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User:Snoteleks/Taxonomy

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  • DIAPHORETICKES Adl et al. 2012: phylogenetic definition.[1]
    • TSAR Strassert, Jamy, Mylnikov, Tikhonenkov & Burki 2019: phylogenetic definition;[2] tubular mitochondrial cristae.[3]
      • Telonemia Shalchian-Tabrizi 2006: phagotrophic biflagellate protists of pyriform shape; flagella emerging on opposite sides of a short protruding antapical rostrum; a food vacuole may be located anteriorly at the stout end of cell, in front of a centro-lateral large nucleus; food uptake in depression in antero-ventral region; characteristic band of vesicles located laterally; tubular mitochondrial cristae; characteristic subcortical lamina of microtubuli supporting layers of microfilaments oriented slightly offset from a right angle to each other;[4] tripartite mastigonemes (in Lateronema and Arpakorses); complex cytoskeleton not found in other eukaryotes.[3] Arpakorses Tikhonenkov & Karpov 2022,[3] Lateronema Cavalier-Smith 2015,[5] Telonema Griessmann 1913.
      • Sar Burki et al. 2008 emend. Adl et al. 2012 (=Harosa Cavalier-Smith 2010): phylogenetic definition;[1] presence of gene Rab1A.[6]
        • Stramenopiles Patterson 1989 emend. Adl et al. 2005: motile cells typically biciliate, typically with heterokont ciliation; anterior flagellum with complex tripartite mastigonemes in two opposite rows, composed of a hollow base, a tubular shaft and non-tubular distal fibres; smooth posterior flagellum; tubular mitochondrial cristae; typically four microtubular kinetosomal roots.[7][8][3]
        • Alveolata: layer of flattened vesicles (cortical alveoli) underneath the plasma membrane,[3] sometimes secondarily lost; with ciliary pit or micropore; tubular or ampulliform mitochondrial cristae.[8]
          • Colponemidia Tikhonenkov et al. 2014: biciliate; three-membraned alveolar pellicle; two microtubule bands armour longitudinal groove; micropore absent; nontubular mastigonemes present; cytotrophic predators and sometimes on microinvertebrates.[8]
          • Ciliophora
          • Myzozoa Cavalier-Smith & Chao 2004: myzocytotic; cytosolic chloroplasts (type II RuBisCo) or leucoplasts; epiplastid membrane separate from rough ER.[11]
            • Apicomplexa Levine 1980 emend. Cavalier-Smith 2013: centriolar doublets; conoid or paraconoid with two lumenal microtubules.[11]
              • Colpodellida Cavalier-Smith 1993 emend. Adl et al. 2005, 2019 (="chrompodellids" Janouškovec et al. 2015; Apicomonada > Apicomonadea Cavalier-Smith 1993 emend. 2018): phototrophs, or predatory heterotrophs; complex plastids, when present bound by four membranes; mitochondrion with tubular cristae; highly flattened cortical alveoli; microtubules beneath alveolae; conoid-like structure with apical complex and rostrum; biciliate; micropore present; cysts at least in some species;[8] myzocytotic biciliates, predominantly apically rostrate, with posterior pointing cilia and pseudoconoid (apically truncated cone of microtubules with dense arms, closed as complete or almost complete ring at apex, open at ventral lower side) and 0-5 eccentric lumenal microtubules, rarely simplified to non-microtubular paraconoid (Colpodella) or missing (Vitrella only), and micronemes; ciliary centre pair microtubules ancestrally with separate globular axosomes and surrounded by slender basal cylinder just distal to axosome pair; ciliary transition zone with proximal, not distal dense plate; multi-microtubule right anterior and posterior ciliary roots; no singlet microtubule between left and right posterior roots.[11]
              • Sporozoa Leukart 1879
            • Dinozoa Cavalier-Smith 1981 emend. 2018
              • Perkinsea > Perkinsida > Perkinsidae Levine 1978 emend. Adl et al. 2005 (=Perkinsozoa Norén & Moestrup 1999): trophozoites parasitic, dividing by successive binary fissions; released trophozoites (termed hypnospores) developing outside host to form zoospores via the formation of zoosporangia or morphologically undifferentiated mononucleate cells via a hypha-like tube; zoospores biciliate; apical organelles including an incomplete conoid (open along one side), rhoptries, micronemes and micropores, and a microtubular cytoskeleton with both an anterior and posterior polar ring.[8]
              • Dinoflagellata Bütschli 1885 emend. Fensome et al. 1993 emend. Adl et al. 2005: cells with two cilia in the motile stage—typically, one transverse cilium ribbon-like with multiple waves beating to the cell’s left and longitudinal cilium beating posteriorly with only one or few waves; nucleus typically a dinokaryon with chromosomes remaining condensed during interphase and lacking typical eukaryotic histones and centrioles; dinoflagellate/viral nucleoproteins package chromatin; closed dinomitosis with extranuclear spindle;[8] Phycodnavirus-like basic chromatin proteins.[11]
        • Rhizaria: fine pseudopodia (filopodia)[3] varying as simple, branching or anastomosing patterns, often supported by microtubules.[8]
          • Gymnosphaerida (?) Poche 1913 emend. Mikrjukov 2000: axopodial microtubules in irregular hexagonal prism; kinetocyst and other types of extrusomes along axopodia; tubular mitochondrial cristae; in some genera, cells attached to substrate with cytoplasmic stalk; free-swimming as amoeboid or motile biciliated cells; one or more nuclei, often located in the amoeboid base of stalk when present; complex life cycle unresolved.[8]
          • Cercozoa Cavalier-Smith 1998 emend. Adl et al. 2005 emend. Cavalier-Smith 2018: lacking distinctive morphological or behavioural characters; biciliated and/or amoeboid, usually with filopodia; most with tubular mitochondrial cristae; cysts common; kinetosomes connecting to nucleus with cytoskeleton; usually with microbodies and extrusomes.[8]
          • Endomyxa Cavalier-Smith 2002 emend. Bass & Berney in Adl et al. 2019: phylogenetic definition.[8]
          • Retaria Cavalier-Smith 2002: mainly marine heterotrophs with reticulopodia or axopodia; usually with various types of skeleton.[8]
    • Archaeplastida: chloroplasts derived from primary endosymbiosis with a cyanobacterium.
