% in (Rueckert et al. 2013), Trichotokaridae are a clade on all recent trees except Figure 4 where Trichotokara oddly moved outside Apicomplexa to join the long-branch Oxyrrhis. But the position of Trichotokaridae varies: in ML trees it was originally
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16 weakly sister to Urosporoidea (Rueckert and Leander 2010), to Urosporoidea plus Porosporoidea (Rueckert et al., 2011a); these contradictory positions or its placement in a mixed long-branch clade (Fig. 1) lack statistical support or biological rationale. By contrast the moderately supported grouping of the polychaete gregarines Polyplicariidae and Trichotokaridae on the evolutionarily
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more realistic heterogeneous tree (Fig. 3), irrespective of whether they have long or short branches, makes evolutionary sense, so I establish order Vermigregarida for this clade. Placing the three
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longest branch clades in distinct superfamilies or orders seems well justified. Their assignment to higher groups is more debatable, but done consistently with morphology and host preference and
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what I judge the best trees, but because of the low bootstrap support needs testing by sequencing at
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least another molecule like Hsp90 less subject to long-branch problems, and ideally by transcriptome analysis and multigene trees. I hope this revised classification stimulates such tests.
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The major long-branch clade comprising insect gregarines, for which I adopt the superfamily name Gregarinoidea used by Chakravarty (1960), consists of the invariably robustly
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related families Gregarinidae (ancestral) and Blabericolidae (derived). In the ML tree that did not include BOLA 566, this groups weakly with the millepede gregarine Stenophora. However CAT-
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model Figs 2-4 slightly more strongly group Stenophora with environmental sequence BOLA566, this clade appearing very weakly sister to Archigregarinida. The third subclade (crustacean gregarines: Porosporoidea) is sister to the short-branch terrestrial gregarines (Actinocephaloidea plus Stylocephaloidea) on both well-converged heterogeneous trees (Figs 2, 3). Gregarinoidea corresponds with terrestrial subclade II of Rueckert et al. (2011a) and Actinocephaloidea with their terrestrial subclade I. However, because Gregarinoidea grouped inside Actinocephaloidea on a later ML tree (Rueckert et al. 2013) they altered the labelling to include a subset of their subclade I in subclade II. However subclade II sensu Rueckert et al. (2013) is evidently a phylogenetic artefact of an ML tree with inadequate taxon (76 gregarines) and sequence (988 nucleotides not 1540 or 1577 as here) sampling. A later basally unresolved 1085-nucleotide tree did not put Gregarinoidea within Actinocephaloidea, and the authors reverted to the clade I and II labelling of Rueckert et al. (2011a)
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17 (Wakeman and Leander 2013b). None of my much better sampled trees place Gregarinoidea within Actinocephaloidea. Heterogeneous-model Figure 3 places them as sister to Actinocephaloidea, without Stenophora, with low (0.46) support. My trees have high 97% BS (Fig. 1) and 0.97 PP (Figs 2-3) support for holophyly of Actinocephaloidea (terrestrial gregarines sensu Rueckert et al.
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(2011) and Wakeman and Leander (2013b)), apparently a very robust clade that includes the new ATCC/stained glass window environmental clade (Cavalier-Smith 2014). The non-converging
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CAT-tree (Fig. 4) grouped Gregarinoidea with Stenophora and BOLA 566 very weakly within Archigregarinida, like ML, both I suspect long-branch artefacts. Stenophora and Pyxinia were
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omitted from all recent trees focusing on marine gregarines only.
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To exclude potentially misleading artefacts caused by long-branch gregarines Fig. 2 used 137 Myzozoa alone, chosen from the Fig. 4 taxa by excluding the longest-branch gregarines and
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long-branch Aggregata and Plasmodium, but supplemented by additional environmental apicomonad sequences (including five from Heidelberg et al. (2013) who obtained 1406
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apicomonad sequences from hypersaline Lake Tyrrell) and two extra Oxyrrhis sequences to break up their long branch. Archigregarines were represented only by the shortest branch Selenidium
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species. It also excluded a few of the fastest evolving sites that were used for Figs 3 and 4 trees to focus more on deep-branch topology. Fig. 2 converged well (maxdiff 0.128) and still placed gregarines and Squirmidea in three separate clades: with CAT Orthogregarinia was sister to Cryptosporidium (0.92 support), rather than to Rhytidocystidae as in other trees; and Squirmidea were not even in Apicomplexa, but weakly sister to Perkinsea (0.47 support, but much higher 0.85 support for including them in Dinozoa, not Sporozoa). Support was strong for a Stenophora/Selenidium clade (0.89) (new class Paragregarea) that was sister to Coccidiomorphea plus Gregarinomorphea; for Paragregarea to move to become sister to Orthogregarinia they would have to cross three nodes with support of 0.65, 0.92, and 0.92. Thus the heterogeneous tree with artefact-inducing long-branches excluded fairly strongly supports the conclusion that gregarines as previously understood are polyphyletic and evolved from smaller celled ancestors three times
Page 17 of 60
18 independently, once within Dinozoa and twice within Sporozoa. Virtually all other published trees have shown the same triphyletic ‘gregarine’ topology, but as they all included so many long-branch taxa and did not use heterogeneous trees or nearly so extensive outgroups, that topology was much more weakly supported and the conclusion of triple polyphyly much less convincing than it now is. I conclude that classical aseptate archigregarines are probably specifically related to but not
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ancestral to the ‘septate’ Stenophorida (not included in most published trees) but are ancestral to the
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also aseptate Velocida; they are apparently not ancestral to or sisters of orthogregarines.
One limitation of present knowledge of gregarine phylogeny is the absence of sequence data
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or ultrastructure for the non-septate blastogregarine Siedleckia (only four species), which unlike
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true gregarines show no evidence of szygy. Grassé (1953) considered them phylogenetically distinct from but distantly related to gregarines. However Levine et al. (1980) included them as a suborder
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within eugregarines primarily because of the absence of merogony, but that is not a sound reason as it does not distinguish them from Cryptosporidium or Histogregaria. As their anisogamy suggests a
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possible closer relationship with Cryptosporidium I prefer to follow Grassé (1953) and retain them as a separate order, as they might be a very early diverging group not within any known rDNA
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clade. Though their general body form and life style within polychaete guts is reminiscent of aseptate gregarines, the latter are probably polyphyletic, so I put blastogregarines incertae sedis in Sporozoa. Unsurprisingly, a preliminary report indicates S. nematoides branching with gregarines or Coccidia depending on taxon sampling and method (Diakin et al. 2012).
New myzozoan class Squirmidea
Previously Filipodium was placed in Lecudinidae, then transferred to Selenidiidae because it robustly grouped with Platyproteum vivax (formerly considered a Selenidium) as a clade here called squirmids, which once weakly grouped with S. terebellae on an ML tree within Selenidium (Rueckert and Leander 2009). Less poorly sampled ML trees (Rueckert and Leander 2010; Rueckert et al. 2011a,b, 2013; Wakeman and Leander 2013a, b) all agree with my even better
Page 18 of 60
19 sampled trees in placing squirmids more deeply than archigregarines, orthogregarines, Cryptosporidium, and coccidiomorphs (with both ML and heterogeneous CAT-model trees); the also weakly sampled ML tree of Rueckert et al. (2010) excluded Filipodium but grouped Platyproteum with a miscellaneous set of long-branch gregarines as sister to S. serpulae (but not in
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the corresponding Bayesian tree). There is no convincing sequence tree evidence for placing squirmids within Archigregarinida, and no cellular reason other than their pendular movement that
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could easily be convergent.
Indeed there is no evidence from ultrastructure concerning their apical complex – whether
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they have even have one or not, and if they do whether it is more like that of Apicomonadea than
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Sporozoa. Thus we do not know that they are even Sporozoa, as now defined as all Apicomplexa that descended from the first one that evolved a true symmetric conoid (Cavalier-Smith and Chao
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2004). Figs 1-4 all make it likely (but not certain) that they are not gregarines or even Sporozoa. It might be suggested that the consistent non-grouping of Squirmida with other gregarines here and in
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every other well-sampled tree is a long-branch artefact caused by the extremely diverse evolutionary rates within marine gregarines, and the absence of close relatives to Squirmida.
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However, their branch is much shorter than many branches that the CAT model places sensibly within archigregarines and orthogregarines and distinctly shorter than the long-branch Myrionecta branch that it correctly places within Ciliophora (Fig. 3; unlike ML which often misleadingly excludes it from Ciliophora leading to past treatment of its environmental sequences CCW75 and CCW100 and DH145-EKD11 as novel kingdoms (Dawson and Pace 2002) or phyla (LópezGarcía et al. 2001; Richards et al. 2005), until their ciliate nature was discovered (Bass et al. 2009)). We should therefore be open to the likelihood that squirmids are a truly independent myzozoan lineage that evolved large-celled gut parasites. Data from other gene sequences are vital to test that, but even now they are sufficiently distinctive genetically and morphologically to merit separation from all other apicomplexans as new order Squirmida.
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20 The limited thin section ultrastructure for Filipodium makes it possible that they are related to archigregarines as pellicle structure is similar except for the presence of hair-like projections. However the similarities consist simply of a wavy or shallowly folded surface (Hoshide and Todd 1996; Schrével 1971), which could easily have evolved independently, as all trees suggest. In
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contrast to some lecudinid archigregarines (e.g. Lankesteria) and some actinocephaloid orthogregarines, which independently lost longitudinal epicytic folds, there is no reason to postulate
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that squirmids had an ancestor with such folds.
Adl et al. (2012) retained Filipodium and Platyproteum in Archigregarinorida as did
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Schrével and Desportes (2013) ‘temporarily’ within Selenidiidae in Archigregarinorida. However,
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there is no sound phylogenetic evidence for that. Though the apical end of the squirmidean trophont is sometimes called a mucron, this term is best avoided as it begs the question of homology with
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that of gregarines for which there is no evidence, and strong contraindications. Its unusual asymmetry suggests that it will not be ultrastructurally the same as in archigregarines and
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orthogregarines, and that trophonts may lack conoids. Though that does not prove they are absent in all life cycle stages, as conoids can be lost secondarily in some developmental stages of some
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gregarines, the simplest interpretation of all my Bayesian trees (branching with Perkinsea Fig. 2; within Apicomonadea Fig. 3; as sister to Apicomonadea or Apicomplexa Fig. 4) and their rostral asymmetry is that Squirmidea have an asymmetric pseudoconoid like Apicomonadea and Dinozoa (see Okamoto and Keeling, 2014). I therefore establish a new class Squirmidea to contain Squirmida. I exclude Squirmidea from Sporozoa and tentatively place it with Perkinsea in Perkinsozoa, now a dinozoan superclass (Cavalier-Smith 2013b):
Class Squirmidea (in superclass Perkinsozoa Norén and Moestrup in Norén et al. (1999) stat. n. Cavalier-Smith 2013a) Squirmidea cl. n. Diagnosis: lumenal parasites of sipunculid intestines, some with intracellular phase; trophonts flattened throughout (Platyproteidae) or just at apical end (Filipodium); epicytic
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21 folds if present shallower than in Gregarinomorphea and underlain by longitudinal microtubules without complex fibrillar skeleton; unlike orthogregarines and Urosporoidea without gliding motility; trophonts with extensible asymmetric rostrum, not a symmetric apical mucron as in many gregarines. Merogony, sexual reproduction, presence or absence of apical complex, cilia, and
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rhoptries unknown. Etymol: squirm E., referring to their squirming motility. Sole order Squirmida
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ord. n. Diagnosis: as for class Squirmidea.
