
==== Front
ISME J
ISME J
ismej
The ISME Journal
1751-7362
1751-7370
Oxford University Press

39077993
10.1093/ismejo/wrae150
wrae150
Review Article
AcademicSubjects/SCI00010
AcademicSubjects/SCI00960
AcademicSubjects/SCI01150
AcademicSubjects/SCI02281
Integrated overview of stramenopile ecology, taxonomy, and heterotrophic origin
Jirsová Dagmar Center for Mechanisms of Evolution, Biodesign Institute, School of Life Sciences, Arizona State University, 1001 S McAllister Avenue, Tempe, Arizona, 85287-7701, United States
Institute of Parasitology, Biology Centre, Czech Academy of Sciences, Branišovská 31, České Budějovice 37005, Czech Republic

Wideman Jeremy G Center for Mechanisms of Evolution, Biodesign Institute, School of Life Sciences, Arizona State University, 1001 S McAllister Avenue, Tempe, Arizona, 85287-7701, United States

Corresponding authors: Dagmar Jirsová, Center for Mechanisms of Evolution, Biodesign Institute, School of Life Sciences, Arizona State University, 1001 S McAllister Avenue, Tempe, Arizona, 85287-7701, United States. Email: djirsova@asu.edu and Jeremy G. Wideman, Center for Mechanisms of Evolution, Biodesign Institute, School of Life Sciences, Arizona State University, 1001 S McAllister Avenue, Tempe, Arizona, 85287-7701, United States. Email: Jeremy.Wideman@asu.edu
1 2024
30 7 2024
30 7 2024
18 1 wrae15029 2 2024
12 6 2024
29 7 2024
30 8 2024
© The Author(s) 2024. Published by Oxford University Press on behalf of the International Society for Microbial Ecology.
2024
https://creativecommons.org/licenses/by/4.0/ This is an Open Access article distributed under the terms of the Creative Commons Attribution License (https://creativecommons.org/licenses/by/4.0/), which permits unrestricted reuse, distribution, and reproduction in any medium, provided the original work is properly cited.

Abstract

Stramenopiles represent a significant proportion of aquatic and terrestrial biota. Most biologists can name a few, but these are limited to the phototrophic (e.g. diatoms and kelp) or parasitic species (e.g. oomycetes, Blastocystis), with free-living heterotrophs largely overlooked. Though our attention is slowly turning towards heterotrophs, we have only a limited understanding of their biology due to a lack of cultured models. Recent metagenomic and single-cell investigations have revealed the species richness and ecological importance of stramenopiles—especially heterotrophs. However, our lack of knowledge of the cell biology and behaviour of these organisms leads to our inability to match species to their particular ecological functions. Because photosynthetic stramenopiles are studied independently of their heterotrophic relatives, they are often treated separately in the literature. Here, we present stramenopiles as a unified group with shared synapomorphies and evolutionary history. We introduce the main lineages, describe their important biological and ecological traits, and provide a concise update on the origin of the ochrophyte plastid. We highlight the crucial role of heterotrophs and mixotrophs in our understanding of stramenopiles with the goal of inspiring future investigations in taxonomy and life history. To understand each of the many diversifications within stramenopiles—towards autotrophy, osmotrophy, or parasitism—we must understand the ancestral heterotrophic flagellate from which they each evolved. We hope the following will serve as a primer for new stramenopile researchers or as an integrative refresher to those already in the field.

Graphical Abstract

Graphical Abstract

stramenopiles
heterotrophic flagellates
plastid evolution
chromalveolate hypothesis
rhodoplex hypothesis
protistology
microbial ecology and evolution
National Science Foundation 10.13039/100000001 2119963 2405455 Gordon and Betty Moore Foundation 10.13039/100000936 GBMF9201 VEDA FELLOWSHIPS within the Operational program Jan Amos Komensky CZ.02.01.01/00/22_010/0008117
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pmcIntroduction

Stramenopiles are mostly microbial eukaryotes, and can be found in virtually all habitats, including freshwater and soil [1, 2], oceans [3, 4], arctic waters [4, 5], the deep sea [6, 7], and deserts [7–9]. Most people are familiar with stramenopiles, even if they have never heard the word. Kelp are stramenopiles—so are diatoms. The Irish Potato Famine was caused by the blight-causing oomycete Phytophthora infestans, a filamentous fungus-like stramenopile [10]. Many people carry one of the subtypes of the intestinal commensal Blastocystis, a heterotrophic stramenopile that forms resilient cysts [11].

Stramenopiles are ecologically and evolutionarily important. From an ecological perspective, stramenopiles contribute greatly to carbon and nutrient cycling. Phototrophic stramenopiles, diatoms in particular, produce almost ~40% of global ocean-derived oxygen, which is comparable to the Amazonian rain forest [12]. Heterotrophic stramenopiles keep the carbon and mineral cycles churning as voracious consumers of bacteria and other microbes [13, 14]. Some stramenopiles are supplementing their photosynthetic capabilities through phagotrophy or osmotrophy, exhibiting mixotrophic life strategy [15–17]. Their position in a food chain is more complex since the ability to function as a primary producer or consumer is difficult to evaluate. Thus, phototrophic, mixotrophic, and heterotrophic groups likely play important roles in most ecosystems [2, 3, 18–20]. However, our ecological knowledge of heterotrophs is mostly limited to 18S rRNA gene amplicon sequencing (e.g. small ribosomal subunit V4 and V9 regions), which are effective [18, 19], though heavily biased [21–23], and uninformative when recovering unsampled or under investigated lineages [24]. As a recent report suggests, “One of the main issues [preventing] us from obtaining a detailed assessment on the ecology of heterotrophic flagellates […] is the lack of cultured strains that effectively represent the dominant heterotrophic flagellate species in the ocean” [18]. Therefore, traditional natural history, taxonomy, cultivation, and whole genome sequencing projects are desperately needed to improve our ecological understanding of heterotrophic flagellated stramenopiles, including their contributions to food webs and the roles they play within diverse ecosystems.