      • Glaucophyta Skuja 1954 (=Glaucoplantae B. Marin & Melkonian in Li et al. 2020): eukaryotic algae with cyanelles, i.e. blue-green plastids with chlorophyll a and phycobilins, stabilized by a peptidoglycan wall between the two envelope membranes; motile biflagellate or non-motile coccoid or colonial algae; with cytoplasmic starch; freshwater algae; in the nuclear-encoded 18S rRNA, base pair 3 of Helix 20 [H505] is A○G instead of C-G.[12] Putative genera: Archaeopsis, Peliaina, Strobilomonas, Chalarodora, Glaucocystopsis.[13]
      • Chloroplastida (=Viridiplantae): chlorophylls a and b.
        • Prasinodermophyta B. Marin & Melkonian 2020: non-motile, coccoid, marine green algae without organic scales, surrounded by a cell wall; occurring as free floating single cells or loose cell colonies, or organized as attached macroscopic palmelloid thalli of up to 25 cm diameter composed of distantly spaced cells embedded in a common gelatinous matrix. Cytokinesis unequal. Marine algae; in the nuclear-encoded SSU rRNA, the third from last nucleotide is G instead of U; in the nuclear-encoded LSU rRNA, base pair 11 of Helix G4 [H2093] is UxC instead of U-A.[12]
          • Prasinodermophyceae B. Marin & Melkonian in Li et al. 2020: marine, coccoid green algae, single-celled, or forming loose planktonic colonies; cells small, ca. 2-6 μm, surrounded by a thick multi-layered cell wall, without pores; the terminal base pair of Helix 30 [H861] in the chloroplast-encoded SSU rRNA is U-A instead of C-G. Prasinoderma Hasegawa & Chihara in Hasegawa et al. 1996.[12]
          • Palmophyllophyceae Leliaert et al. 2016 emend. B. Marin & Melkonian in Li et al. 2020: cells coccoid, (sub)spherical, ca. 3-14 μm, surrounded by a thin cell wall perforated by pores; occurring as free floating single cells, or forming attached macroscopic palmelloid thalli of up to 25 cm diameter composed of distantly spaced cells embedded in a common gelatinous matrix; in the chloroplast-encoded 23S rRNA, the terminal nucleotide of the spacer between Helices C1 and D1 [H533 and H579] is U instead of A.[12]
            • Prasinococcales Guillou et al. ex Leliaert et al. 2016 emend. B. Marin & Melkonian in Li et al. 2020: solitary, coccoid green algae, about 3-8 μm; cells surrounded by a thin cell wall perforated by pores, surrounded by a thick gelatinous capsule; planktonic marine algae; in the nuclear-encoded 18S rRNA, base pair 3 of Helix 50 [H1506] is C-G instead of U-A.[12]
            • Palmophyllales > Palmophyllaceae Zechman, Verbruggen, Leliaert, Ashworth, M.A.Buchheim, Fawley, H.Spalding, Pueschel, J.A.Buchheim, Verghese & Hanisak 2010: benthic marine algae; thallus green to deep green, of firm gelatinous texture; thallus macroscopic, crustose or erect; thallus composed of subspherical cells in gelatinous matrix; cells irregularly distributed throughout the whole gelatinous matrix but more frequent near surface; cell diameter 6–10 μm; cells with a single cup-shaped chloroplast without pyrenoids. Palmophyllum, Verdigellas.[14]
        • Chlorophyta
        • Streptophyta Bremer & Wanntorp 1981: asymmetric motile cells, when present, with pair of cilia without mastigonemes; basal bodies with distinctive multilayered structure of microtubular rootlet and cytoskeletal anchor; thylakoids stacked; open mitosis; usually with phycoplast,but some with phragmoplast and cell plate; with primary plasmodesmata between adjacent cells in filamentous forms; filaments branching or nonbranching; with non-motile vegetative phase; some with multinucleate cells; with or without sexual reproduction; sexual species with haplobiontic life cycle; with desiccation-resistant cysts (zygospores); glycolate oxidase in peroxisomes; Cu/Zn superoxide dismutase; ciliary peroxisome.