Filipodiidae fam. n. Diagnosis: highly contractile (bending, twisting) trophonts covered in hair-like
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projections (cytopili) supported by tubular extensions of the cortical alveoli, but without a
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microtubular or fibrillar axoneme; in some species at least cytopili arranged in longitudinal rows within every third or fifth longitudinal groove; cell apex flattened, often laterally expandable as a
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blunt cornered triangle wider than body. Type and sole genus Filipodium Hukue, 1939. Platyproteidae fam. n. Diagnosis: trophozoite with temporary fine transverse striations, associated
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with peristaltic motility, but no hair-like projections or longitudinal folds; bending and peristaltic shape changes; an extensible asymmetric flattened rostrum; with lumenal and intracellular stages;
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no evidence for merogony. Type and sole genus Platyproteum Rueckert and Leander, 2009.
Revised Gregarine Taxonomy
Classically gregarines were called Gregarinae (Haeckel 1866; Pritchard 1861), Grégarines (Claparède 1861), Gregarinidae (Lankester 1863) or Gregarinida (Pritchard, 1861) treated as a class in the animal kingdom even before Coccidea were recognized or Sporozoa invented to embrace both, though sometimes only treated as an order within Rhizopoda (Saville Kent 1880-1882). Other spellings are subclass Gregarinidea (Lankester 1885), Gregarinomorpha (Grassé 1953), or Gregarinia (Levine et al. 1980) or Gregarinasina (Perkins 2002 dated 2000) if ranked as subclass or Gregarinea when a class (Cavalier-Smith 1993a; Cavalier-Smith and Chao 2004). I here adopt the broader class name Gregarinomorphea and use the vernacular gregarinomorphs to refer collectively
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22 to all its members (i.e. gregarines, Cryptosporidium, and Histogregaria), and recommend that ‘gregarines’ is continued as in traditional usage for classical Gregarinea alone (i.e. orthogregarines and paragregarines), allowing their contrast collectively with Cryptosporidium, Histogregaria, and Squirmidea. As Gregarinomorphea include both many classical gregarines and two orders that are not gregarines, its suffix -morph usefully emphasises that it includes both gregarines and non-
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gregarine organisms (as in ‘nucleomorph’ which means related to, but not the same as, a nucleus);
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for the same reason I adopt Doflein’s class name Coccidiomorphea to include both classical
coccidia and organisms like Hematozoa and nephromycids that are not coccidia, but related to or
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derived from them. Diagnoses of new taxa follow:
Superclass Sporozoa Leuckart, 1879
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Gregarinomorphea Grassé, 1953
Histogregaria subcl. n. Diagnosis: apicoplast present; gametogony and merogony both absent;
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inhabit invertebrate tissues, not gut lumen. Etymology: histos Gk tissue, because of habitat. Sole order Histogregarida nom. nov. pro order Agamococcidiorida Levine, 1979; substitution because
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Levine’s name misleadingly implies they are coccidia. Cryptogregaria subcl. n. Diagnosis: epicellular parasites of vertebrate epithelia; initial round trophont becomes spherical whilst being surrounded by host lamellar extension of plasma membrane; after rhoptry and microneme discharge a single-membrane anterior vacuole develops that fuses with host plasma membrane after trophozoite plasma membrane fuses with that of host; the anterior vacuole membrane on the trophozoite cytosol side folds extensively to construct a feeder organelle with no homologues in Coccidomorphea, but some similarities to certain gregarines. Anisogamous; microgametes non-ciliate. Oocytes have 4 naked sporozoites without sporocysts. Unlike coccidiomorphs, Histogregaria, and archigregarines, lack apicoplast. Etymology: crypto Gk hidden: refers to hidden, epicellular location of the trophozoite (i.e. not in
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23 gut or coelomic lumen as in most mature gregarines) and to sole genus Cryptosporidium. Sole order Cryptogregarida ord. n. Diagnosis: as for subclass Cryptogregaria.
Orthogregarinia subcl. n. Diagnosis: Trophozoites ambicellular, i.e. partially embedded in host
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cell, partially extracellular (Cavalier-Smith 2004b). Aseptate or ‘septate’ gregarines with mucron but no apicoplasts; invertebrate hosts (arthropods or annelids); gamonts of similar size. Unlike
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archigregarines or Platyroteum, usually not strongly flattened. Well-developed longitudinal epicytic folds in gut lumenal forms; motile by gliding, without dramatic contractions seen in
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Archigregarinida, Veloxidium, and squirmids. Etymol: ortho Gk straight, correct, true, as includes
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the standard originally described gregarines (Gregarina Dufour, 1828).
Vermigregarida ord. n. Diagnosis: Trophozoites aseptate, with densely packed, deep epicytic
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folds; mucron often elongate, cylindrical and hairy, sometimes squat; typically in gut lumen of polychaete hosts; szygy lateral or end-to-end; gamonts without mucron; gametes, sporozoites,
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oocysts or merogony not described; inhabit polychaete guts or rarely foraminifera. Etymol: vermis L. worm emphasises that known hosts are generally polychaete worms.
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Trichotokaridae fam. n. Diagnosis: Polychaete-infecting gregarines, commonly with elongate hairy mucron. Type genus Trichotokara (Rueckert and Leander 2010). Other genus Paralecudina Rueckert et al., 2013.
Polyplicariidae fam. n. Diagnosis: Polychaete-infecting gregarines, with blunt non-hairy mucron, and high density of longitudinal epicytic folds (4-5/m). Type and sole genus Polyplicarium (Wakeman and Leander 2013a).
Arthrogregarida ord. n. Diagnosis: ancestrally ‘septate’ gregarines living in gut lumen of arthropod hosts, plus derived parasites of other body cavities or tissues in insects and oligochaete annnelids, and some simplified aseptate forms. Mucron ranges from short to elongated as an epimerite. Ancestrally without merogony but some morphologically reduced genera secondarily
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24 evolved it. Szygy usually late and caudofrontal, sometimes frontal. Male gamont often uniciliate, never biciliate. Etymol: arthro Gk joint as all hosts are jointed/segmented animals; also refers to the trophonts’ joint-like appearance in the ‘septate’ majority. Porosporina subord. n. Diagnosis: ‘septate’ or aseptate lumenal parasites of crustacean (rarely
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mollusc) intestines; syzgy early, usually caudofrontal, sometimes involving more than two cells; merogony absent. Etymol: Named after type genus Porospora Schneider, 1875 of the oldest
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included family. Contrary to Rueckert et al. (2011a) the sole superfamily cannot be called
Cephaloidophoroidea if it includes Porosporidae, because superfamily names must be based on the
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oldest included family and attributed to the authors of that family not the person(s) raising it to that
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rank (article 23.3 of the International Code of Zoological Nomenclature, applicable to Sporozoa). Terragregarina subord. n. Diagnosis: ancestrally ‘septate’ insect, arachnid or oligochaete parasites
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plus some secondarily simplified non-sepate forms; merogony ancestrally and commonly absent but present in some insect parasites (former neogregarines). Etymol: terra L. earth, emphasizes that
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hosts are terrestrial (or as nymphs in freshwater) not marine. Superfamily Actinocephaloidea Léger, 1892 stat. n. Diagnosis: szygy frontal, late; insect,
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arachnid, myriapod hosts. Phylogenetically defined as the clade containing all gregarines more closely related to Monocystis, Ascogregarina, and Mattesia than to Stylocephalus or Gregarina.
Class Paragregarea
Paragregarea cl. n. Diagnosis: aseptate marine gut-dwelling or coelomic gregarines; annelid, nemertean, mollusc, deuterostome hosts; trophozoite with longitudinal or transverse folds or knoblike projections and surface crenulations; often with marked bending or thrashing motility; mucron short, pointed or blunt. Etymol: para Gk but, against, contrary to; emphasizes they are probably not directly related to orthogregarines. Velocida ord. n. Diagnosis: Aseptate. Trophont either contractile with bending and twisting motility, but neither gliding nor with longitudinal folds, or else non-contractile with longitudinal
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25 folds and gliding motility; mucron round or nipple-like. Etymol: velox, velocidis L. rapid because of rapid twisting of Veloxidium and gliding of Urosporoidea. Velocid- is the combining stem form that grammatically ought to have been used for Veloxidium. Urosporoidea Léger, 1892 superfam. n. Diagnosis: trophont aseptate, with longitudinal folds,
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knobs, and gliding motility; unlike Veloxidium, no bending or twisting motility; mucron round. Veloxidioidea superf. n. Diagnosis: as for sole family: Veloxidiidae fam. n. Diagnosis: trophont
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contractile with bending and twisting motility; lacking gliding motility or longitudinal folds. Type and sole genus Veloxidium Wakeman and Leander, 2012.
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Evolution and Taxonomy of Coccidiomorphea
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Stenophoridae Léger and Dubosq, 1904 (e.g. Stenophora).
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Stenophorida ord. n. Diagnosis: myriapod hosts; ‘septate’. Etymology: Named after sole family
Removal of Cryptosporidium makes Coccidiomorphea holophyletic on sequence trees.
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Recently it was shown that the sea-squirt symbionts Nephromyces (Saffo 1981) and Cardiosporidium (order Nephromycida (Cavalier-Smith 1993b)) belong in Sporozoa and are
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phylogenetically related to piroplasms (Saffo et al. 2010). Fig. 2 strongly confirms that but contradicts their non-CAT ML and Bayesian trees in showing coccidians as paraphyletic, with the nephromycid/piroplasm clade as sister to Eimeriida only not to all Coccidia. However, Saffo et al. (2010) did not include Haemosporida, so could not exclude an alternative relationship with them. Fig. 4 therefore included Haemosporida and other previously excluded coccidiomorphs to clarify their position. Nephromycids remain sister to piroplasms, with slightly reduced but still reasonable support and their joint clade remains sister to Eimerida only but with sharply reduced support. Haemosporida, though currently grouped with piroplasms as class Hematozoa, group with weak but significant support with adelinid coccidians not with piroplasms (Fig. 4). This makes it possible the grouping Haemosporida and piroplasms as Aconoidasida (Mehlhorn et al. 1980) or Hematozoa (Vivier 1982) was artificial and shared features are convergent. On present evidence retaining a
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26 separate class uniting piroplasms only with Haemosporidia is unjustified. Multigene trees strongly group piroplasms as sister to Eimeriida, but as transcriptomes are unavailable for Adeliida do not reveal whether they are nested within Coccidia or their sisters. Until they are it may be premature to abolish Hematozoa, retained here as a subclass of Coccidiomorphea to which Nephromycida is added (Table 1); the original rationale for Aconoidasida embracing piroplasms and Haemosporidia
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(Mehlhorn et al. 1980) however is defunct, as some ookinetes in Haemosporidia have conoids
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(Paterson and Desser 1989). A new superorder is needed for piroplasms plus nephromycids:
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Aconoidia superord. n. Intracellular parasites of ascidian haemocytes with posterior biciliate phase
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or non-ciliate parasites of mammalian blood cells with microtubule-supported projections from gametes; conoid absent. Etymol: a- without + conoid; Aconoidasida is a misleading name for
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Hematozoa as Haemosporidia often have conoids (Paterson and Desser 1989), never seen in piroplasms and nephromycids (Ciancio et al. 2008).