From an evolutionary perspective, stramenopiles have colonized and adapted to many unique environments and are key to understanding cell biological transitions to phototrophy, mixotrophy, osmotrophy, and parasitism. Again, though we know substantially more about phototrophs and parasites, we know very little about the free-living heterotrophic flagellates [18], from which phototrophs (including mixotrophs) and parasites emerged (e.g. [16, 25]). As eukaryotic microbial dark matter comes into light, we must develop strategies to culture and investigate newfound heterotrophic lineages that occupy important phylogenetic positions on the stramenopile tree of life. In-depth investigations of these lineages will be critical to understanding both their ecological and biochemical importance [26] and their ancestor’s capacity to diversify and occupy open niches.

Stramenopiles were originally called heterokonts, with both names deriving from shared morphological characters. Heterokont simply means “two different flagella,” and stramenopile (stramen = straw; piles = hair) derives from the hair-like tripartite tubular protrusions (mastigonemes) (Fig. 1). Mastigonemes have been recognized as characteristic of stramenopile cells for many decades [27–30]. Although they are lost in some clades (e.g. diatoms [31], Cafileria marina [32]) and present in some nonstramenopiles (e.g. cryptophytes [33], bodonids [34]), their tripartite structure and placement on both sides of the anterior flagellum make them unique to stramenopiles. In some stramenopiles, mastigonemes are not very obvious features but remain present for motile lifecycle stages (e.g. zoospores of kelp [35], oomycetes [36], and labyrinthulomycetes [37]). So far, we know that mastigonemes enhance flagellar propulsive force by increasing surface area and also act as mechanosensors [38, 39], though other potential functions remain enigmatic [40]. Genomics and cell biological investigations of stramenopiles have revealed a few more shared characters. Stramenopiles universally lack mitochondrial-encoded threonine tRNA [41], contain a fused version of the glycolysis enzymes glyceraldehyde-3-phosphate dehydrogenase and triosephosphate isomerase [42, 43], and have relocalized the ATP-yielding part of glycolysis from the cytosol to the mitochondria [43, 44]. More clade-defining features are likely present, but we lack the model systems to investigate the vast diversity of stramenopiles.

Figure 1 A representation of a typical heterotrophic stramenopile. Stramenopiles have two unequal flagella; the anterior is equipped with two rows of tripartite tubular hairs called mastigonemes. Some taxa have a specific phagocytic apparatus that acts like a “cell mouth” called a cytostome. Early phototrophic stramenopiles were likely similar to this representation but with an addition of a complex red plastid [209].

In this review, we will introduce the broad diversity of stramenopiles and highlight important aspects of their morphology, ecology, and evolutionary history. Fortunately, the interest in free-living heterotrophic stramenopiles is growing [3, 32, 45–48], which only highlights the need to further investigate this group.

Stramenopiles are the “S” in SAR

Stramenopiles, Alveolates, and Rhizarians form the monophyletic group called SAR [49] (Fig. 2). A fairly clear branching order of the three groups has emerged with rhizarians at the base of the group and alveolates + stramenopiles as sister clades to one another [50–52], although some studies place rhizarians as a sister group to alveolates and stramenopiles at the base (e.g. [53, 54]). Apart from the fact that they always branch together in gene-based phylogenetic trees, these three groups do not seem to have much in common (though some molecular synapomorphies have been identified, e.g. [55]). Though there are no apparent cellular or morphological similarities observed between SAR groups, it has been clear for several decades that stramenopiles form a monophyletic group. This early conjecture was based on their unique flagellar traits described above [56].

Figure 2 A simplified phylogeny of the SAR clade, with a focus on stramenopiles. Rhizarians and alveolates are distant relatives of stramenopiles. Within stramenopiles, there are two major well-supported groups: Bigyra and Gyrista. Whereas Bigyra consist of heterotrophs only, Gyrista include the only plastid possessing group—Ochrophyta. Within the Bigyra and Gyrista, branching orders are still being worked out as new lineages and species of stramenopiles are consistently being discovered (e.g. MArine STrameonpiles (MASTs) [4] and Platysulcus tardus [150]) or the phylogenetic position of already known species is being reclassified by more robust genomic and transcriptomic data (e.g. P. flagellatus [48, 210], ﻿A. sol [45, 48]).

Stramenopiles comprise two usually well-supported clades, the Bigyra and Gyrista [27, 57–60] (Fig. 2), although lately, the monophyly of Bigyra has been shaken [61]. Gyrista includes the predominantly osmotrophic pseudofungi and the photosynthetic ochrophytes, which consist of some lineages known only from sequence data called Marine OCHrophytes (MOCHs) [4, 48], encompassing marine as well as freshwater and terrestrial ochrophytes [48]. Bigyra are strictly heterotrophic and include bikosians, labyrinthulomycetes, and placidozoans [57, 59, 60, 62, 63]. Both Bigyra and Gyrista contain several lineages that are not well characterized, many of which are referred to as MArine Stramenopiles (MASTs) [4], even if some are not marine [4, 64]. In the following, we outline what is known about the major groups of stramenopiles and highlight avenues for future research.

Ochrophytes are the only photosynthetic group of stramenopiles

Compared to heterotrophic stramenopiles, ochrophytes have enjoyed significant scientific attention. They represent a clade of extremely diverse eukaryotic autotrophs [65, 66], which includes photosynthetic nano- and pico- algae, giant kelps, and even derived heterotrophs that secondarily lost their photosynthetic capabilities [45, 48, 67–71]. One of the peculiar features of many ochrophytes is their ability to be mixotrophs (e.g. chrysophytes [15], Microchloropsis sp. [16], Parmales/Bolidophyceae [17]). Mixotrophy occurs in many plastid-bearing lineages (e.g. green algae Prasinophyta [72], dinoflagellate Karlodinium veneficum [73], haptophytes such as Phaeocystis [74]) in which autotrophs maintain heterotrophy as a compensation for specific nutrient deficiencies [75]. As important primary producers and secondary consumers, mixotrophs have complex roles in food webs [76, 77], though they are excluded from most ecological models [76].