          • Phragmoplastophyta Lecointre & Guyander 2006: cell division by way of some form of phragmoplast; some oogamous, others anisogamous with nonmotile female gamete and motile male gamete.
            • Charophyceae Smith 1938 emend. Karol et al. 2009: thallus attached to substrate with rhizoids; thallus a central axis of multinucleate internodal cells, with whorls of branchlets radiating from uninucleate cells at nodes; calcium carbonate accumulates in cell wall of many species; haplobiontic life cycle; sexual reproduction oogamous with sperm cells; differentiated sperm and egg producing organs; antheridium with several shield cells and a manubrium that gives rise to spermatogenous filaments; primarily in freshwater.
            • Coleochaetophyceae Jeffrey 1982: thalli discs of cells or branched filaments; sheathed hairs as extensions of the cell wall.
            • Anydrophyta Rensing 2020: molecular features to cope with drought, i.e. land plant auxin repressors Aux/IAA, plant-like A/B type auxin response factor (ARF), ABA receptor, and GRAS gene family.[15]
              • Zygnematophyceae van den Hoek et al. 1995 emend. Hall et al. 2009: without ciliated stages; sexual reproduction via conjugation; thalli unicellular or filamentous; no centrioles.
              • Embryophyta Engler 1886 emend. Lewis & McCourt 2004 (=Plantae Haeckel 1866): ciliated basal bodies, when present, with distinctive multilayered structure of microtubules and cytoskeletal anchor; open mitosis with phragmoplast at cytokinesis; plasmodesmata and other characteristic cell–cell junctions; diplobiontic life cycle, with vegetative propagation possible in many; alternation of generations with fertilization of egg by sperm inside protective test; embryology with tissue differentiation coordinated by hormones; differentiated sperm and egg cells, may be on different sexual individuals, on different organs of the same individual or in the same organ.
                • Bryobionta: dominance of gametophyte; single sporangium in each sporophyte.
                • Polysporangiomorpha Kenrick & Crane 1997 (=Pan-Tracheophyta P.D. Cantino & M.J. Donoghue 2007): free-living sporophyte; multiple sporangia per sporophyte.
                  • “Protracheophytes” (†)
                  • Tracheophyta Sinnott 1935 ex Cavalier-Smith 1998 (=Tracheidatae Bremer 1985; =Apo-Tracheophyta P.D. Cantino & M.J. Donoghue 2007): walls of water-conducting cells with a thick, lignified, decay-resistant layer; sterome, a well developmental peripheral zone of the stem consisting of thick-walled, decay-resistant cells; pitlets in the tracheid wall; tracheids.
                    • Lycopodiophyta Cronquist, Takhtajan & Zimmermann 1966 emend. P.D. Cantino & M.J. Donoghue 2007 (=Microphyllophyta sensu Bold 1957; Lepidophyta sensu Smith 1955; Lycopodiopsida sensu Bierhorst 1971): association of a single axillary or adaxial sporangium with a sporophyll; absence of vasculature in the sporangium; metaxylem tracheids pitted; root stele bilaterally symmetrical, with phloem located on only one side of the stele; crescent-shaped root xylem; microphylls; exarch xylem differentiation in stem; stellate xylem strand in stem; reniform sporangia with transverse dehiscence.
                      • Lycopodiales DC. ex Bercht. & J. Presl 1820 (=Lycopodiaceae P. Beauv. ex Mirb., 1802; Huperziaceae Rothm. 1962; Lycopodiellaceae Val. N. Tikhom. 2018; Phylloglossaceae Kunze 1843; Urostachyaceae Rothm. 1944 nom. illeg.): isospory.
                      • Isoëtopsida Cronquist et al. 1972 emend. P.D. Cantino & M.J. Donoghue 2007 (=Glossopsida Bold 1957): heterospory; ligules.
                    • Euphyllophyta P.D. Cantino & M.J. Donoghue 2007: roots with monopodial branching and endogenous lateral roots; sporangia terminating lateral branches and dehiscing longitudinally; lobed, mesarch primary xylem strand, which has been modified in the stems of most extant members; multiflagellate spermatozoids (convergent with Isoëtes); a 30 kb inversion in the chloroplast genome.
                      • Monilophyta P.D. Cantino & M.J. Donoghue 2007: exclusively centrifugal development of the spore exine.
                      • Pan-Spermatophyta P.D. Cantino & M.J. Donoghue 2007 (=Radiatopses Kenrick & Ccrane 1997): tetrastichous branching; a distinctive form of protoxylem ontogeny with multiple strands occurring along the midplanes of the primary xylem ribs.
                        • Lignophyta M.J. Donoghue & J.A. Doyle 2007 (=Lignophytia Kenrick & Crane 1997): bifacial vascular cambium; cork cambium, producing periderm; cortical fiber strands; heterospory (except in Aneurophytales).