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Aggregata, initially considered a gregarine, has very long branches on 18S rDNA trees so was excluded from both coccidians and gregarines and left incertae sedis in Apicomplexa by Adl et
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al. (2012). However, its elongate triciliate microgametes are typical for Coccidia, and Fig. 4 strongly supports its grouping with Hepatozoon within Adeleida, as found by Castellanos-Martínez et al. (2013), so it clearly belongs in that order. Eimeria and Goussia are both found in two highly divergent clades that are genetically more different from each other than is either from Sarcocystidae, partly because rDNA has evolved much faster in one of these clades (Figs 2, 4). Clearly both genera have been overlumped and inaccurately split; Calyptospora forms one of three subclades within the longer branch Eimeriidae clade and probably does not deserve its own family. Important phylogenetically are Coelotrophiida (=Protococcidiida), e.g. the polychaete parasites Coelotropha and Grellia with early intracellular and later coelomic phases, for which I once established a separate class Coelotrophea (Cavalier-Smith 1993a). Sequence trees are needed to test their unity and phylogenetic position: a preliminary report of an ML tree suggests that they
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27 may be nearer to apicomonads than to Sporozoa (Diakin et al. 2012), but sequences are unavailable and could not be included here. However, their biciliate gametes do not resemble apicomonads (Bardele 1966; Porchet-Henneré 1967), and are distinct from those of Coccidia and Haemosporida in not being elongated with strongly reflected backward pointing centrioles; unlike Coccidia the centrioles are well separated and not closely attached behind a pointed
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microtubule-supported perforatorium. Moreover, as their complete conoid is typically
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coccidian-like with two intraconoid microtubules (Bardele 1966; Porchet-Henneré 1971),
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from Coccidia and Hematozoa merits a new third subclass:
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Table 1 conservatively keeps them in Sporozoa and Coccidiomorphea. Microgamete distinctness
Coelotrophia subcl. n. Diagnosis: Gametogony and sporogony extracellular, unlike other
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coccidiomorphs. Biciliate microgametes near-spherical, not elongated as in Coccidia and Haemosporida, not aciliate as in piroplasms; mitochondrion lies between anteriorly indented
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nucleus and the centrioles, not laterally as in Coccidia and Haemosporida. Microtubule-supported microgamete perforatorium absent, unlike Coccidia. Etymol: Named after sole included order
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Coelotrophiida.
Changing views of gregarine and sporozoan evolution The large and predominantly extracellular gregarines were known long before the small-celled intracellular coccidia. Soon after Leuckart (1879) united both groups as Sporozoa, Bütschli (1880-1882) suggested that Sporozoa evolved from flagellates by losing cilia as an adaptation to parasitism. Sporozoa are united primarily by having a non-flagellate cell invasion stage, the sporozoite, with radially symmetric apical cytoskeleton adapted for penetrating the host plasma membrane. Bütschli thought they might have evolved from euglenoids, now clearly not so. More presciently, the discovery of the coccidian-like syndiniid intranuclear parasite Coccidinium by Chatton and Biecheler (1934) led Chatton to suggest that evolution of multiple fission by an
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28 intracellular parasitic dinoflagellate could have led to the origin of Coccidia and Sporozoa generally. Levine favoured perkinsids as the closest flagellates to Sporozoa, and recent thinking has been dominated by his view (Levine, 1985) that gregarines are ancestral to coccidians (e.g. Leander 2008; Perkins et al. 2002 dated 2000; Schrével and Desportes 2013; Vivier and Desportes
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1990) even though I am unaware of any compelling explicit arguments for it. Grassé drew an analogy between bodonids and coccidian male gametes, but was inclined to think similarities
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convergent and regarded gregarines as two clades diverging from a common sporozoan ancestor and did not postulate that one evolved from the other. Vivier and Desportes (1990) wrote
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‘gregarines, which are exclusively parasitic on invertebrates, surely evolved earlier than other
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apicomplexan classes, which parasitise vertebrates.’ However that argument is fallacious because it confuses secondary invasion of vertebrate hosts with the primary origin of each group; coccidia and
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Hematozoa both parasitize invertebrates too so could be just as old as gregarines or even older. Anyway, if we now accept Cryptosporidium as a gregarinomorph then one gregarinomorph lineage
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did invade vertebrate hosts. Gregarines were still called ‘the early Apicomplexa’ by Shrével and Desportes (2013) even though their schematic tree showed gregarines, coccidia, and
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Cryptosporidium branching from the same point. The true flagellate ancestors of Sporozoa became clearer when other French protistologists, Brugerolle and Mignot (1979), showed that a myzocytotic predatory flagellate then called Spiromonas perforans (transferred by Patterson and Zölffel (1991) to Colpodella, then by me to Chilovora (Cavalier-Smith and Chao 2004)) had ultrastructure suggesting an intermediate position between dinoflagellates and Sporozoa, and was most unlike the bodonid Dimastigella described under the name Spiromonas angusta (MacDonald and Darbyshire 1977). Discovery of another myzocytotic predator (Foissner and Foissner 1984) led Krylov and Mylnikov (1986) to make a subclass Spiromonadomorphina and order Spiromonadida within Sporozoa for these free-living predators and Cavalier-Smith (1991) to place Colpodella within Apicomplexa and formulate the concept of alveolates as ancestrally photosynthetic eukaryotes with cortical alveoli and tubular
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29 mitochondrial cristae, and formally establish infrakingdom Alveolata to embrace ciliates, dinoflagellates, Apicomplexa, and protalveolates (Colponema). Earlier the haploid dinoflagellates and Sporozoa were formally grouped as Miozoa (Cavalier-Smith 1987), now a phylum with subphyla Protalveolata (sole class Colponemea) and Myzozoa (Cavalier-Smith 2013a,b).
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Spiromonadida was renamed Colpodellida by Cavalier-Smith (1993) (as Spiromonas is confusing and probably invalid) and the class Apicomonadea outside Sporozoa (but within Apicomplexa) was
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established for colpodellids and perkinsids, it being proposed that Sporozoa and dinoflagellates evolved independently from such myzocytotic apicomonads. When the first sequence tree to
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include a colpodellid grouped it, but not Perkinsus, with Sporozoa (Cavalier-Smith 2000) I
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explicitly proposed free-living Colpodella-like myzocytotic predatory flagellates as the ancestors of Sporozoa. After more colpodellid and perkinsid sequences became available Perkinsus was
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removed from apicomonads when it become clearer that Perkinsea was the deepest branch within Dinozoa and Apicomonadea the deepest within Apicomplexa (Cavalier-Smith and Chao 2004).
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Now sequences of both rDNA genes for Colponema place it as a sister clade to Myzozoa (Janouškovec et al. 2013) in accord with that classification; I mistrust their concatenated 6-gene
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tree that put it weakly as sister to all other alveolates, as the individual trees for the four proteins (the same used in the accompanying paper: Cavalier-Smith 2014) were confused and none of the six genes individually placed it so distantly.
The first explicit suggestions of how sporozoa evolved from an apicomonad emphasized that most gregarines are not truly extracellular but ambicellular (Cavalier-Smith 2004b; CavalierSmith and Chao 2004). I argued that the conoid, composed of conically arranged spiral tubulin fibres arranged like a compressed spring (Hu et al. 2002), was an innovation for cell penetration, and that the extracellular condition of bigger gregarines was developmentally and evolutionarily secondary. At least in Toxoplasma conoidal tubulin fibres are not microtubules but sharply curved protein sheets, comma-like in transverse section (Hu et al. 2002). Now that my CAT trees make it likely that orthogregarines and paragregarines evolved independently from early non-gregarine
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30 Sporozoa we can see that they constitute two somewhat distinct organismal types. Trophozoites of orthogregarines may start off as intracellular parasites but those inhabiting intestines spend most of their growth phase as ambicellular. From their broad distribution on the trees such ambicellular gut parasites are clearly the ancestral state for orthogregarines, so this lifestyle imposed the initial
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selective forces on their origin, outlined previously (Cavalier-Smith 2004b; Cavalier-Smith and Chao 2004). Orthogregarines secondarily invaded other habitats, notably the coelom of oligochaetes
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and then lost ambicellularity and became secondarily purely extracellular trophozoites. By contrast archigregarines are not ambicellular but always initially intracellular and in some but not all cases
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become truly extracellular by extrusion into the gut lumen before growth finishes. Some Selenidium
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even undergo multiple fission within the intestinal epithelium.
This basic difference in lifestyle between archigregarines and orthogregarines is consistent
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with their having evolved large trophozoites independently. The fact that only orthogregarines are truly ambicellular explains why only they have evolved true epimerites that anchor the partially
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embedded cell tip into the epithelium and are autotomised when growth finishes, making them truly extracellular only when differentiating into gamonts. The constricted condition (traditionally
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misleadingly called ‘septate’; they have a constriction not a septum) is also restricted to ambicellular arthropod-infecting orthogregarines and Stenophora, the constriction being arguably an adaptation to facilitate later autotomy and deutomerite release. The facts that ‘septate’ orthogregarines do not all cluster together on Fig. 3 supports Grassé’s (1953) judgement that septation has only weak systematic value; yet even though he rejected order Septata (Lankester 1885), Levine (1885) resurrected it as suborder Septatorina), whence it has lingered on as suborder Septatorina in the latest revisions of both of The Illustrated Guide to Protozoa (Clopton 2002 dated 2000) and more surprisingly of Grassé’s own Treatise chapter (Schrével and Desportes, 2013). It is now evident that Septatorina are polyphyletic, most comprising the ancestrally ambicellular orthogregarines that inhabit arthropod guts (a paraphyletic subgroup of septates) here placed in the new order Arthrogregarinida that also includes non-septate gregarines that clearly
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31 evolved from them (e.g. Monocystis) by switching life style to extracellularity. Stenophorids apparently evolved septation independently through convergent adoption of the ambicellular lifestyle in arthropod (millipede) guts (though protein sequence data are sorely needed to check they are really sisters to core paragregarines; relationship instead with septate Gregarinomorphea is not
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strongly excluded); however unlike ‘septate’ orthogregarines their mucron is retractible (Grassé 1953) and thus not a true caducous epimerite. Aseptatorina Chakravarty, 1960 (= Acephalina von
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Kölliker, 1848; Haplocyta Lankester, 1885) by contrast is a polyphyletic mixture of archigregarines and velocids which are never ‘septate’ and derived orthogregarines which lost the constriction
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secondarily.
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Grassé (1953) argued that the division of gregarines into schizogregarines with multiple fission unrelated to sex (merogony) and eugregarines without merogony by Léger (1900) was no
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less artificial; he thought that some gregarines had evolved merogony secondarily (his order Neogregarinida), whereas others had it primitively (his Archigregarinida). Sequence phylogeny
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shows he was right about the secondary nature of neogregarines, but that merogony evolved several times independently in different groups of orthogregarines (Fig. 3). Thus neogregarines are
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polyphyletic and the distinction evolutionarily too trivial to be used at ordinal level. Therefore the new phylogenetic classification (Table 1) abandons Neogregarinida, Eugregarinida, Aseptatorina, and Septata/Septatorina (all polyphyletic). Grassé’s ideas about archigregarines and merogony remain untested as there are no sequences for merogonic archigregarines (Merogregarinidae, Exoschizonidae), only for Selenidium of Selenidiidae, a family that Levine transferred from archigregarines to eugregarines (Levine 1971). Contrary to Levine, recent discussions of gregarine sequence phylogeny have all referred to Selenidium as an archigregarine (Rueckert et al. 2010; Rueckert and Leander 2009), followed here as I regard the presence or absence of merogony as evolutionary relatively trivial changes compared with cell morphology. Like Grassé (1953) but unlike recent Levine-influenced compilations (Perkins et al. 2002 dated 2000; Schrével and Desportes 2013; Vivier and Desportes 1990) I include Selenidiidae within archigregarines. It is
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32 noteworthy that Grassé (1953 p. 348) remarked that the most archaic gregarines are Selenidiidae, Urosporidae, and Lecudinidae – essentially Archigregarinida plus Urosporoidea (Table 1), i.e. all paragregarines then known. Sequence trees now confirm that he rightly singled out these groups as differing from other gregarines, as all three form a completely non-septate paragregarine clade, but
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suggest that he was wrong in assuming gregarine monophyly. Merogonic archigregarines might possibly be primitive for Paragregarea, but there is no reason to think they are ancestral to
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orthogregarines. The intracellular nature of merogonic multiple fission in Merogregarina reflects the fact that unlike the larger celled non-merogonic archigregarines trophozoites remain
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intracellular. Further growth prior to syngamy that forces older trophozoites of non-merogonic
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archigregarines to become extracellular is arguably a derived character for Sporozoa. However, that does not mean that non-merogonic archigregarines or orthogregarines
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evolved from merogonic ancestors by losing merogony. More likely both evolved divergently from non-merogonic ancestors independently of several lineages that independently evolved merogony.