Ochrophytes have a specific combination of photosynthetic pigments that gives their plastid its brown colour (chlorophyll c, chlorophyll a, and the accessory pigment—fucoxanthin) [78]. Exceptions include Xanthophyceae and Eustigmatophyceae (yellow-green algae) as they have only chlorophyll a and β-carotenes [79, 80]. The best studied ochrophytes are the diatoms (Bacillariophyceae). These microalgae are known for their spectacularly shaped silica-based shells (aka frustules). Their significant impact on oxygen production [12] and carbon fixation [81] makes them the most successful ocean phototrophs. Their natural production of ﻿triacylglycerols and fucoxanthin makes them a potential candidates for the biofuel industry [82], whereas their silica shells are used for pest control [83] and as environmental fertilizers [84]. Diatoms are also one of the few genetically tractable stramenopiles [85, 86].

Though diatoms are the most scientifically explored of all stramenopiles, the most famous ochrophytes are the kelp. Kelp dominate cold ocean areas of low water motion, with kelp forests providing interesting ocean habitats [87, 88]. Kelps (Phaeophyceae, aka brown algae) have the typical brown colouring of ochrophytes and their characteristic stramenopile sexual zoospore stage, which is complete with mastigonemes that have been noted for many decades [89]. As primary producers, kelp and diatoms are essential part of food webs [90, 91], and any environmental changes will affect the species representation causing a domino effect in other ecosystem tiers [88, 92].

Besides diatoms, other highly abundant and omnipresent ochrophytes include the chrysophytes (aka golden algae) [3, 18, 93, 94]. Chrysophytes are mostly solitary biflagellate algae but can form colonies, and similar to diatoms, some species have silica shells (lorica) [15]. Chrysophytes encompass wide variation in their nutritional modes, from plastid-bearing phototrophs (e.g.﻿ Ochromonas sp. and Chromulina sp. [94]), to photosynthetic mixotrophs (e.g. some Ochromonas species [95]), bacteriovorus phago-mixotrophs (e.g. Ochromonas [95, 96]), and even heterotrophs (e.g. Spumella sp. and Spumella-like sp. [97]), which no longer photosynthesize but retain a leucoplast—a colourless plastid. Mixotrophy is not exclusive to chrysophytes but also occurs in other ochrophytes like diatoms [98, 99] and coccolithophores [100, 101]. Heterotrophic nutrient uptake in mixotrophy can occur through phagotrophy or osmotrophy, with osmotrophy being common when cells have physical barriers like silica shells [98–101]. A complete shift to osmotrophy can lead to plastid simplification and loss of photosynthetic ability, as seen in Nitzschia putrida [102]. Mixotrophy may offer a selective advantage over phototrophy or heterotrophy alone [99].

One reason chrysophytes might have maintained mixotrophy is their relatively poor capacities to fix carbon, assimilate nitrogen [103–105], and uptake phosphorus [106, 107]. Additionally, ﻿chrysophytes cannot make thiamine (B1 vitamin) [25], which leads to a dependency on specific members of the microbial community [108]. The mixotrophy amongst algae seems to be more prevalent than we originally thought and was adopted or maintained multiple times in different lineages [109, 110], perhaps rightfully so, as it is the transitional stage between heterotrophy and obligate autotrophy. Therefore, mixotrophy is an important part in plastid acquisition and its genomic and metabolic integration (see Ochrophyte plastid: count the gains, not the losses section).

As mixotrophs, chrysophytes are metabolically and genetically diverse [25, 95]. With the plastid comes advantages and trade-offs as the new organelle is adopted into a functioning eukaryotic cell; some metabolic pathways show significant differences compared to heterotrophs. The most prominent alterations can be seen in the heme pathway [70, 111], isoprenoid [67], and fatty acid synthesis [112, 113]. In addition to their significant ecological impact, this makes the chrysophytes great model organisms for studying plastid genome reduction and metabolic evolution [69]. Besides these well-described taxa, 10 MOCH clades are scattered across ochrophytes, which are known exclusively from sequence data [4, 48].

Ochrophyte plastid: count the gains, not the losses

Although ochrophytes are the most studied group of stramenopiles, their plastid’s origin story remains incomplete. The group evolved over 500 million years ago [114], but the evolutionary origin of its plastid is only just coming to light. Although it is undisputed that the ochrophyte plastid has a red algal origin [115–117], there are two predominating theories of how this arose (Fig. 3). In the first scenario, the ancestor of the SAR lineage had already acquired a secondary red plastid, which was subsequently lost in all related heterotrophic lineages (Fig. 3A). The second scenario starts with a heterotrophic ancestor and requires multiple plastid acquisitions across the evolutionary tree, including one in stramenopiles (Fig. 3B).

Figure 3 The origin of complex plastids. (A) The chromalveolate hypothesis. The originally proposed chromalveolate hypothesis [118] suggests that all CASH plastids arose from a single secondary endosymbiotic event. Complex plastids were thought to be lost in a few known heterotrophic lineages related to complex plastid-bearing lineages. The chromalveolate hypothesis was supported by early phylogenies that erroneously grouped the Cryptophyta, Alveolata, Stramenopila, and Haptophyta (CASH lineages) together based on striking similarities of their plastids genes. However, with new data, for this hypothesis to remain in contention, organellar loss must be an easier accomplishment than once thought. (B) The rhodoplex hypothesis. As more heterotrophic lineages were shown to be more closely related to individual CASH lineages than the CASH lineages were to one another, more and more losses were required for the chromalveolate hypothesis to remain true. Instead, data now strongly suggest that the ancestor of stramenopiles (and SAR) was heterotrophic. Thus, instead of a single plastid gain followed by multiple losses, four serial gains of complex plastids have been reported explaining plastid possession in the CASH lineages. In the stramenopiles, recent studies strongly suggest that the plastid of ochrophytes was obtained through a tertiary endosymbioses of a cryptophyte [147].