                          • Spermatophyta Britton & Brown 1896 emend. P.D. Cantino & M.J. Donoghue 2007: endarch primary xylem in the stem; meiospores and microgametophytes with distal aperture; linear tetrad of megaspores; platyspermic ovules; sealed micropyle; honeycomb alveolar pollen infratectal structure; heterospory; ovule, an integumented, indehiscent megasporangium that develops after fertilization into a seed; embryo dormancy; axillary branching; eustele; cataphylls.
                            • Acrogymnospermae P.D. Cantino & M.J. Donoghue 2007: phylogenetic definition; abaxial microsporangia.
                            • Angiospermae Lindley 1830 emend. P.D. Cantino & M.J. Donoghue 2007: closed carpel, which develops into a fruit; ovule with two integuments; cuticle of megasporangium thick; lack of cutinized megaspore membrane; highly reduced female gametophyte, usually with no more than eight nuclei; endosperm resulting from double fertilization; microgametophyte with three nuclei; scalariform pitting or perforations in secondary xylem; two or more orders of leaf venation; poles of stomatal guard cells level with aperture; axially aligned companion cells derived from the same mother cells as the sieve elements; pollen with unlaminated endexine; stamen with two pairs of pollen sacs.
  • AMORPHEA Adl et al. 2012: phylogenetic definition.[1]
    • Amoebozoa Lühe 1913, sensu Cavalier-Smith 1997:[17] almost all demonstrating amoeboid activity in stages of their life cycle; amoeboid locomotion through steady flow of the cytoplasm or occasional eruptions, or through the extension and retraction of pseudopodia and/or subpseudopodia with little coordinated movement of the cytoplasm; cells naked, often with well-developed differentiated glycocalyx; tectum or cuticle covering cells in some groups, other are testate; tubular mitochondrial cristae (ramicristate); frequent sporocarpic or sorocarpic fruiting; uniciliated, biciliated and multiciliated stages in some groups; reduction of bikont kinetid to unikont kinetid in some groups.[8]
      • Tubulinea Smirnov et al. 2005: lobose pseudopodia (lobopodia), variable cell projections, smooth in outline, with rounded tips, which participate in the relocation of the main cytoplasmic mass of the cell and include both the granuloplasm and the hyaloplasm; entire cell or pseudopodia are tubular, cylindrical or subcylindrical, with a rounded cross-section; monoaxial flow of the cytoplasm in every pseudopodium or entire cell; no evidence of ciliate stages; two groups testate, two groups sorocarpic; no sporocarpy reported.[8]
        • Corycida Kang et al. 2017: phylogenetic definition; cells covered with flexible, leather-like coating forming one or several openings used to protrude pseudopodia or are enclosed in hard test made of spicules with multiple apertures.[8]
        • Echinamoebida Cavalier-Smith 2004:
        • Elardia Kang et al. 2017: phylogenetic definition; cells naked or covered with a hard test; tubular or produce tubular pseudopodia; if flattened or branched, capable of altering the locomotive form to monopodial or polypodial, with tubular pseudopodia.[8]
          • Leptomyxida Pussard & Pons 1976 sensu Smirnov et al. 2017:[18] naked amoebae with locomotive form altering from a flattened expanded or reticulate one to a subcylindrical monopodial one when in rapid movement or under specific conditions; adhesive uroidal structures always present.[8]
          • Arcellinida Kent 1880: cell covered with hard or highly rigid organic or mineral extracellular test consisting of either self-secreted elements (calcareous, siliceous or chitinoid), a sheet-like chitinoid structure, or recycled organic or mineral particles bound together, with a single main opening.[8]
            • Organoconcha Lahr et al. 2019: organic, highly flexible and hemispheric test; aperture reaching almost the entire diameter of the whole test, often without rims; pseudopods blunt.[19]
            • Phryganellina Bovee 1985: organic test, sometimes with a calcite internal coating, or agglutinated test; terminal aperture in general; pseudopods conical; comprises both fast growing, small terrestrial species and slow-growing, large aquatic organisms.[19]
            • Glutinoconcha Lahr et al. 2019: shell generally made of agglutinated autogenous (idiosomes) or exogenous (xenosomes) material.[19]
              • Sphaerothecina Kosakyan et al. 2016: test rigid or slightly flexible, either completely chitinoid or comprising recycled organic or mineral particles held together by an organic cement, or composed of self-secreted chitinoid or siliceous elements; test always rounded in radial symmetry but varying in height from flattened saucer-shaped, hemispheric or more elongated to egg-shaped; pseudostome circular or lobed, surrounded by a collar; produce thick, digitate pseudopodia.[19]
              • Longithecina Lahr et al. 2019: test rigid, agglutinated from mineral particles, recycled diatom frustules, scales or plates, and/or composed of siliceous self-secreted particles (idiosomes) held together by an organic cement.[19]
              • Cylindrothecina González-Miguéns, Todorov, Blandenier, Porfirio-Sousa, Ribeiro, Ramos, Lahr & Lara 2022 > Cylindriflugiidae González-Miguéns, Todorov, Blandenier, Porfirio-Sousa, Ribeiro, Ramos, Lahr & Lara 2022: distinguished by specific sequences of the mitochondrial and nuclear DNA markers (COI, NADH and SSU) and its phylogenetic placement.[20]
              • Excentrostoma Lahr et al. 2019: shells compressed in the axis perpendicular to the aperture ("dorso-ventrally compressed"); aperture off-center; position of aperture considered an adaptation to dry environments.[19]
              • Hyalospheniformes Lahr et al. 2019 > Hyalospheniidae Schultze 1877 emend. Kosakyan et al. 2012: Test rigid, colorless or yellowish-brown, flask or vase shaped, oval or pyriform, compressed in the axis paralell to the apertural plane ("laterally compressed"); shell entirely self-secreted composed of organic matrix, or reinforced through addition of self-secreted or recycled siliceous plates; aperture terminal and bordered by an organic rim.[19]
              • Volnustoma Lahr et al. 2019: test reinforced with mineral particles, compressed in the axis paralell to the apertural plane ("laterally compressed"), with conspicuous slit-like aperture.[19]
          • Euamoebida
      • Evosea Kang et al. 2017: phylogenetic definition; wide range of morphologies; many with complex life cycles including amoeboid, ciliated and fruiting stages.[8]
    • Obazoa Brown et al. 2013: phylogenetic definition.