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Cell biologically, multiple fission, whether preceding syngamy (gametogony), following it to produce sporozoites (sporogony) or in between periods of vegetative growth, is the same process.
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Multiple fission cell cycles are widespread in protists, especially in parasites and in free-living unicellular algae like Chlamydomonas (Craigie and Cavalier-Smith 1982) and must therefore be easy to evolve. They can evolve from normal binary fission cell cycles merely by imposing a reversible block to division, temporarily uncoupling normal size-dependent division control so cells become manyfold (not just two-fold) larger than daughter cells before dividing (Cavalier-Smith 1980). A round of multiple fission can be introduced at any point in a life cycle merely by relieving that block by some developmental signal – a simple switch, nothing complicated like evolving a conoid or myzocytosis. Clearly multiple fission had already evolved in the apicomonad ancestor prior to the origin of Sporozoa by evolving the conoid and non-ciliate sporozoite. Apicomonads grow manyfold whilst feeding and then the large cell encysts and undergoes multiple fission to make new flagellates; an adaptation to maximising food uptake and the number of daughters during
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33 a single feeding bout, as the rareness of food encounters makes it important to get as many offspring as possible from each. Commonly a division cyst yields four daughters. Almost nothing is known about apicomonad sex, so we cannot say whether their fission is gametogony, merogony or sporogony or more than one. Anteriorly connected quadriciliate cells of Algovora (=Colpodella) pugnax were suggested to represent syngamy (Simpson and Patterson
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1996), which may be correct if mitotic division occurs only within a cyst; it is less likely that it is a
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freak misdivided cell (common in flagellates) as one would expect division to begin anteriorly and sister hemicells to be posteriorly linked. However, this species could actually be a dinozoan not an
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apicomonad (Cavalier-Smith and Chao 2004), making its relevance to sporozoan origins unclear.
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Nonetheless it is likely that some apicomonads still are sexual; the one ancestral to sporozoa must have been and would probably have had both post-feeding and post-syngamy multiple fissions,
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probably into four daughters, and thus have two rather than three multiple fission stages in its life cycle. Almost certainly it would have been isogamous and copulating freely in water not within a
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cyst; thus anisogamy and gamont szygy were sporozoan innovations. I suggest that both evolved in the ancestral sporozoan. Contrary to what is sometimes stated, enclosure of gamonts within a cyst is
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not a unique gregarine feature but also occurs in some adelinid coccidians, even though the cyst is tenuous. Therefore it was probably lost by Eimeriida and Hematozoa.
Ciliary evolution, cell and genome size, and the polyphyletic origins of gregarines Sequence trees strongly indicate that gregarines are polyphyletic, not paraphyletic as commonly assumed since Levine (1985). Three arguments from cell evolution also make it improbable that gregarines are ancestral to coccidiomorphs. First consider ciliary evolution. Myzozoan cilia are distinctly different from the ancestral alveolate state represented by Colponema with normal triplet centrioles and two 9 + 2 cilia with equal sized central pair microtubules. In virtually all Myzozoa, unlike any other eukaryotes, one central pair microtubule is narrower than the other; though most clearly obvious and only explicitly previously noticed in Perkinsea
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34 (Leander and Hoppenrath 2008) it is also true of dinoflagellates, apicomomonads and Coccidiomorphea. A few images suggest equal sized central microtubules (e.g. Speer and Danforth 1976 Fig. 7) suggesting an occasional reversal to the ancestral state, presumably by losing or modifying some special protein(s) needed for making the narrow microtubule. With the
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notable exception of Aggregata, which has normal triplets (Heller 1970), Apicomplexa have generally reduced their ciliated centrioles largely to doublets (true of apicomonads and Sporozoa:
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Brugerolle and Mignot 1979; Colley 1967); only at the extreme base of the Chromera centriole can one clearly see triplets (Oborník et al. 2011). Sporozoa further modified their mitotic
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centrioles only by blocking centriolar development to yield short nine-singlet procentrioles capable
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of further duplication when ciliary development was suppressed during the evolutionary origin of sporozoites with complete conoids. Orthogregarines apparently lost the centre pair of their cilia and
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have 9 + 0, 6 + 0, or 3 +0 cilia, with only slightly reduced 8-doublet centrioles (Schrével and Besse 1975; Schrével and Desportes 2013). The 9 + 2 cilia of Coccidiomorphea could not have evolved
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from them. Gregarines have either no cilia or a single posterior cilium only in male gametes, so the
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ancestors of both orthogregarines and paragregarines lost the posterior cilium and its associated centriolar roots. Thus neither gregarine group can be ancestral to Coccidiomorphea, most of which have two cilia; Piroplasmida lost cilia altogether, but their nephromycid sisters have two posterior 9 + 2 cilia and apparently a two-microtubule anterior ciliary root and posterior directed multiplemicrotubule root as well as a dorsal microtubule fan in their microgametes (Cianco et al. 2008). Eimeriid and adelinid coccidian microgametes have two posterior-pointing cilia arranged as in apicomonads with an amorphous connector between them, but their cell apex is modified; nonetheless the dense apical perforatorium is some species clearly includes a short anterior fan of microtubules. Some have a third cilium, probably a multiply derived state. Several Eimeria and the adelinid Aggregata have a posterior microtubular root emanating between the two primary centrioles, but this root has several times been misinterpreted as a third or fourth recurrent
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35 intracellular ciliary axoneme (e.g. Scholtyseck et al. 1972). Eimeria perforans unusually has a third cilium (Scholtyseck 1965); Colley (1967) thought Eimeria nieschulzii also has a third intermediate ciliary stub, which may be correct even though some of the structures he labeled basal bodies are probably the amorphous centriolar connector not a centriole; however the structures labeled third flagellum might instead represent the posterior root of five singlet microtubules plus
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dense fibrillar material found there in Eimeria truncata (Gajadhar and Stockdale 1986) or 8-10
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microtubules in Eimeria magna (Speer and Danforth 1976) or four microtubules in Sarcocystis (Vetterling et al. 1973). These singlet microtubules cannot be relics of a hypothetical third cilium
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as Vetterling and others assumed. They are associated with dense material and probably are
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homologous with the most prominent right posterior root of apicomonads (R1) whose two centrioles are connected and oriented in the same way, and therefore provide evidence that Coccidia
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evolved from ancestors having two connected centrioles and a prominent posterior microtubular root. Haemosporida have either two posterior cilia (Haemoproteus: Bradbury and Trager 1968)
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with highly attenuated/modified centrioles or one (Plasmodium) with no trace of microtubular roots. It seems clear that the ancestral coccidiomorph had two cilia later reduced in one lineage to one, in
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another to zero, and increased in two others to three. In addition to a centriolar connector it would have had a slender anterior ciliary root (arguably R3), perhaps of 2 or a fan of about 8 microtubules, and a posterior root with several microtubules. Therefore Coccidiomorphea could not have evolved from either group of gregarines, which have a single cilium, which as primary differentiations after a non-ciliate stage would be developmentally homologous with the anterior cilium of biciliate eukaryotes even though they may point and swim posteriorly (see Cavalier-Smith 2013a for this argument) and should therefore lack homologues of the posterior roots associated with the older cilium. According to this argument, Coccidiomorphea ancestrally retained ciliary and centriolar transformation and features specific to the posterior cilium and its roots, but gregarines probably lost such features independently in Orthogregarinia and Paragregarea. Unfortunately ciliary
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36 ultrastructure is unknown for Paragregarea, most of which lack cilia, so we cannot say whether it is more like that of Orthogregarinia or Coccidiomorphea. A second argument relates to cell and genome size, a key difference between gregarines and Coccidiomorphea. As previously emphasized, coccidian cells are much smaller than the ancestral
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apicomonads and have correspondingly miniaturized their genomes, whereas most gregarines are far larger than their host cells and have greatly magnified their cells, nuclei and genome sizes
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compared with apicomonads or coocidia (Cavalier-Smith 1985 p. 123); thus their chromosomes and genomes are generally thousands of times more massive than those of Coccidia as first emphasized
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by Goldschmidt (1955) which led him to doubt that DNA can be the genetic material, because he
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did not envisage that in eukaryotes it actually has both genic functions and separate structural functions that make the amount of non-genic DNA highly dependent on cell size (Cavalier-Smith
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1978, 1982, 1985, 2005). These correlations reflect the universal eukaryotic scaling laws between cell, nuclear, and genome size, and have been explained by the theory of optimal cytonuclear ratio
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coupled with the skeletal function of all nuclear DNA whether it is genic or not (Cavalier-Smith 2005). The immense size of gregarines was favoured by their gut luminal life style, which offers an
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essentially unlimited food source allowing much more protracted cell growth than is possible for a predator of single cells, whereas the tiny size of coccidiomorphs is required for rapid intracellular reproduction within a similarly spatially restricted food source. The large-celled ambicellular or maturely extracellular habit would lead to an inexorable increase in cell size and in associated genome size through thousands of independent duplications of non-genic nucleoskeletal DNA. I argue that this would sooner rather than later become irreversible, because gregarine cells would become too large to shift lifestyle to purely intracellular location; large reductions in genome size would be impossible in a single step, as that would require thousands of concerted deletions, so gregarines would be trapped in the largely extracellular trophic condition and could not secondarily become minute intracellular parasites like coccidia; eventually they would pass a point where they could not even fit into single animal cells.
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37 The ciliary evolution and cell and genome size arguments both reinforce Grassé’s view that gregarines and coccidians are independent lines. It is historically incorrect to attribute to him the recent dominant Levinean view that coccidia evolved from gregarines. It is now clear that both coccidia and gregarines in the classical sense are polyphyletic. That is hardly surprising because the
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small cell intracellular versus large cell extracellular dichotomy is very simple, involving transitions crossed many times during protist evolution, so one should not expect organisms sharing these
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features to be phylogenetically coherent unless they share unique morphological features, which is not the case within Sporozoa. Relying too much on them led to the small-celled intracellular
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Cryptosporidium being wrongly classified with coccidians rather than orthogregarines. Interestingly
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unlike Coccidia, but like Piroplasmida and some but not all Haemosporida, it has lost the conoid (Matsubayashi et al. 2008). Loss of the apicoplast in the common ancestor of Cryptosporidium
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and orthogregarines is a third reason why this gregarinomorph subclade cannot have been ancestral
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to Coccidiomorphea, many of which have apicoplasts.