How many losses are too many losses?

The first hypothesis—the “chromalveolate hypothesis” [118]—posits the ancestor of a now-defunct photosynthetic clade called chromalveolates (stramenopiles, alveolates, cryptophytes, and haptophytes) had a secondary red algal-derived plastid [119, 120]. The chromalveolate hypothesis presumes that all heterotrophic lineages related to any of these groups independently lost their plastid. When this hypothesis was first proposed, only a few related heterotrophic SAR lineages were known (e.g. ciliates—a group of alveolates covered with hair-like cilia, oomycetes, goniomonads—aplastidial cryptists), suggesting that multiple plastid losses were the most feasible scenario. Shared characters between chromalveolates such as the presence of chlorophyll c [78, 118] and robust plastid phylogenies [121–123] supported the monophyly of this group. Additionally, some evidence of the historical plastid was found in heterotrophic lineages (e.g. oomycetes [124], labyrinthulomycetes [125]). Unfortunately, these early analyses had suffered from a lack data for known groups. They focused nearly exclusively on phototrophs, and conclusions were based largely on plastid genes [126], or very small datasets in which signal was indistinguishable from noise [127]. Thus, although the multiple plastid losses proposed by the chromalveolate hypothesis seemed likely at the time, the rarity of organellar loss was not yet fully appreciated or understood.

Now we know that once an endosymbiotic organelle is acquired, its complete loss is exceedingly rare—although reduction and loss of organellar functions and even genomes are quite common (e.g. highly reduced mitochondria in anaerobic lineages [128, 129]). In the case of mitochondria, the only confirmed loss of the organelle occurred in the anaerobe Monocercomonoides [130]. In parasitic lineages, organellar reduction is not surprising, as most simplify their metabolism because of capitalizing on host-derived substrates [131–133]. Despite the commonness of functional simplification, we know of only two examples of complete plastid loss in parasites: the intestinal apicomplexan Cryptosporidium [134] and the parasitic dinoflagellate Hematodinium sp. [135]. An example of a putative loss in a free-living lineage has only recently come to light in the Picozoa, which branch sister to red algae and Rhodelphis [136]. Finally, recently discovered plastid losses have been identified in the ochrophyte lineage (e.g. the flagellate ﻿Picophagus flagellatus and the heliozoan ﻿Actinophrys sol [48]). Although these cases demonstrate the possibility of total organelle loss, it speaks to the rarity of the event.

In acknowledgement of the rarity of plastid loss, the chromalveolate hypothesis became less feasible. Furthermore, as rhizarians were shown to be related to stramenopiles and alveolates, and cryptophytes and haptophytes were revealed to branch only distantly to the SAR lineage [137, 138], more and more plastid losses were required for the chromalveolate hypothesis to remain true. As more heterotrophs were found, a growing number of aplastidial lineages were shown to be more closely related to individual lineages within chromalveolates than the photosynthetic chromalveolates were to each other. These discoveries resulted in the eventual downfall of the chromalveolate hypothesis. Thus, the field concluded that plastid gain is not as rare as we once thought, and it is now accepted that multiple plastid gains is the more likely scenario. For example, considering only alveolates and stramenopiles, the minimum number of plastid losses required for the chromalveolate hypothesis to remain in contention is between five and seven [46], whereas only two red-algal derived plastid gains are required within the entire SAR clade.

How can multiple gains be a realistic explanation?

With the chromalveolate hypothesis disproven, a multiple origins hypothesis became necessary. Remember, the chromalveolate [now sometimes referred to as “CASH” (Cryptophytes, Alveolates, Stramenopiles, and Haptophytes)] lineages of plastids all contain chlorophyll c and are very closely related based on plastid phylogenies [121, 139]. These data suggest that their plastid came either from multiple secondary endosymbioses of very closely related red algae or a series of higher-order endosymbioses that occurred between CASH groups. Again, the first option seems simpler. However, the secondary plastids of cryptophytes, stramenopiles, and haptophytes all share a phylogenetically related and specialized protein sorting machinery derived from the ER-Associated protein Degradation system (ERAD) of red algae [140–142]. This machinery is called SELMA (Symbiont-specific ERAD-Like MAchinery) and is responsible for transporting proteins into and through the second (from the outside) plastid membrane [143, 144]. These data strongly suggest that CASH plastids all have a common endosymbiotic origin. But how can that be if multiple endosymbioses are necessary?

The rhodoplex hypothesis starts with a heterotrophic ancestor and introduces a new element of complexity. Instead of multiple secondary origins, only a single secondary endosymbiotic event occurred, and other red algal-derived plastids were acquired by tertiary or even quaternary endosymbioses [139, 145, 146]. The rhodoplex hypothesis was originally proposed as agnostic towards which lineage contained the original secondary endosymbiosis. Recent papers have suggested that molecular timescales and phylogenetic analyses are still compatible with multiple downstream scenarios; however, the original secondary endosymbiosis has likely been narrowed to the cryptophyte lineage [117, 147, 148]. These findings indicate that ochrophyte, alveolate, and haptophyte algae could have been derived from tertiary endosymbioses of cryptophytes or even more complex quaternary endosymbioses [117]. Additionally, modern cryptophytes have diverged quite recently (~200–300 MYA [147]); thus, most of the available evidence suggests that ochrophytes acquired their plastid via tertiary endosymbioses of an extinct lineage of cryptophytes [66, 147, 142] plausibly via kleptoplasty as the nucleomorph is missing [149].