      • Apusomonadida Karpov & Mylnikov 1989: gliding cells (5–15 µm), with dorsal cell membrane underlain by thin theca extending laterally and ventrally as flanges that delimit a broad ventral region from which pseudopodia develop in most genera; with two heterodynamic cilia, the anterior enclosed by sleeve-like extension of flanges to form a proboscis, and the posterior cilium lying within the ventral region; tubular mitochondria cristae; phagocytosis of bacteria.
      • Breviatea Cavalier-Smith 2004: amoeboid gliding cells (10–15 µm) with single anteriorly directed apical cilium and in some isolates a second posteriorly directed cilium; filopodia projecting unilaterally from cell, perpendicular to anteroposterior axis and direction of movement; filopodia forming at anterior end, moving posteriorly as cell moves forward (filopodia appearing attached to substrate), and resorbed at posterior; cell can also produce broad lamellopodia; anaerobic or microaerophilic, with large mitochondrion-like organelle; ingests bacteria; can form cysts.
      • Opisthokonta Cavalier-Smith 1987 emend. Adl et al. 2005: single posterior flagellum without mastigonemes, present in at least one life cycle stage or secondarily lost; with a pair of kinetosomes or centrioles, sometimes modified; flat (rarely tubular) mitochondrial cristae in the unicellular stage.
        • Nucletmycea Brown 2009 (=Holomycota Liu 2009)
          • Rotosphaerida Rainer 1968 (=Cristidiscoidida Page 1987, Cavalier-Smith 1993, 1998; Nucleariidae Patterson 1983, 1999)
          • Fungi R.T. Moore 1980: phylogenetic definition; no unambiguous synapomorphies;[8] polarisome; gene families MedA and APSES;[21] ancestrally phagotrophic amoeboflagellates with endobiont feeder stage and free-living dispersal stage.[22]
            • Rozellomyceta Tedersoo et al. 2018 > Rozellomycota Doweld 2013 (=Opisthophagea Galindo et al. 2022): vegetative cells amoeboid, with pseudopodial extensions extending around host organelles; zoospores with a posterior flagellum that has a solid rhizoplast associated with a long kinetosome; one single large mitochondrion (missing in Microsporidea); resting spores thick-walled; chitinous wall present only in some life stages; penetration of host cells via germ tube; intracellular obligate parasites of fungi, animals and protists that consume host organelles via phagocytosis;[23] ancestrally chitin-based cell-walled taxa as food source.[22]
            • Phytophagea Galindo et al. 2022: ancestrally cellulose-based cell-walled taxa as food source.[22]
        • Holozoa Lang et al. 2002: phylogenetic definition.[8]
          • Ichthyosporea Cavalier-Smith 1998 (=Mesomycetozoea Mendoza et al. 2002): single-celled trophic organisms, Ichthyophonus with hyphal multinucleated filaments; flat mitochondrial cristae but some may have tubular mitochondrial cristae; if present, single cilium; without collar or cortical alveoli; some species form only elongate amoeboid cells; most animal parasites, some free-living and saprotrophic; chitin reported in cell wall (proven by staining with wheat germ agglutinin and molecular phylogeny of chitin synthases); both marine and freshwater.[8]
          • Pluriformea Tikhonenkov, Hehenberger, Mylnikov & Keeling 2017: phylogenetic definition. Syssomonas Tikhonenkov, Hehenberger, Mylnikov & Keeling 2017, Corallochytrium Raghu-Kumar 1987.