Multiple fission in apicomplexan evolution
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The third traditional dichotomy between coccidia and gregarines lies in gametogony. In all Sporozoa gametes are anisogamous, unlike virtually all other protists except a few volvocalean green algae, whose life cycle involves multicellular coenobia so they are really multicellular, not true unicellular protists. In gregarines male and female gamonts divide a similar number of times, whereas in classical coccidia female gamonts do not divide. This dichotomy also has a simple adaptive explanation that allows it to evolve easily more than once. I argued above that szygy evolved in the ancestor of all sporozoa and was independently secondarily lost by Eimeriida and Hematozoa. The enclosure of gamonts by a cyst (unique in protists) inevitably led to anisogamy with respect to ciliary development because as syngamy occurred within a cyst it would be wasteful for both gametes to develop cilia and better for one to focus its resources on food storage for zygote development; only one needed to make cilia to find the other. The presence of the cyst however
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38 radically changed selective forces relating to cell size anisogamy (see Bulmer and Parker 2002; Parker 1978). In that conventional argument a major driver of cell miniaturization to make sperm is the benefits of fertilizing ever more genetically different eggs. But by being locked up within the cyst wall with access to only one female genotype this driver for evolving many more divisions into
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ever smaller sperm is absent in gregarines, so there is no benefit of having more multiple fissions than the female. Because of that constraint gregarines kept the ancestral condition of an equal
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number of fissions in both sexes. Those Coccidiomorphea that lost the zygy cyst probably did so because, unlike in the large gut where there can be many sexual partners, a strictly intracellular life
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style (specially in small host cells) greatly reduces the chances of finding a genetically different
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partner in that cell, so multiple divisions have to precede partner association (making merogony almost a sine-qua non for an intracellular parasite) and at least one partner must be much more
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mobile and seek the other across tissues.
Moreover, as soon as szygy was lost for that reason, male gametes suddenly were for the
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first time exposed to the selective force favouring numerous tiny sperm, and thus evolved more divisions than the female; conversely this tipping of the balance imposed a new force on the female
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to reduce the number of divisions eventually to zero to maximise egg size (Bulmer and Parker 2002; Parker 1978). Intracellularity of Cryptosporidium exposed it to the same selective forces as in most coccidiomorphs, so it independently evolved female gamonts that do not divide before fertilization. As apicomonads probably have equal biciliate gametes little different from vegetative cells, both the gregarine and the coocidia-like, gametogony patterns are derived compared with the ancestral condition. But both almost certainly diverged independently polyphyletically from a less sexually dimorphic ancestral sporozoan, which was probably an intracellular parasite like coccidia rather than a purely extracellular one. An intracellular ancestry is more likely because in both orthogregarines and Paragregarea development starts intracellularly and because, as I explain in a separate paper, full penetration into host cells prior to intracellular development provides a marked contrast to the extracellular nature of apicomonads and thus a novel selective force that favoured the
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39 origin of symmetrical conoids, two preconoidal rings, and microtubule nucleating polar ring from the asymmetric pseudoconoid and pellicles of apicomonads. Thus the contrasting patterns of sexual dimorphism in Sporozoa are also consistent with gregarine polyphyly, as they are simply explicable
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as multiple, mutationally and developmentally easy, responses to similar selective forces.
Derived gliding motility
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It is notable that actin-myosin based gliding motility of most orthogregarines is similar to the coccidiomorph cell entry machinery (Valigurová et al. 2013); coccidiomorphs use actin-based
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gliding past the host/parasite cell junction ring to enter host cells. I suggest that this gliding
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machinery evolved in a common ancestor of Gregarinomorphea and Coccidiomorphea, possibly but not necessarily after it diverged from Paragregarea. The absence of translational gliding and
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presence of pendular or threshing movements in archigregaines and Veloxidium is a marked difference consistent with an early divergence of Paragregarea from a putative
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gregarinomorph/coccidiomorph clade, as suggested by the CAT tree of Fig. 2. Also suggestive of such a clade is the fact that their conoids have two intraconoid microtubules, never more. Though
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evidence concerning paragregarine intraconoid microtubules is scrappy and inconclusive, Lankesteria has at least one (Garnham et al. 1971 Fig. 3; possibly two if the two densities between the two right hand rhoptries on the transversely sectioned conoid in Fig. 1 are microtubules). One image for the archigregarine Selenidium hollandei (Schrével 1971 Fig. 24) suggests that there may be three, like the anterior rostral root of Chilovora perforans (Brugerolle and Mignot 1979). If that difference were to be confirmed it could be explained as early divergence from an apicomonad ciliary root incorporated into the conoid lumen when it closed (Cavalier-Smith unpublished). Gliding motility probably evolved independently in orthogregarines and Urosporoidea. Though orthogregarines and Paragregarea most likely evolved independently from the earliest ancestral sporozoans, non-gliding Squirmidea are probably not even Sporozoa. Squirmids probably lack not only conoids (which can be secondarily lost in both coccidiomorphs and
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40 gregarinomorphs) but even a symmetrical apical complex found even in all secondarily aconoidal sporozoans – even Cryptosporidium which has largely lost pellicular microtubules retains a radially symmetrical and extra thick polar ring; Cryptosporidium also retained the two gregarinomorph intraconoidal microtubules with an historical central asymmetry, as in all sporozoan apical
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complexes. Even Plasmodium, where some species have no conoid or intraconoid microtubules (despite their radially symmetric apical protrusion being called the conoid region or sometimes
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carelessly even conoid (Ladda et al. 2001)), retains two preconoidal rings (Hepler et al. 1966) - at least one species kept an asymmetrical dense fibre connecting its degenerate apical complex to the
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centriole, which is likely to have descended from an apicomonad anterior root, an idea reinforced
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by the discovery of the highly conserved corticate anterior striated root protein SF-assemblin in the
Conclusions
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probably homologous centriolar/conoidal connector of Toxoplasma (Francia et al. 2012).
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1. The improved CAT-GTR-GAMMA phylogenies provide a firmer myzozoan tree onto which cellular and host-interaction differences across Sporozoa map in a much more
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evolutionarily comprehensible way than before. 2. Gregarines are polyphyletic. Orthogregarines are most closely related to Cryptosporidium and then rhytidocystids (here placed in three separate subclasses of Gregarinomorphea) than to Paragregarinea (Archigregarinida, and new orders Velocida and Stenophorida). Platyproteum and Filipodium are probably not gregarines (i.e. neither orthogregarines nor paragregarines) nor Sporozoa and placed in a new class Squirmidea. 3. The three former gregarine groups represent similar convergent adaptation towards terminally extracellular giant trophic cells within intestines, but show significant differences in detail reflecting their probably independent origins from broadly similar myzocytotic ancestors. Environmental DNA BOLA566 is probably from a stenophorid paragregarine.
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41 4. Many improvements to sporozoan classification have been made, especially at highest ranks and within orthogregarines. Most notably, Eugregarinida have been split into four orders (orthogregarine Vermigregarida and Arthrogregarida; paragregarine Velocida and Stenophorida) and Neogregarinida merged into Arthrogregarinida.
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5. I explained the key selective forces that led to the differentiation of sporozoan cell structure and life-histories in an often convergent manner that was previously taxonomically
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confusing.
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Appendix A. Supplementary data
Supplementary data associated with this article can be found, in the online version, at
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http://dx.doi.org/...
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Valigurová, A., Hofmannová, L., Koudela, B., Vávra, J., 2007. An ultrastructural comparison of the attachment sites between Gregarina steini and Cryptosporidium muris. J. Eukaryot. Microbiol. 54, 495-510. Valigurová, A., Vaškovicová, N., Musilová, N., Schrével, J., 2013. The enigma of eugregarine epicytic folds: where gliding motility originates? Front. Zool. 10, 57.
Page 51 of 60
52 Vetterling, J.M., Pacheco, N.D., Fayer, R., 1973. Fine structure of gametogony and oocyst formation in Sarcocystis sp. in cell culture. J. Protozool. 20, 613-621. Vivier, E., 1982. Reflexions et suggestions à propos de la systématique des Sporozoaires: création d'une classe des Hematozoa. Protistologica 18, 449-453.
ip t
Vivier, E., Desportes, I., 1990. Phylum Apicomplexa, in: Margulis, L., Corliss, J.O., Melkonian, M., Chapman, D.J. (Eds.), Handbook of Protoctista. Jones & Bartlett, Boston, pp. 549-573.
cr
Wakeman, K.C., Leander, B.S., 2013a. Identity of environmental DNA sequences using
from capitellid polychaetes. Mar. Biodiv. 43, 133-147.
us
descriptions of four novel marine gregarine parasites, Polyplicarium n. gen. (Apicomplexa),
an
Wakeman, K.C., Leander, B.S., 2013b. Molecular phylogeny of marine gregarine parasites (Apicomplexa) from tube-forming polychaetes (Sabellariidae, Cirratulidae and Serpulidae),
M
including descriptions of two new species of Selenidium. J. Eukaryot. Microbiol. 60, 514-525. Yabuki, A., Toyofuku, T., Takishita, K., 2014. Lateral transfer of eukaryotic ribosomal RNA
doi:10.1038/ismej.2013.252
d
genes: an emerging concern for molecular ecology of microbial eukaryotes. ISME J. 1-4
Ac ce pt e
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65
Page 52 of 60
53 Figure legends
Fig. 1. GTR gamma maximum likelihood tree for 18S rDNA of 652 eukaryotes using 1540 nucleotide positions, showing gregarine branches collapsed on Fig. 1. Numbers on branches
ip t
show bootstrap support (400 resamplings) for almost all bipartitions (those ≥ 85% in bold); black blobs indicate 100% support. To compress the tree to a single page some speciose clades
cr
are collapsed, the number of taxa in each being indicated to the right of the clade name.
Percolozoa are shown non-collapsed and Dinozoa and Euglenozoa less collapsed in Fig 2.
us
Fig. 2. PhyloBayes CAT-GTR-GAMMA tree for 18S rDNA of 137 Myzozoa only using 1614
an
nucleotide positions, excluding long-branch gregarines. Support values for this consensus tree from two well converged chains (maxdiff 0.128) are posterior probabilities; black blobs indicate
M
1.0 PP. Exclusion of longer-branch gregarines coupled with the evolutionarily more realistic site-heterogeneous CAT model makes it even clearer than in Fig. 1 that Squirmidea formerly
d
treated as gregarines do not branch within Sporozoa but with Dinozoa and that archigregarines are less closely related to orthogregarines than is Cryptosporidium. The arrow marks the
Ac ce pt e
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65
putative time of loss of the apicoplast in a common ancestor of Cryptosporidium and orthogregarines.
Fig. 3. PhyloBayes CAT-GTR-GAMMA tree for 18S rDNA of 196 alveolates only using 1577 well-aligned nucleotide positions, including long-branch gregarines. Support values are posterior probabilities; black blobs indicate 1.0 PP. Sum of two well-converged chains (maxdiff 0.101316). Though an ML tree was also run for this alignment, for clarity its support values are not shown, but some key ones are mentioned in the text. Fig. 4. PhyloBayes CAT-GTR-GAMMA tree for 18S rDNA of 276 alveolates only using 1577 well-aligned nucleotide positions. Support values are posterior probabilities; black blobs indicate 1.0 PP. This tree is for one of two chains that did not converge (maxdiff 0.6801); the other chain differed in only two respects as discussed in the text.