Towards an understanding of the ancestor of ochrophytes

With the fall of the chromalveolate hypothesis came an understanding that plastid loss has not happened in the stramenopile heterotrophic lineages. Thus, stramenopiles are ancestrally heterotrophic. So far, no evidence of historical plastid presence has ever been demonstrated in any plastid-lacking stramenopile lineage, with the above-mentioned exceptions of ﻿P. flagellates and ﻿A. sol [48]. No evidence of historical plastid loss remains for the major lineage comprising pseudofungi and bigyromonads [46], nor any bigyran, nor the deep-branching Platysulcus tardus [150, 59]); therefore, we can conclude that the ancestral stramenopile was a free-living heterotroph.

Oomycetes: pseudofungi with heterotrophic flagellated relatives, the ﻿bigyromonads

Unlike autotrophic stramenopiles that cluster into a single group, heterotrophic stramenopiles can be found across several major stramenopile clades [50, 57]. Sister to ochrophytes are the pseudofungi (Oomycetes) and bigyromonads [57, 151], which includes saprotrophs [152, 153], though the best known and studied representatives are the parasitic oomycetes (aka water moulds). The lineages closely related to oomycetes are nonparasitic, but maintain an osmotrophic, fungi-like lifestyle (e.g. Hyphochytrium [27, 154]), Although they look like fungi, pseudofungi have zoospores with two unequal flagella and mastigonemes as a part of the life cycle [36]. Oomycetes can be found in aquatic and terrestrial habitats, have a filamentous structure, and because of this, were historically considered as a basal group of “true” fungi [155]. The similarities between fungi and pseudofungi, including their cell structure and osmotrophic lifestyle, were greatly affected by horizontal gene transfers (HGTs) from bacteria, animals [156], and fungi [157]. In particular, many genes associated with parasitism were transferred, including those encoding enzymes that break down polysaccharides and associated transporters that enable a parasite to feed on nutrients obtained from the host [158]. Historically, oomycetes provided key evidence for the chromalveolate hypothesis as their genomes contained genes of “red algal” origin [124]. However, the red algal ancestry of these genes was disproven by analysing a larger data set with different methods [120, 127].

Sister to these filamentous lineages [46], and somewhat overlooked due to the general lack of data, are the bigyromonads. This group of heterotrophic phagotrophs include the eukaryovore Develorapax marinus [159], the bacteriovores Developayella elegans [160], and Mediocremonas mediterraneus [161]. With bigyromonads as sisters to the oomycetes, investigations of these free-living heterotrophic flagellated stramenopiles are required to better understand transitions to osmotrophy, parasitism, and a filamentous cellular structure.

Bigyra are a melange of mostly heterotrophic flagellates

In comparison to Gyrista, we know substantially less about Bigyra. Most known taxa are free-living heterotrophs, though some parasitism/commensalism has evolved in the opalinids and labyrinthulomycetes (e.g. Blastocystis sp. [162], Aplanochytrium kerguelensis [163]). Beyond Blastocystis, the best studied groups are the labyrinthulomycetes and bicosoecids (discussed below). Most of the assorted unidentified stramenopiles are known almost completely by sequence data and are called MArine STramenopiles (MASTs) [4, 64, 93, 164, 165]. Over the past years, the number of newly identified organisms in these groups is constantly rising due to environmental DNA surveys. Environmental studies (e.g., Tara Oceans expeditions) confirm this richness and estimate thousands of unidentified species belonging to MAST clades [4, 23, 59, 166]. Unfortunately, for the vast majority of MASTs, we lack microscopic observation and only have 18S or single-cell omics data, making it almost impossible to pair an organism with its sequence data [59, 164, 166, 167]. As MASTs are being cultured, morphologically described, and formally classified, the moniker will slowly disappear (e.g. MAST-3 are called Nanomonadea [168], and MAST-4 and 6 are called Eogyrea [59, 169]). To date, only a few MASTs have been phylogenetically placed [60, 164], with many more awaiting rediscovery and further investigation.

Branching outside both the Bigyra and Gyrista is a single species, Platysulcus tardus, an enigmatic heterotrophic flagellate [150, 59]. This stramenopile shares some key stramenopile features—two uneven flagella, the anterior equipped with mastigonemes, microtubular root, and its mitochondria exhibit tubular cristae [150]—but this species also has a few unique characteristics (﻿e.g. a flat vesicle surrounds the cytoplasm containing a nucleus, mitochondria, and microbodies). As we explore the enigmatic heterotrophs, we will no doubt begin to understand their diversified morphology and ecological strategies.

Labyrinthulomycetes comprise slimy heterotrophs, flagellates, and some surprises

Bigyra also includes a somewhat well-studied group of filamentous heterotrophs called Labyrinthulomycetes. They are named for their most famous members, which create labyrinthine slime nets or “slime tubes.” These tubes create a network on which cells slither and absorb nutrients [170]. Most described species are marine saprotrophs, though some parasites of algae and animals also exist [171–174]. Beyond the more famous members for which they are named, labyrinthulomycetes also contain phagotrophs, including species with amoeboid stages that nutritionally exploit single-cell algae [175] and mixotrophic taxa with endosymbiotic green algae [176]. In the same way as oomycetes, labyrinthulomycetes have motile spores with two flagella, the anterior equipped with mastigonemes [177]. Though the group is mostly known for being marine, freshwater and terrestrial members are also being identified [177–179]. For a time, the labyrinthulomycetes entered the Chromalveolate debate [170] because of their abilities to produce omega-3 polyunsaturated fatty acids using a plastid-like desaturase [125]. However, this biochemical anomaly was later attributed to HGT from marine bacteria [180]. The biochemistry of this group has become industrially relevant, and as a result, a few species have been recently developed as genetically tractable models (e.g. Parietichytrium sp. and Thraustochytrium sp. [181]). This lineage is only just beginning to receive the attention it deserves. Similar to ochrophytes and oomycetes, an ancestral heterotrophic flagellated stramenopile was inevitably the predecessor to this clade, and once again, a better understanding of the transition to a fungal-like form requires deeper investigations into heterotrophic flagellates.