          • Filozoa Shalchian-Tabrizi et al. 2008
            • Filasterea Shalchian-Tabrizi et al. 2008
            • Choanozoa Brunet & King 2017
              • Choanoflagellata Kent 1880-1882:
              • Metazoa Haeckel 1874 emend. Adl et al. 2005 (=Animalia Linnaeus 1758): sexual reproduction through an egg cell, fertilized usually by a uniciliated sperm cell with acrosome; embryonic development with blastula followed by gastrulation that begins the differentiation into endoderm, ectoderm, mesoderm, and neuroderm (except Porifera); tissues organized into organs that share tasks for the individual, unless secondarily lost; coordination of cells and tissues by membrane receptors that respond to ligands through elaborate signal transduction; cell–cell junctions with belt desmosomes or zonulae adherentes; basal lamina and extracellular matrix with collagen and other fibrous proteins (laminin, nidogen, perlecan); heterotrophic nutrition with secretion of digestive enzymes and osmotrophy through a digestive tract; without cell wall; ectoderm completely surrounding body, and endoderm surrounding a digestive tract; sensory cells in epithelium; nervous tissue in organized network; epithelial actin–myosin-based contractile cells between endoderm and ectoderm; some tissues with phagotrophic cells.[8]
                • Porifera Grant 1836: flat mitochondrial cristae; if sexual reproduction present, zygotes forming larva (nine known larval types) or juveniles; asexual reproduction by gemmules, budding or fragmentation; sessile adult; differentiation of larva to a variety of cell types, including choanocytes, amoeboid cells and cells with granular inclusions; cell types transformable into other types as necessary; cells more or less independent; without mesoderm, nervous tissue, desmosomes, localized gonad or glandular digestive cells.[8]
                • Ctenophora
                • Planulozoa
                  • Placozoa
                  • Cnidaria
                  • Bilateria
                    • Xenacoelomorpha
                    • Nephrozoa
                      • Deuterostomia: gill slits (faringotremy).[25]
                      • Protostomia
                        • Spiralia
                        • Ecdysozoa
                          • Scalidophora (?)
                          • Cryptovermes Howard et al. 2022: phylogenetic definition.[28]
                            • Nematoida
                            • Panarthropoda sensu Nielsen 1995: true metameric limb pairs.[29]
                              • Tardigrada
                              • Onychophora
                              • Arthropoda Gravenhorst 1843 (not von Siebold 1848):[30] arthrodized appendages.[29]
                                • “Dinocaridids” (†)
                                • Isoxyidae (†)
                                • Euarthropoda Lankester 1904 (=Heptopodomera Aria et al. 2015): arthrodization of both body and all appendages; arthrodized biramous appendage, composed of basipod, endopod and exopod; seven-podomerous endopods.[29]
                                  • “Megacheirans” (†)
                                  • Cenocondyla Aria 2019: phylogenetic definition; antennules, protochelicerae or chelicerae and, in at least early representatives, with cephalon housing biramous appendages whose basis is differentiated as a masticatory device (dented gnathobase, spinose endites or mandible).[29]
                                    • Arachnomorpha (?) Lameere 1890: all cephalic endopods fully developed; third cephalic appendage gnathobasic (gnathobasipod);[29] presence of gnathobase(s); trunk endopods ending in set of three claws ("apotele"); posteriormost trunk tergites fused into single plate.[31]
                                      • Panchelicerata sensu Aria & Caron 2017: ground pattern of a seven-segmented prosoma; trunk appendages with reduced or vestigial endopods; differentiation of the seventh prosomal appendage.[31]
                                        • Habeliida (†) Aria & Caron 2017: cephalic shield with sub-triangular, sub-horizontal pleural expansions and with antero-lateral notches accommodating pair of lateral compound eyes with no peduncle; cephalic shield with large mesio-dorsal bulge accommodating stomach; five pairs of anterior, slender and segmented antennule-like exopods likely inserted below the eyes and dorsally to other head appendages; on ventral side of head, reduced pair of appendages inserted anteriormost (presumed in Sanctacarididae), followed by five pairs of appendages composed of gnathobasic basipods increasing in size posteriad and bearing seven-segmented spinose/setose enditic endopods projecting anteriad; trunk bearing paddle-like exopods fringed with thin lamellae.[31]
                                        • Chelicerata Heymons 1901: chelicerae.[29]
                                      • Artiopoda (†)