Page 53 of 60
54 Table 1. Revised classification of parvphylum Sporozoa Leuckart, 1879 and its 4 classes and 13 orders Class 1. Paragregarea cl. n. (mainly marine; annelid, nemertean, myriapod, mollusc, deuterostome hosts) (three orders, two new) Order 1. Archigregarinida* Grassé, 1953 (largely marine and aseptate; longitudinal ridges; contractile, non-gliding)
Levine, 1971) (Selenidioides, Meroselenidium, Merogregarina)
ip t
Family 1. Merogregarinidae Porter, 1908 (invalid junior synonym Selenidioididae
Family 3. Exoschizonidae Levine, 1971 (Exoschizon)
us
Order 2. Velocida ord. n. (aseptate; motility varied)
cr
Family 2. Selenidiidae* Brasil, 1907 (e.g. Selenidium*, Selenocystis)
Superfamily 1. Veloxidioidea superfam. n. (contractile, non-gliding; no longitudinal folds)
an
Family Veloxidiidae fam. n. (Veloxidium)
Superfamily 2. Urosporoidea Léger, 1892 superfam. n. (formerly eugregarines;
M
gliding, longitudinal folds)
Family 1. Lecudinidae Kamm, 1922 (e.g. Lecudina, Lankesteria, Difficilina) Family 2. Urosporidae Léger, 1892 (e.g. Urospora, Pterospora, Lithocystis)
d
Order 3. Stenophorida ord. n. (millipede hosts; ‘septate’) Family Stenophoridae Léger and Dubosq, 1904 (e.g. Stenophora)
Ac ce pt e
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65
Class 2. Gregarinomorphea Grassé, 1953 em. (four orders; three new) Subclass 1. Histogregaria subcl. n.
Order Histogregarida nom. n. pro Agamococcidiorida: Levine, 1979 = Agamococcidiida Levine, 1980.
Family 1. Rhytidocystidae Levine, 1979 Family 2. Gemmocystidae Upton and Peters, 1986
Subclass 2. Cryptogregaria subcl. n. Order Cryptogregarida ord. n.
Family Cryptosporidiidae Léger, 1911 Subclass 3. Orthogregarinia subcl. n. Order 1. Vermigregarida ord. n. (mainly polychaete gregarines) Family 1. Trichotokaridae fam. n. (Trichotokara, Paralecudina) Family 2. Polyplicariidae fam. n. (Polyplicarium) Order 2. Arthrogregarida ord. n. (mainly arthropod hosts; often ‘septate’) Page 54 of 60
55 Suborder 1. Porosporina subord. n. (crustacean, seldom mollusc, hosts) Superfamily Porosporoidea Labbé, 1899 (not Porosporicae Chakravarty, 1960; syn. Cephaloidophoroidea Simdyanov & Aleoshin in Rueckert et al., 2011) Family 1. Porosporidae Labbé, 1899 (Porospora, Nematopsis, Pachysporospora, Thiriotia) Family 2. Cephaloidophoridae Kamm, 1922 (e.g. Cephaloidophora)
ip t
Family 3. Uradiophoridae Grassé, 1953 (e.g. Heliophora, Uradiophora) Family 4. Ganymedidae Huxley, 1910 (Ganymedes)
cr
Suborder 2. Terragregarina subord. n. (terrestrial arthropod or oligochaete hosts; cilia without central pair microtubules)
us
Superfamily 1. Stylocephaloidea Léger, 1892 (rank given by Clopton, 2009) Family 1. Stylocephalidae Léger, 1892 (e.g. Stylocephalus, Xiphocephalus, Colepismatophila) has one cilium szygy frontal, late; beetle, Diplura, Thysanura hosts.
an
Superfamily 2. Actinocephaloidea Léger, 1892 superfam.. n. (szygy frontal, late; insect, arachnid, myriapod hosts)
M
Family 1. Actinocephalidae* Léger, 1892 (e.g. Geneiorhynchus, Prismatospora, Hoplorhynchus, Pyxinia)
Paraschneideria)
d
Family 2. Monocystidae Bütschli, 1882 (Monocystis, Ascogregarina,
Family 3. Ophryocystidae Léger and Dubosq, 1908 (Ophryocystis, Apicystis)
Ac ce pt e
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65
Family 4. Syncystidae Schneider, 1886 (Syncystis, Steinina) Family 5. Lipotrophidae Grassé, 1953 (e.g. Mattesia) Family 6. Sphaerocystidae Chakravarty, 1960 (Sphaerinina)
Superfamily 3. Gregarinoidea Labbé, 1899 (not Chakravarty, 1960) szygy early Family 1. Gregarinidae* Labbé, 1899 (e.g. Gregarina, Leidyana, Amoebogregarina) Family 2. Blabericolidae Clopton, 2009 (e.g. Blabericola, Protomagalhaensia)
Terragregarina incertae sedis***: Dactylophoridae Léger, 1892 (centipede hosts)
Class 3. Coccidiomorphea Doflein, 1901 (five orders) em. Subclass 1. Coccidia Leuckart, 1879 (3 orders: Adeleida Léger, 1911 (with szygy) including families Adeleidae and Aggregatidae; Eimeriida Léger, 1911 (Eimeriidae, Calyptosporidae, Sarcocystidae, Elleipsisomatidae); Ixorheida Levine, 1984 (no gamogony). Subclass 2. Hematozoa Vivier, 1982 (=Aconoidasida Mehlhorn et al., 1980**) Superorder Haemosporidia Danilewsky, 1885; order Haemosporida Danielwsky, 1885
Page 55 of 60
56 Superorder Aconoidia superord. n. orders Nephromycida Cavalier-Smith, 1993 (ascidian symbionts: Nephromyces, Cardiosporidium); Piroplasmida Wenyon, 1926) Subclass 3. Coelotrophia subcl. n. Order Coelotrophiida Vivier, 1982 (=Protococcidiida Kheisin, 1956) (5 families, e.g. Coelotropha)
Sporozoa incertae sedis: Order Blastogregarinida Chatton and Villeneuve, 1936 (no folds)
ip t
Family Siedleckiidae Chatton and Villeneuve, 1936 (Siedleckia) For all taxa I use the uniform zoological endings recommended by Pearse (1936) for phyla (-
cr
a) classes (-ea), subclasses (-ia) orders (-ida), and suborders (-ina), which have become
standard for Protozoa and phagotrophic or parasitic non-algal chromists, not the longer
us
replacement forms suggested by Levine (1959) for all zoological nomenclature but not adopted by anyone else except anomalously for Sporozoa. I suggest that Levine’s idiosyncratic respellings of older names (e.g. Piroplasmorida for Piroplasmida) be now abandoned for
an
Sporozoa also. Families listed only for gregarines. * paraphyletic short 2-microtubule conoid.
M
** name unsuitable as conoid present on one haemosporidan ookinete and apparently has
d
***Many other mainly insect-host, probably terragregarine, families are not listed because no sequences are available and it is therefore unclear to which superfamily they belong.
Ac ce pt e
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65
Page 56 of 60
Figure 1
6
87
10
20 Apicomonadea 8 93 Coccidea/Hematozoa 7 4 87 Cryptosporidium Rhytidocystis cyamus GQ149767
17 33
Sporozoa
9
31
49
22 23 42
C3_E012 AY046842 C2_E026 AY046816 BOLA212 AF372767 BOLA458 AF372771 environmental clone BOLA327 AF372770 Paralecudina polymorpha AY196706
40
31 2
26
3
85
BAQA65 AF372825
Blabericola migrator FJ459754
97 Blabericola haasi FJ459753 Protomagalhaensia granulosae FJ459757 Protomagalhaensia wolfi FJ459758 Blabericolidae 99 99 Gregarina basiconstrictonea FJ459740
Gregarinoidea
1
85 37
4
24
67
Veloxidium leptosynaptae JN857966
Filipodium phascolosomae FJ832163 Platyproteum vivax AY196708
63 Dinozoa 36
Chilodonella uncinata
2
0
Diphylleia rotans AF420478 triciliatum ContigKST Diphylleida 16 RigifilaCollodictyon ramosa AB686266 Rigifilida Micronuclearia podoventralis anoxic marine sediment DNA CCA32 AY179990 NOT dinoflagellate Telonema subtilis RCC3587 Telonema antarcticum AJ564773 Telonemea 24 54 Ministeria vibrans AF271998 63 29 Capsaspora owczarzaki 59 Monosiga ovata AF271999 Monosiga brevicollis 80 28 Salpingoeca infusionum Amoebidium parasiticum Y19155 29 Ichthyophonus hoferi 43 Corallochytrium limacisporum Fonticula alba FJ816018 72 36 RT5iin16 AY082991 Cristidiscoidea RT5iin14 AY082985 42 Nuclearia simplex 4 Rozellidea 52 36 Aphelidida Rozellida 4 35 47 60 25 FUNGI 13 85 Manchomonas bermudensis Apusomonas proboscidea 40 microbial mat cold seep DNA RM1-SGM31 AB505488 62 30 marine marine mIirobial mat cold seep DNA RM2-SGM40 AB505548 marine microbial mat cold seep DNA RM2-SGM42 AB505550 33 Thecamonas trahens EU542595 12 AT4-11 AF530526 Thecamonas sp. CCAP 9795 contaminant 28 marine microbial mat cold seep DNA RM2-SGM44 AB505552 12 25 90 marine microbial mat cold seep DNA RM1-SGM33 AB505490 95 marine microbial mat cold seep DNA RM1-SGM37 AB505494 marine microbial mat cold seep DNA DSGM-75 AB275075 marine microbial mat cold seep DNA RM1-SGM35 AB505492 marine microbial mat cold seep DNA RM2-SGM45 AB505553 lake sediment NAMAKO-2 DNA AB252742 90 RM2-SGM71 AB505579 RM1-SGM28 AB505485 95
Varisulca Sulcozoa
kingdom ANIMALIA 7
Ac ce pt e 0
10
2
2
21
3
95
0 7 12
53
28
77
34
38
91
79
26
99
77 49
D3P05C09 EF100280 BOLA366 AF372746
70
D4P07D12 EF100407 SA2_2C8 EF527166 BOLA187 AF372745
Subulatomonas tetraspora HQ342676
RM1-SGM29 AB505486 NAMAKO-1 AB252741 RM2-SGM39 AB505547
7
Choanozoa podiates Apusomonadida
Amoebozoa 12 clade
Breviatea
Breviata anathema WIM30 AM114802 Mantamonas plastica Mantamonadida Mantamonas sp.2 Africa 74 Planomonadida 5 Glaucophyta 3
91
6
19
86
picobiliphyte 3 Cryptista Viridiplantae 12
77
28
cr
Hacrobia p. p. (kingdom CHROMISTA)
submarine fumarole anoxic sediment DNA TAGIRI-28 AB1914 V. costatus contaminant EF483923 Microhelida Microheliella maris AF534711
d
3
Dinozoa
M
0
Urosporoidea Velocida
Ciliophora
AF300282
Heliozoa 7
96
'core paragregarines'
53 Heterokonta + Rhizaria 48
Haptophyta 5 Palpitomonas bilix AB508339 Palpitia