Bikosia are key heterotrophs in aquatic ecosystems

As prime bacterial consumers, bikosians are an essential part of food webs and nutrient cycling through remineralization [13, 182, 183]. About 20 well-defined species of bicosoecids have been described with some affiliations to MASTs [57, 164, 184]. Despite their previous assignment near photosynthetic ochrophytes based on their ultrastructure [185, 186], phylogenetic analyses consistently show that bicosoecids are definitively bigyrids [187], with Opalinata (Slopalinida and Blastocystis) and placidideans branching nearby [57, 59, 60]. Bicosoecids share with chrysophytes a putatively ancestral-like feeding basket with a unique microtubular organization [169, 187] and also the heterotrophic/phagotrophic life strategy [95, 188]. The bicosoecid flagellar apparatus has a unique microtubular root (R3) that loops around the cell and creates additional physical support [184].

Bicosoecids include one of the best studied free-living heterotrophic stramenopiles, Cafeteria roenbergensis [189], as well as several more recently described species (e.g. new species in Cafeteria [9], Cafileria marina [32], and Bilabrum latius [190]). Cafeteria is a common and highly abundant part of marine plankton with many well-identified species [7, 166]. Its mitochondrial genome was the first to be assembled from Bikosia [191], and drafts of nuclear genomes were recently published [192]. Even prior to obtaining genomic data, C. roenbergensis was a model for studying bacterial grazing [193–195] and viral infection for the giant marine DNA virus CroV [196, 197]. Thus, Cafeteria should be an ideal model organism to study heterotrophic stramenopiles. However, we still lack basic information about its cell biology, further necessitating the development of more genetically tractable model organisms in the stramenopile lineage.

Placidozoa span the normal to the bizarre

Placidozoans were first introduced in 2013 [168] as a closely related group to Bikosia [59, 60, 168], consisting of several groups of heterotrophic flagellates from the MAST-3 or Nanomonadea (including the parasitic Solenicola setigera [198] and the marine uniflagellated heterotroph Incisomonas marina [168]) as well as strictly intestinal protists in Opalinata. Placididea were recognized as a separate group in 2002 [199] with two taxa Wobblia lunata [62] and Placidia cafeteriopsis [199], but the number of described taxa is growing (e.g. [200]). Placidideans are often isolated from the deep sea halophilic environments, but in cultured conditions, they can tolerate lower levels of salinity [201, 202]. Their kidney-shaped cells, ultrastructural features (presence of mastigonemes on anterior flagellum), and movement all resemble Cafeteria sp. [201]. However, despite continuous effort, unlike Cafeteria, they are difficult to cultivate and remain mostly known from sequence data [60].

Opalinids are rather unusual protists, as all known taxa are intestinal commensals. Their cells are large (up to 3 mm) with multiple nuclei (2–200) and flagella or shorter cilia [203]. It is not surprising that based on these ciliate-like features, they were originally misclassified as ciliates [204]. They are mostly known to inhabit the cloacae of amphibians and lizards (e.g. Opalina sp. [205, 206]). Due to their obscure life style, opalinids are very under-sampled and their inner phylogenetic relationships are not resolved [207]. The related vertebrate intestinal commensal Blastocystis sp. [11, 162] is unlike its more decorated cousins. Its cells are spherical with a large central vacuole and one or two nuclei and lack flagella completely [208]. With extreme morphological plasticity and lifecycle complexity, this group can reveal how radiations can fill several available niches; however, due to the lack of a free-living heterotrophic model, our understanding is largely limited to phylogenies and comparative morphology.

Conclusion

Why should we care about free-living heterotrophic stramenopiles? Heterotrophic stramenopiles are among the most abundant heterotrophs in nearly every habitat, likely playing vital roles in every ecosystem. However, our limited understanding of their life strategies hinders our assessment of their ecological importance and evolutionary impacts. By delving into their cell biology, we can unravel transitions to autotrophy, osmotrophy, and parasitism. Furthermore, as our knowledge grows, we can better integrate these organisms into complex food webs, illuminating their ecological significance.

Apart from ochrophytes, all other photosynthetic eukaryotes branch sister to extremely derived heterotrophs. Thus, stramenopiles are possibly among the best candidates to study the entire evolutionary journey of the plastid—from its acquisition, genomic and metabolic assimilation, to its potential loss. The next steps towards better understanding the process of higher-order endosymbioses would be to establish model heterotrophic stramenopiles to determine which endogenous pathways might be recruited to support an incoming plastid. Whatever the future holds, a better understanding of heterotrophic stramenopiles will help determine their pivotal roles in ecosystems and their evolutionary dynamics.

Conflicts of interest

None declared.

Funding

This work was funded by National Science Foundation grants (2119963 and 2405455), Gordon and Betty Moore Foundation grant (GBMF9201) and VEDA FELLOWSHIPS within the Operational program Jan Amos Komensky (OP JAK), https://opjak.cz/en/ Project No. CZ.02.01.01/00/22_010/0008117.