                                    • Mandibulata Snogdrass 1938: mandibles[29] (the appendage of the post-tritocerebral segment, embedded in a chewing chamber between the clypeolabrum and hypopharynx); gnathal edge of mandibles (a modified coxal endite) modified into a dentate incisor part and a molar part for chewing, with a surface formed from rows of fused spines; gradient of decreasing expression of gene Distal-less, wholly lacking along the inner margin of the mandible (area corresponding to the gnathal edge); expression of Dachshund strong in the area corresponding to the tooth-like parts of the mandible; appendage of the deutocerebral segment modified as an antenna; five appendage-bearing head (antennal/antennular, intercalary/second antennal, mandibular, first maxillary, and labial/second maxillary segments) with a differentiation of maxillules as mouthparts;[32] paired lateral buds on the mandibular sternum, that give rise to either the paragnaths or components of the hypopharynx; first maxilla as a gnathal appendage; a conserved midline neuropil embedded in the protocerebral matrix (lacking in diplopods), not lying superficial to the protocerebrum; midline neuropil 2, a unique neuropil in the central body of the brain; size variability among the somata that supply cerebral neuropils, not uniform; olfactory lobe contained within the deutocerebrum; stomatogastric and labral nerves connected to the tritocerebrum, not deutocerebrum; crystalline cone developed in the dioptric apparatus; low, fixed cell numbers in each subunit of lateral eyes; interommatidial pigment cells; reduced and more fixed number of serotonergic neurons, cells individually identifiable and typically developed singly or in pairs, to a maximum of four neurons in each group; pair of serotonergic neurons with neurites that cross to the contralateral side.[33]
                                      • Myriapoda
                                      • Fuxianhuiida (†)
                                      • Pancrustacea Zrzavý & Štys 1997 (=Tetraconata Dohle 2001): tritocerebral appendage pair absent; in crown-group Pancrustacea, protocerebral bridge, paired olfactory lobes, serotonin immunoreactivity in the ventral nerve cord, and ultrastructure of the compound eye comprising four crystalline cone cells.[32]
                                        • Hymenocarina (†) Clarke 1882: bivalved carapace with a highly convex cross section covering the cephalothoracic region; cephalothorax bearing well-developed multisegmented antennules and endopods with well-developed paired terminal claws; limb basis enditic and externally subdivided; frontalmost inter-ocular complex composed of a median sclerite flanked by lobate protrusions; post-antennular pair of appendages generally not developed; mandibles broad and rounded with uniform masticatory margins; posterior tagma (abdomen) with segments forming tergo-pleural rings; tailpiece bearing well-developed caudal rami.[34]
                                        • Clypecarididae (†) Hou, 1999 emend. Zhai et al. 2019: bivalved euarthropods with smooth shield, paired stalked eyes (absent in some forms), well-developed uniramous antennules, biramous appendage-bearing trunk segments, three or eight appendage-less abdominal ring-like segments, and an elongate cylindrical terminal telson associated with paired segmented caudal rami with posterior-facing straight setae; serrations on antero-lateral margins of bivalved shield expressed in some forms. Segments posterior to shield hinge bear multiple pairs of dorsal spines connected to the body by round sockets; biramous trunk appendages consist of endopod with rounded endites, and setae-bearing exopods.[32]
                                        • Oligostraca Zrzavý, Hypša & Vlášková 1998: short oligomeric trunks.[35]
                                        • Communostraca
                                        • Allotriocarida Oakley et al. 2012.[36]
  • EXCAVATES (Excavata Cavalier-Smith 2002 emend. Simpson 2003) (P)
    • Metamonada
    • Discoba
      • Jakobida
      • Tsukubamonadida
      • Discicristata
        • Heterolobosea
        • Euglenozoa
          • Euglenida Butschli 1884 emend. Simpson 1997: with a pellicle of proteinaceous strips, fused in some taxa; when unfused and with more than ~20 strips capable of active distortion (metaboly); where known, paramylon is the carbohydrate storage product.[8]
            • Petalomonadida Cavalier-Smith 1993: strictly non-photosynthetic, phagotrophic bacterivorous euglenoids with an aplastic pellicle of a few longitudinally arranged strips;[37] rigid cells without supporting rods or cemented oral supports; vanes present or absent; glide on anterior cilium.[38] Dolium, Notosolenus, Petalomonas, Scytomonas, Sphenomonas.[39]
            • Alistosa Lax & Simpson 2021: phylogenetic definition. Ploeotia, Serpenomonas, Keelungia, Lentomonas, Decastava.[40]
            • Gaulosia Lax, Cho & Keeling 2023: free-living, rigid, biflagellated heterotrophic euglenid, oblong profile with a markedly rounded, blunt posterior; cell slightly flattened ventrally; anterior flagellum ~0.8× cell length, thickened posterior flagellum ~2.2× cell length; slow gliding speed, when under stress slowly retracts on posterior flagellum; 10 faint broad pellicle strips can be seen with light microscopy.[41]
            • Karavia Lax, Cho & Keeling 2023: phylogenetic definition. Hemiolia, Liburna.[41]
            • Olkaspira Lax & Simpson 2021: phylogenetic definition.[40]
              • Olkasia
              • Spirocuta
                • Anisonemia
                • Peranemida (P)
                • Euglenophyceae Schoenichen 1925 emend. Marin & Melkonian 2003 (=Euglenea Butschli 1884 emend. Busse & Preisfeld 2002): phototrophic, with one to several plastids of secondary origin with three bounding membranes and chlorophylls a and b; some species secondarily non-photosynthetic; most with extraplastidic eyespot and photosensory apparatus associated with cilia; most motile.[8]
                  • Rapazia > Rapazida > Rapazidae Cavalier-Smith 2016: phagophototrophs;[42] cells solitary, cytotrophic on microalgae with two heterodynamic cilia of unequal length; pellicle with helically arranged strips capable of metaboly locomotion; discoidal chloroplast(s) with pyrenoids surrounded by three membranes; marine species. Rapaza Yamaguchi et al. 2012.[8]
                  • Euglenophycidae Busse & Preisfeld 2003: non-phagotrophs.