16
Veloxidiidae Squirmidea
Mesodinium/Myrionecta 12
Blepharisma americanum BJA16SLRNA Loxodes striatus AM946031 Remanella sp. R101 AM409181 Trachelocerca ditis GQ167153 Tracheloraphis sp. TCIRGSSA AF149979 Paramecium tetraurelia Halteria grandinella AF194410 Pseudourostyla cristata DQ019318
AT4-68 AF530543
19
us
chelyosomae EU670240 92Lankesteria 67 Lankesteria ascidiae JX187607 78 CCI7 AY179977 70 Lecudina tuzetae AF457128 70 53 Lankesteria abbotti DQ093796 99 Pterospora schizosoma DQ093793 Pterospora floridiensis DQ093794 Urosporidae Lithocystis sp. DQ093795 96 99 environmental clone DSGM-6 AB275006
94 26 33
Archigregarinida
an
20
(CHROMISTA)
Vermigregarida
environmental clone SS1E0145 EU050982 environmental clone NAMAKO-25 AB252765 Blabericola cubensis FJ459751
Selenidium idanthyrsae JN857967
37
Trichotokaridae
Selenidium cf. mesnili JN857968 Selenidiidae 86 AY196709 Selenidium serpulae DQ683562 Selenidium terebellae 27 Selenidium pisinnus FJ832162 Selenidium orientale FJ832161 Difficilina paranemertis FJ832159 Difficilina tubulani FJ832160 Lecudina longissima FJ832157 Lecudina phyllochaetopteri FJ832156 Lankesteria cystodytae EU670241 Lecudinidae
52 23
Trichotokara eunicae@JX426618 Trichotokara nothriae GU592817 Trichotokara japonica JX426617
Gregarina cuneata FJ459744 LEMD145 AF372805 LEMD003 AF372797 Gregarina cloptoni FJ459742 Gregarina niphandrodes FJ459747 55 gregarines insect gregarines Gregarina polymorpha FJ459748 81 Gregarina diabrotica FJ459745 80 Gregarina coronata FJ459743 Gregarinidae 19 34 Leidyana erratica FJ459752 Gregarina blattarum FJ459741 42 Gregarina chortiocetes NA18SRRB 99 Gregarina caledia GNA18SRRA Gregarina tropica FJ459749 Amoebogregarina nigra FJ459737 49 Gregarina kingi FJ459746 Stenophorida 64 Uncultured DNA AB275069 98 diplopod gregarinesStenophora robusta FJ459760 Thiriotia pugettiae HQ876006 Porosporidae Ganymedes themistos FJ976721 Ganymedidae Heliospora caprellae HQ876007 Uradiophoridae BOLA48 AF372821 Porosporoidea crustacean gregarines 97 DH148-EKD18 AF290084 85 CS_R003 AY046643 Cepahaloidophoridae 98 Cephaloidophora cf. communis HQ876008 63 Selenidium boccardellae JN857969
28
1
Harosa
81
22
4
Alveolata
short-branch gregarinesTerragregarina pro parte
DSGM_8 AB275008
3
Myzozoa
'core orthogregarines' BS 79%
Rhytidocystidae 49 Stylocepahaloidea + Actinocephaloidea49 parasite of Ammonia beccarii ABU07937 unidentified clade BOLA267 AF372774 clone CCI31 AY179976 clone CCA38 AY179975 Polyplicariidae
Tridacna hemolymph parasite AB000912
ip t
Apicomplexa
Sulcozoa
Glissodiscea
kingdom PLANTAE
clade
83 Rhodophyta 4
Anaeromonadea 10
Metamonada
Malawimonas jakobiformis Malawimonadea Malawimonas californiana Andalucia godoyi AND19 Andalucia godoyi AND28 Andalucia incarcerata Jakobea Guyamas hydrothermal marine sediment DNA CS_E022 AY046649 5 Histionina saltmarsh pool water DNA CCW8 AY180011 Tsukubamonas globosa AB576851
77 Euglenoida/ Postgaardea
Excavata Percolozoa
Eozoa 45
121 Kinetoplastea/Diplonemea Euglenozoa
0.4
Page 57 of 60
Figure 2
Tridacna haemolymph parasite AB000912 Rhytidocystis polygordiae DQ2739887 Rhytidocystis cyamus GQ149767
Cryptosporidium fragilis
Stylocephalus giganteus FJ459761 Colepismatophila watsonae FJ459738
0.38 0.75 0.56
Eimeria leucisci
DNA D5P09F10 EF100363 DNA D5P09C08 EF100358 Steinina (=Gregarina) ctenocephali GU320208 Monocystis agilis AF457127 Ophryocystis electroscirrha FJ865354
Actinocephaloidea
Goussia szekelyi Goussia koertingi Goussia pannonica Goussia janae Goussia desseri Goussia kessleri Goussia chalupski Goussia sp. Goussia balatonica Calyptospora spinosa Calyptospora serrasalmi Calyptospora funduli
ip t
Gregarinomorphea
Eimeriida
Frenkelia glareoli Hyaloklossia lieberkuhni Toxoplasma gondii Besnoitia oryctofelis Nephromyces sp. MO-3F HM469391 Nephromyces sp. MO-5F HM469379 Nephromyces sp. MO-4G HM469380 Nephromyces sp. MO-6H HM469378 Nephromyces sp. MO-4B HM469377 .75 Nephromyces sp. MO-6H HM469376 Nephromyces sp. MO-4F HM469375 Nephromyces sp. MR-1N HM469383 Nephromyces sp. MM-3N HM469382 Nephromyces sp. MC-2N HM469384 intrahemocytic apicomplexan from Halocynthia EF558768 Cardiosporidium cionae Babesia microti Babesia conradae Cytoauxzoon felis Theileria parva Babesia odocoilei Babesia gibsoni Babesia orientalis Babesia bovis Klossia helicina Hepatozoon catesbianae Hepatozoon canis Adelina grylli 0.99 Adelina bambarooniae Vitrella brassicaformis DQ174731 environmental DNA LT85_AS KC487674 environmental DNA LT85_K21 KC487845 environmental DNA LT35_H13 KC486567 environmental DNA LT35_ET3 KC486520 0.98 environmental DNA KC487652 environmental DNA GU067926 0.99 Colpodella (= Alphamonas) edax AY234843 BOLA914 AF372772 BOLA553 AF372785 .87
0.54 0.36 0.98
.93
0.98
Coccidia p. p.
Coccidiomorphea
Nephromycida Aconoidia
M
0.82
cr
0.99 Eimeria variabilis Eimeria anguillae 0.84 Goussia neglecta Eimeria arnyi 0.69 Eimeria myoxi Eimeria mitis 0.65 0.95 Cyclospora papionis Cyclospora colobi Cyclospora cayetanensis 0.93 0.99 Sarcocystis oviformis Sarcocystis sinensis Sarcocystis hirsuta 0.96 Sarcocystis gallotiae
d
0.99 0.95 0.98 0.54
Ac ce pt e
Adeleida
Piroplasmida
Sporozoa
Coccidia p. p.
Vitrella/Colpodella subclade
Apicomonadea
Chromerida Voromonas pontica AY078092
0.48
Chromera velia DQ174731
0.44 0.64
'Colpodella' sp. HEP GQ4111073 'Colpodella' sp. ATCC5059 AY142075 'Colpodella' tetrahymenae AF330214 BOLA176 AF372786 Psammosa pacifica JN873311 Psammosa atlantica JN873310 .54 Kofoidinium sp. FG256 GU355681 Syndinium sp. ex Corycaeus DO146406 .12 Hematodinium sp. MF-2000 AF286023 Amoebophrya sp. ex Prorocentrum micans AY208893 0.99 Amoebophrya sp. ex Ceratium tripos AY208892 Amoebophrya sp. ex Karlodinium 0.53 Amoebophrya sp. .57.58 0.69 Amoebophrya sp. AF069516 Heterocapsa triquetra GU594638 .47 Cochliodinium polykrikoides Karenia brevis Abedinium dasypus .21 Alexandrium tamarense AB088333 .41 Prorocentrum gracile .27 Polarella glacialis Haplozoon praxillellae .98 Haplozoon axiothellae
0.99
0.85
0.97
0.83 0.97 0.99
0.29
PROSOPE.E3_95m.109 EU793257 marine DNA 65 DQ916410 uncultured Dubosquella 3905 JF791022 Ichthyodinium chabelardi FJ440625 Kofoidinium cf. pavillardii GU355680 Spatulodinium pseudonoctiluca GU355685
Noctiluca scintillans AF022200
Dinozoa
Syndinea p. p. Dinoflagellata Oxyrrhida 0.99
Syndinea p. p.
Oxyrrhis marina AF280077 Oxyrrhis marina AB033717 Oxyrrhis marina AF482425
Dubosquellidae
Noctilucea Thalassomyces fagei AY340590
IN242 NA2_2D10 EF526760 Parvilucifera procentrum FJ424512 Parvilucifera sinerae EU502912 Parvilucifera infectans AF133909 Perkinsus atlanticus AF509333 marine hydrothermal vent DNA AT4-98 AF530536
0.47 .49 0.99 0.38 0.47 0.32
Apicomplexa
Voromonadida
Peridinea = Dinophyceae
.18 .12
Orthogregarinia
Paraschneideria metamorphosa DQ462456 Ascogregarina culicis Pseudomonocystis lepidota PNT18SRRNA Mattesia geminata AY334568 Apicystis bombi FN546182
0.86 0.94
0.65
0.45
Histogregaria
Cryptogregarida Cryptosporidium serpentis Cryptogregaria baileyi 0.970.95 Cryptosporidium Xiphocephalus ellisi FJ459762 Xiphocephalus triplogemmatus FJ459763 Stylocephaloidea
0.99
0.27
Rhytidocystidae
Cryptosporidium parvum
0.91
0.99
Stenophorida
Stenophora robusta FJ459760
us
0.92
Paragregarea
Archigregarinida
Selenidium terebellae AY196709 Selenidium pisinnus FJ832162 Selenidium orientale FJ832161 BOLA566 AF372760
an
0.51 0.89 0.69 0.92
0.94 0.3
Ellobiopsida Ellobiopsis chattonii FJ593706
Perkinsea Squirmidea
Perkinsozoa Platyproteum vivax AY196708 Filipodium phascolosomae
Page 58 of 60
Figure 3 Colpodella (=Alphamonas) edax AY234843
Vitrella/Colpodella clade Voromonadidida Voromonas pontica AY078092 Chromera/Voromonas clade Chromerida Chromera velia DQ174731 Platyproteum vivax AY196708
environmental DNA GU0679
'Colpodella'-like 3
Squirmidea 0.99 Rhytidocystidae Histogregaria 0.98
Filipodium phascolosomae FJ832163
Cryptosporidium 4
Cryptosporidiidae
Tridacna haemolymph parasite AB000912 Rhytidocystis polygordiae DQ2739887
Rhytidocystis cyamus GQ149767
0.99
0.5
clone CCI31 AY179976 clone CCA38 AY179975
0.48
0.23
Polyplicariidae
parasite of Ammonia beccarii U07937
BOLA267 AF372774
0.58
0.62
0.67 0.68
0.51 0.19
0.34
0.99 0.86
0.91
0.99
environmental clone BOLA327 AF372770
Stylocephalidae ES
Blabericola haasi FJ459753
Blabericolidae
Amoebogregarina nigra FJ459737 Leidyana erratica FJ459752
0.71
Gregarinidae
Actinocephalidae ES
Pyxinia crystalligera FJ459759 cooling tower water DNA CTW-5c JF774861
stained glass window DNA clone 1195 DQ451601
0.99
Hoplorhynchus acanthatholius FJ459750