Data availability

Data sharing is not applicable to this article as no datasets were generated or analyzed during the current study.
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References

1. Lefèvre  E, Roussel  B, Amblard  C  et al.  The molecular diversity of freshwater picoeukaryotes reveals high occurrence of putative parasitoids in the plankton. PLoS One  2008;3 :e2324. 10.1371/journal.pone.0002324 18545660
2. Singer  D, Seppey  CVW, Lentendu  G  et al.  Protist taxonomic and functional diversity in soil, freshwater and marine ecosystems. Environ Int  2021;146 :106262. 10.1016/j.envint.2020.106262 33221595
3. Seeleuthner  Y, Mondy  S, Lombard  V  et al.  Single-cell genomics of multiple uncultured stramenopiles reveals underestimated functional diversity across oceans. Nat Commun  2018;9 :310. 10.1038/s41467-017-02235-3 29358710
4. Massana  R, del Campo  J, Sieracki  ME  et al.  Exploring the uncultured microeukaryote majority in the oceans: reevaluation of ribogroups within stramenopiles. ISME J  2014;8 :854–66. 10.1038/ismej.2013.204 24196325
5. Thaler  M, Lovejoy  C. Environmental selection of marine stramenopile clades in the Arctic Ocean and coastal waters. Polar Biol  2014;37 :347–57. 10.1007/s00300-013-1435-0
6. Xu  Z, Wang  M, Wu  W  et al.  Vertical distribution of microbial eukaryotes from surface to the hadal zone of the Mariana trench. Front Microbiol  2018;9 :2023. 10.3389/fmicb.2018.02023 30210485
7. Schoenle  A, Hohlfeld  M, Rosse  M  et al.  Global comparison of bicosoecid Cafeteria-like flagellates from the deep ocean and surface waters, with reorganization of the family Cafeteriaceae. Eur J Protistol  2020;73 :125665. 10.1016/j.ejop.2019.125665 31978633
8. Park  JS, Cho  BC, Simpson  AGB. Halocafeteria seosinensis gen. Et sp. nov. (Bicosoecida), a halophilic bacterivorous nanoflagellate isolated from a solar saltern. Extremophiles  2006;10 :493–504. 10.1007/s00792-006-0001-x 16874468
9. Schoenle  A, Hohlfeld  M, Rybarski  A  et al.  Cafeteria in extreme environments: investigations on C. burkhardae and three new species from the Atacama Desert and the deep ocean. Eur J Protistol  2022;85 :125905. 10.1016/j.ejop.2022.125905 35868212
10. Haas  BJ, Kamoun  S, Zody  MC  et al.  Genome sequence and analysis of the Irish potato famine pathogen Phytophthora infestans. Nature  2009;461 :393–8. 10.1038/nature08358 19741609
11. Lhotská  Z, Jirků  M, Hložková  O  et al.  A study on the prevalence and subtype diversity of the intestinal protist Blastocystis sp. in a gut-healthy human population in the Czech Republic. Front Cell Infect Microbiol  2020;10 :544335. 10.3389/fcimb.2020.544335 33123491
12. Benoiston  A-S, Ibarbalz  FM, Bittner  L  et al.  The evolution of diatoms and their biogeochemical functions. Philos Trans R Soc B Biol Sci  2017;372 :20160397. 10.1098/rstb.2016.0397
13. Nakano  S, Ishii  N, Manage  P  et al.  Trophic roles of heterotrophic nanoflagellates and ciliates among planktonic organisms in a hypereutrophic pond. Aquat Microb Ecol  1998;16 :153–61. 10.3354/ame016153
14. Sanders  RW . Protists: Flagellates and amoebae. In: Mehner T and Tockner K (eds), Encyclopedia of Inland Waters, 2nd edn. Elsevier, 2022, 630–8.
15. Kristiansen  J . Chrysophytes – Golden algae. In: Likens  G.E. (ed.), Plankton of Inland Waters, 1st edn. Elsevier, 2009, 123–9.
16. Oborník  M, Dorrell  RG, Tikhonenkov  DV. Editorial: mixotrophic, secondary heterotrophic, and parasitic algae. Front Plant Sci  2021;12 :798555. 10.3389/fpls.2021.798555 34899815
17. Ban  H, Sato  S, Yoshikawa  S  et al.  Genome analysis of Parmales, the sister group of diatoms, reveals the evolutionary specialization of diatoms from phago-mixotrophs to photoautotrophs. Commun Biol  2023;6 :697. 10.1038/s42003-023-05002-x 37420035
18. Obiol  A, Muhovic  I, Massana  R. Oceanic heterotrophic flagellates are dominated by a few widespread taxa. Limnol Oceanogr  2021;66 :4240–53. 10.1002/lno.11956
19. Delmont TO, Gaia  M, Hinsinger  DD  et al.  Functional repertoire convergence of distantly related eukaryotic plankton lineages abundant in the sunlit ocean. Cell Genomics  2022;2 :100123. 10.1016/j.xgen.2022.100123 36778897
20. Jamy  M, Biwer  C, Vaulot  D  et al.  Global patterns and rates of habitat transitions across the eukaryotic tree of life. Nat Ecol Evol  2022;6 :1458–70. 10.1038/s41559-022-01838-4 35927316
21. Poulsen  CS, Ekstrøm  CT, Aarestrup  FM  et al.  Library preparation and sequencing platform introduce bias in metagenomic-based characterizations of microbiomes. Microbiol Spectr  2022;10 :e0009022. 10.1128/spectrum.00090-22 35289669
22. McLaren  MR, Willis  AD, Callahan  BJ. Consistent and correctable bias in metagenomic sequencing experiments. elife  2019;8 :e46923. 10.7554/eLife.46923 31502536