                    • Eutreptiales Leedale 1967 emend. Marin & Melkonian 2003 > Eutreptiidae Hollande 1942: 2–4 emergent heterodynamic cilia of equal or unequal length; cells not rigid, usually capable of metaboly; mostly marine or brackish species, rarely freshwater.[8] Eutreptia, Eutreptiella (=Tetreutreptia).[43]
                    • Euglenales Leedale 1967 emend. Marin & Melkonian 2003
          • Glycomonada Cavalier-Smith 2016: heterotrophs, ancestrally phagotrophs; microbodies are glycosomes containing glycolytic enzymes, not peroxisomes; mitochondria ancestrally polykinetoplastid; mitochondrial genomes of multiple heterogeneous circles; transcripts undergo RNA editing including uridine insertion; ciliary pocket without dorsal row microtubules, unlike Euglenida and Postgaardia; pellicle microtubules ancestrally in a continuous cross-linked corset, nucleated posteriorly; cytostome ancestrally present at tip of pronounced apical rostrum separated from ciliary pocket by narrow preoral crest; feeding apparatus ancestrally with left and right ‘jaw-bone’-like cement lip supports, each associated with short lateral dense rods that extend only a short way from cytostome; ancestrally with hairs on preoral crest, and circumferential encircling microtubules surrounding cytostome; three microtubule sets (nucleated in ciliary pocket) loop over to support the left side of cytopharnx: 4–8 central widely spaced microtubules reinforced by characteristic dense material (MTR) flanked by close-set less reinforced microtubules on dorsal (parallel microtubule loop (PML) microtubules, mostly paired with intrapair thin laminas) and ventral external microtubule band (EMB) sides; ancestrally MTR microtubules had flanges on inner face of loop and vanes on cytostome-associated outer part, secondarily lost in most kinetoplastids.[42]
            • Kinetoplastea Honigberg 1963: cells with a kinetoplast (large mass(es) of mitochondrial DNA); mitochondrial RNA editing; trans-splicing of splice leader RNA; polycistronic transcription.[8]
            • Diplonemea Cavalier-Smith 1993 emend. Simpson 1997: keterotrophic cells exhibiting pronounced metaboly; both cilia are short and usually supported with paraciliary rod; apical papilla, feeding apparatus with 'pseudovanes'; few giant, flattened mitochondrial cristae.[8]
          • Symbiontida Yubuki et al. 2009 (=Postgaardia Cavalier-Smith 2016; Postgaardea Cavalier-Smith 1998 emend. 2016):[42] microaerobic or anaerobic cells that possess rod-shaped epibiotic bacteria above superficial layer of mitochondrion-derived organelles with reduced or absent cristae.[8]
            • Postgaardida Cavalier-Smith 2003 emend. 2016: microtubule band of four or about sixteen microtubules loops over from ciliary pocket to cytopharynx dorsal margin in a plane orthogonal to five reinforced microtubules that loop over apically from ciliary pocket to cytopharynx via six finger-like projections inside mouth of cytopharynx; no supporting rods or dorsal/left jaw support homologues. Rigid cells with homogeneous pellicle with a dense thin (∼25 nm) lamina underlying microtubules; cytostome ventral groove.[42]
              • Calkinsiidae Cavalier-Smith 2016: phagotrotrophs with MTR pocket cytopharynx that eat diatoms and bacteria; cytostome opens on right of dorsal cilium into a shared apical depression (vestibulum) close to ciliary pocket; the cytopharyngeal loop has about 16 microtubules with dense flanges on the inner side of the loop at the cytopharyngeal end; glide on largely rigid anterior cilium. Calkinsia Lackey 1960.[42]
              • Postgaardidae Cavalier-Smith 2016: heterotrophs with cytopharynx opening separately from ciliary pocket; 5 reinforced microtubules (MTR) loop from ciliary pocket, along right side of U-shaped gutter oriented posteriorly from the cytostome, turn anteriorly along its left side halfway back towards cilia, then loop inwards and posteriorly alongside cytopharynx that opens into left gutter; an orthogonal looping band of four non-flanged microtubules passes from ciliary pocket end of MTR along the anterior half of the left gutter; gutter covered by two overlapping longitudinal flaps, inner reinforced ridge on left and anterior lip on right; swim, not glide. Postgaardi Fenchel et al. 1995.[42]
            • Bihospitida > Bihospitidae Cavalier-Smith 2016: metabolic anaerobic bacterivorous phagotrophs with pellicle longitudinally subdivided into extrusome-delimited S-shaped regions formed by grooves containing epibiotic bacteria and ridges with underlying mitochondria; cytostome opens on right of dorsal cilium into shared apical depression (vestibulum) close to ciliary pocket; strongly curved C-shaped rod apparatus originates at vestibulum, loops round nucleus within a nuclear envelope groove; nucleus-attached ‘main rod’ comprises a dense lamina (immediately adjacent to adhering accessory rod) bearing a stack of ∼75 broad, parallel, laminas; ‘accessory rod’ is a membrane-associated row of ∼40 reinforced microtubules with proximal dense flanges and distal paired vanes like those of diplonemids, initiated at a cytostomal funnel at the ciliary pocket and looping over to cytopharynx; an adjacent bundle of non-reinforced microtubule pairs at the ciliary pocket is not part of the ‘accessory rod’. Bihospites Breglia et al. 2010.[42]

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