Prismatospora evansi FJ459756 Monocystidae Monocystis agilis AF457127 saltmarsh sediment DNA alveolate clone CCA5 AY179988
strain ATCC 50646 GQ377652
EA
Actinocephalidae ES
us
0.97
Gregarinoidea
Gregarina tropica FJ459749
DNA D5P09F10 EF100363 DNA D5P09C08 EF100358
0.29 0.47 0.4
Stylocephaloidea
Protomagalhaensia wolfi FJ459758 Protomagalhaensia granulosae FJ459757 Blabericola migrator FJ459754
Geneiorhynchus manifestus J459739
0.95
0.45
Porosporoidea 8 crustacean gregarines
Blabericola cubensis FJ459751 Gregarina blattarum FJ459741 0.99 Gregarina chortiocetes NA18SRRB Gregarina caledia GNA18SRRA Gregarina kingi FJ459746 0.85 Gregarina diabrotica FJ459745 0.88 Gregarina coronata FJ459743 Gregarina polymorpha FJ459748 Gregarina niphandrodes FJ459747 0.32 0.4 Gregarina cloptoni FJ459742 LEMD145 AF372805 LEMD003 AF372797 Gregarina cuneata FJ459744 0.99 0.86 0.89 0.43 Gregarina basiconstrictonea FJ459740
0.33
13 soil DNA clones (Lesaulnier et al. 2008) wrongly annotated as Eimeriidae + Lake AF37277 & EU910605 or Cryptosporidiidae Syncystidae N Syncystis mirabilis DQ176427 Steinina ctenocephali GU320208 Actinocephalidae ES 0.98 0.99 Psychodiella chagasi FJ865354 0.85 Ophryocystis electroscirrha FJ865354 Ophryocystidae N 'Lecudinidae' Paraschneideria metamorphosa DQ462456 Sphaerocystidae EA 0.91 0.98 Ascogregarina 4 0.99 0.83 Pseudomonocystis lepidota PNT18SRRNA E Mattesia geminata AY334568 0.8 Lipotrophidae N 0.99 Apicystis bombi FN546182 Stenophora robusta FJ459760 BOLA566 AF372760 0.45 0.99 Selenidium idanthyrsae JN857967 0.23 0.99 Selenidium serpulae DQ683562
0.99
an
gregarine env. DNACHI_S2_24 AY821921
0.14
0.84 Selenidium cf. mesnili JN857968 Selenidium boccardellae JN857969 Selenidium pisinnus FJ832162 Selenidium orientale FJ832161 Selenidium terebellae AY196709 Veloxidium leptosynaptae JN857966 0.67 environmental clone DSGM-6 AB275006 0.59 Difficilina tubulani FJ832160 Difficilina paranemertis FJ832159 Pterospora schizosoma DQ093793 0.88 Pterospora floridiensis DQ093794 Lithocystis sp. DQ093795 Lecudina longissima FJ832157 0.98 Lecudina phyllochaetopteri FJ832156 0.27 Lankesteria cystodytae EU670241 0.78 Lankesteria chelyosomae EU670240 0.99 Lankesteria ascidiae JX187607 Lankesteria abbotti DQ093796 0.99 Coccidiomorphea 7 Lecudina tuzetae AF457128 0.99 0.86 CCI7 AY179977
Selenidiidae
0.77
Kofoidinium sp. FG256 GU355681
AF022200 0.7 Noctiluca scintillans Thalassomyces fagei AY340590 0.45 0.24 Ellobiopsis chattonii FJ593706 Ellobiopsida 0.99 Oxyrrhida 0.99 Psammosa pacifica JN873311 Oxyrrhis marina AF482425 Psammosa atlantica JN873310 0.36 0.11 Amoebophryidae 5 0.84 Dubosquellidae 4 0.96 0.48 Syndinea Syndinium sp. ex Corycaeus DO146406 Hematodinium sp. MF-2000 AF286023 0.63 0.82 0.97 Peridinea 9
0.96 0.6 0.99
Stenophorida
Archigregarinida
Urosporidae
Lecudinidae
Urosporoidea
Velocida Paragregarea
Dinozoa
Syndiniidae
0.99 Mesodinium pupula N412739
Mesodinium pulex QD-I DQ845294 Mesodinium pulex DQ411865 Mesodinium pulex JN412740
CCW75 AY180032 0.99 Myrionecta rubra AB364286 Mesodinium rubrum JN412736 rubrum JN412738 0.87 Mesodinium Ciliophora Myrionecta rubra AY587129 Mesodinium rubrum CCMP2563 JN412738 0.89 AF290065 AY180041 0.88 DH145-EKD11 Mesodinium chamaeleon JN084213 0.99 CCW100 AY180041
Karyorelictea 4 other Intramacronucleata 4
Blepharisma americanum BJA16SLRNA
0.98
Terragregarina
Noctilucea
Ac ce pt e
0.99 Spatulodinium pseudonoctiluca GU355685
Actinocephaloidea 0.97
Lecudinidae
d
Perkinsea 7
BAQA65 AF372825
A r t h r o g r e g a r i d a
Veloxidiidae Veloxidioidea
M
0.23
Kofoidinium cf. pavillardii GU355680
Trichotokara nothriae GU592817 Trichotokara eunicae@JX426618 Trichotokara japonica JX426617
Trichotokaridae Vermigregarida
BOLA458_AF372771 BOLA212 AF372767
Stylocephalus giganteus FJ459761 Colepismatophila watsonae FJ459738 0.99
0.75 0.77
0.46
0.35
environmental cloneNAMAKO-25 AB252765 C3_E012 AY046842 C2_E026 AY046816 Paralecudina polymorpha AY196706
0.6
Xiphocephalus triplogemmatus FJ459763 Xiphocephalus ellisi FJ459762
0.51
0.43
unidentified clade
environmental clone SS1E0145 EU050982
0.99
0.5 0.68
Orthogregarinia
DSGM_8 AB275008
0.76
0.1
0.44
Apicomonadea
Vitrella brassicaformis DQ174731
ip t
0.99
cr
0.51 0.75 0.3
0.4
Page 59 of 60
Figure 4 0.46 Voromonas pontica AY078092 Colpodella-like 3 0.99 Voromonadida 0.76 Chromera velia DQ174731 Chromerida Apicomonadea 0.21 Vitrella brassicaformis DQ174731 0.57 environmental DNA GU0679 Vitrella/Colpodella clade (=Alphamonas) edax AY234843 0.21 Colpodella 0.99 Platyproteum vivax AY196708 Filipodium phascolosomae FJ832163 0.77 0.95 Cryptosporidium 4 Cryptosporidia Squirmidea Gregarinomorphea Rhytidocystidae Agamococcidia 0.15 0.99 Polyplicariidae + unidentified subclade 0.57 DSGM_8 AB275008 Porosporoidea 8 crustacean gregarines 0.1 Paralecudina clade .62 0.89 0.65 Stylocephaloidea 4 0.1
Apicomplexa
Geneiorhynchus manifestus J459739
.93 0.41 DNA D5P09F10 EF100363
GQ377652 0.99 stained glass window DNA clone 1195 DQ451601 cooling tower water DNA CTW-5c JF774861
0.97 0.83 0.27 0.5
other Actinocephaloidea
Selenidium idanthyrsae JN857967 Selenidium serpulae DQ683562 Selenidium cf. mesnili JN857968 Selenidium boccardellae JN857969
0.76
BOLA566 AF372760
0.35
.83
Velocida
cr Eimeriidae
Eimeriida
0.7 Calyptospora spinosa FJ904637 Calyptospora serrasalmi FJ904641 0.79 Calyptospora funduli GU479670 Sarcocystis 6 0.65
Frenkelia glareoli AF009245
Sarcocystidae
Nephromyces sp. MO-6H HM469378
.99 Nephromyces sp. MO-4B HM469377 .99.99.99 Nephromyces sp. MO-6C HM469376
Halocynthia intrahemocytic apicomplexan EF558768
d
0.62 .99 Cardiosporidium cionae EU052685
.94
Coccidiomorphea
Nephromycida
Nephromyces sp. MO-4F HM469375 Nephromyces sp. MR-1N HM469383 Nephromyces sp. MM-3N HM469382 .96 Nephromyces sp. MC2-2N HM469384
.99 .61
0.73
Gregarinoidea
(Gregarinomorphea)
lieberkuehni AF298623 .76 Hyaloklossia Toxoplasma gondii X75453 .65 Besnoitia oryctofelisi GU479632 .96 Nephromyces sp. MO-3F HM469381 .99 Nephromyces sp. MO-5H HM469379 .98 .99 Nephromyces sp. MO-4G HM469380
.99
Paragregarea
Stenophorida
Stenophora robusta FJ459760
Goussia szekelyi GU479656 Goussia koertingi GU479647 Goussia pannonica GU479642 Goussia janae GU479644 Eimeria leucisci GU479649 Goussia desseri GU479665 Goussia kessleri GU479645 0.9 Goussia chalupskyi GU479653 Goussia sp. BMR-2011d GU479643 0.89 Goussia balatonica GU479650 Eimeria variabilis GU479674 Eimeria anguillae GU479633 Goussia FJ009242 0.99 Eimeria neglecta arnyi AY613853 0.88 Eimeria myoxi JF304148 Cyclospora papionis AF111187 Cyclospora colobi AF111186 0.91 Eimeria mitis EMU40262 Cyclospora cayetanensis AF111183
0.07
Archigregarinida
an
0.41
0.73
0.4
Selenidium pisinnus FJ832162
orientale FJ832161 0.6 SelenidiumSelenidium terebellae AY196709
M
0.28 0.21 0.99 0.82 0.97
0.97
.86
0.94
Actinocephaloidea
Hoplorhynchus acanthatholius FJ459750
0.98
0.19
Pyxinia crystalligera FJ459759
strain ATCC 50646
DNA D5P09C08 EF100358
ip t
0.48
us
0.34
Piroplasmida 8
Plasmodium reichenowi Z25819 Plasmodium gallinaceum Plasmodium malariae PFARGBAB Plasmodium ovale PFASSUR 0.44 Plasmodium gonderi AB287270 0.93 0.75 Plasmodium fieldi AB287282 cynomolgi PFARRNASSA 0.86 .32Plasmodium Plasmodium vivax VU07367 0.86 Plasmodium inui EU400396 Plasmodium fragile asexually expressed PFARRSS 0.88 0.7 Plasmodium vivax El Salvador sporozoite PVU07368 0.8 Plasmodium knowlesi PKU83876 Plasmodium coatneyi EU400393 Plasmodium chabaudi DQ241815 Plasmodium berghei PFARGSS
Ac ce pt e
0.39
Hematozoa
Haemosporida
0.57 0.98
0.95
Hepatocystis 6
Adelina dimidiata DQ096835 Klossia helicina HQ224956 0.35 Hepatozoon canis DQ111754 Hepatozoon catesbianae AF130361 0.59 Aggregata octopiana KC188342 0.56 0.84 Aggregata octopiana DQ096837 Aggregata eberthi KC188343 0.5 Aggregata eberthi DQ096838 Adelina grylli DQ096836 .99 Adelina bambarooniae AF494059
0.71
.39
anoxic marine DNA EF526760
Perkinsida 5 0.84
Mid-Atlantic Ridge hydrothermal sediment marine DNA AF530534 0.69
Ellobiopsis chattonii FJ593706 Spatulodinium pseudonoctiluca GU355685
0.99 Noctiluca scintillans AF022200 0.37
Kofoidinium sp. FG256 GU355681
Noctilucida
Kofoidinium cf. pavillardii GU355680 .23 0.08 Amoebophryidae 5 0.82 Dubosquellidae 4 0.93 .55 Syndinium sp. ex Corycaeus DO146406 .97 0.53
Adeleida Aggregatidae Adeleidae
Perkinsea 7
Dinozoa
Oxyrrhis marina AF482425 Oxyrrhida
pacifica JN873311 0.56 0.99 Psammosa Psammosa Psammosa atlantica JN873310 Thalassomyces fagei AY340590 0.12
0.36 .7
Adeleidae
Hematodinium sp. MF-2000 AF286023
4
Syndiniidae
17
Coccidiomorphea
Trichotokara 3 Gregarinomorphea
Ellobiopsida
Oxyrrhea
Dinozoa
Syndinea
Peridinea 9
Ciliophora 0.5
Page 60 of 60