23. Sunagawa  S, Acinas  SG, Bork  P  et al.  Tara Oceans: towards global ocean ecosystems biology. Nat Rev Microbiol  2020;18 :428–45. 10.1038/s41579-020-0364-5 32398798
24. Sibbald  SJ, Archibald  JM. More protist genomes needed. Nat Ecol Evol  2017;1 :0145. 10.1038/s41559-017-0145
25. Graupner  N, Jensen  M, Bock  C  et al.  Evolution of heterotrophy in chrysophytes as reflected by comparative transcriptomics. FEMS Microbiol Ecol  2018;94 :fiy039. 10.1093/femsec/fiy039 29518196
26. Hu  A, Meng  F, Tanentzap  AJ  et al.  Dark matter enhances interactions within both microbes and dissolved organic matter under global change. Environ Sci Technol  2023;57 :761–9. 10.1021/acs.est.2c05052 36516075
27. Cavalier-Smith  T, Chao  EE-Y. Phylogeny and megasystematics of phagotrophic heterokonts (kingdom Chromista). J Mol Evol  2006;62 :388–420.16557340
28. Petersen  JB . Beiträge zur Kenntnis der Flagellatengeißeln. Bot Tidsskr  1929;40 :373–89.
29. Dodge  JD . Flagella and associated structures. In: Dodge  JD (ed.), The Fine Structure of Algal Cells, 1st edn. Academic Press Inc, London and New York. 1973, 57–79.
30. South  GR, Whittick  A. An Introduction to Phycology, 1st edn. Wiley, 1988, Hoboken, NJ.
31. Shukla  SK, Mohan  R, Sudhakar  M. Diatoms: a potential tool to understand past oceanographic settings. Curr Sci  2009;97 :1726–34.
32. Jirsová  D, Füssy  Z, Richtová  J  et al.  Morphology, ultrastructure, and mitochondrial genome of the marine non-photosynthetic bicosoecid Cafileria marina gen. et sp. nov. Microorganisms  2019;7 :240. 10.3390/microorganisms7080240 31387253
33. Santore  UJ . Flagellar and body scales in the Cryptophyceae. Br Phycol J  1983;18 :239–48. 10.1080/00071618300650251
34. Brooker  BE . Mastigonemes in a bodonid flagellate. Exp Cell Res  1965;37 :300–5. 10.1016/0014-4827(65)90178-3 14298942
35. Fu  G, Nagasato  C, Oka  S  et al.  Proteomics analysis of heterogeneous flagella in brown algae (Stramenopiles). Protist  2014;165 :662–75. 10.1016/j.protis.2014.07.007 25150613
36. Walker  CA, van West  P. Zoospore development in the oomycetes. Fungal Biol Rev  2007;21 :10–8. 10.1016/j.fbr.2007.02.001
37. Iwata  I, Kimura  K, Tomaru  Y  et al.  Bothrosome formation in Schizochytrium aggregatum (Labyrinthulomycetes, Stramenopiles) during zoospore settlement. Protist  2017;168 :206–19. 10.1016/j.protis.2016.12.002 28314190
38. Nakamura  S, Tanaka  G, Maeda  T  et al.  Assembly and function of Chlamydomonas flagellar mastigonemes as probed with a monoclonal antibody. J Cell Sci  1996;109 :57–62. 10.1242/jcs.109.1.57 8834790
39. Liu  P, Lou  X, Wingfield  JL  et al.  Chlamydomonas PKD2 organizes mastigonemes, hair-like glycoprotein polymers on cilia. J Cell Biol  2020;219 :e202001122. 10.1083/jcb.202001122 32348466
40. Hee  WY, Blackman  LM, Hardham  AR. Characterisation of Stramenopile-specific mastigoneme proteins in Phytophthora parasitica. Protoplasma  2019;256 :521–35. 10.1007/s00709-018-1314-1 30302550
41. Sengupta  S, Yang  X, Higgs  PG. The mechanisms of codon reassignments in mitochondrial genetic codes. J Mol Evol  2007;64 :662–88. 10.1007/s00239-006-0284-7 17541678
42. Liaud  MF, Lichtlé  C, Apt  K  et al.  Compartment-specific isoforms of TPI and GAPDH are imported into diatom mitochondria as a fusion protein: evidence in favor of a mitochondrial origin of the eukaryotic glycolytic pathway. Mol Biol Evol  2000;17 :213–23. 10.1093/oxfordjournals.molbev.a026301 10677844
43. Nakayama  T, Ishida  K, Archibald  JM. Broad distribution of TPI-GAPDH fusion proteins among eukaryotes: evidence for glycolytic reactions in the mitochondrion?  PLoS One  2012;7 :e52340. 10.1371/journal.pone.0052340 23284996
44. Río Bártulos  C, Rogers  MB, Williams  TA  et al.  Mitochondrial glycolysis in a major lineage of eukaryotes. Genome Biol Evol  2018;10 :2310–25. 10.1093/gbe/evy164 30060189
45. Azuma  T, Pánek  T, Tice  AK  et al.  An enigmatic stramenopile sheds light on early evolution in ochrophyta plastid organellogenesis. Mol Biol Evol  2022;39 :msac065. 10.1093/molbev/msac065 35348760
46. Cho  A, Tikhonenkov  DV, Hehenberger  E  et al.  Monophyly of diverse Bigyromonadea and their impact on phylogenomic relationships within stramenopiles. Mol Phylogenet Evol  2022;171 :107468. 10.1016/j.ympev.2022.107468 35358688
47. Khanipour Roshan  S, Dumack  K, Bonkowski  M  et al.  Stramenopiles and Cercozoa dominate the heterotrophic protist community of biological soil crusts irrespective of edaphic factors. Pedobiologia - J Soil Ecol  2020;83 :150673. 10.1016/j.pedobi.2020.150673
48. Terpis  KX, Salomaki  ED, Barcytė  D  et al.  Multiple plastid losses within photosynthetic stramenopiles revealed by comprehensive phylogenomics. bioRxiv  2024; 2024.02.03.578753
49. Burki  F, Shalchian-Tabrizi  K, Minge  M  et al.  Phylogenomics reshuffles the eukaryotic supergroups. PLoS One  2007;2 :e790. 10.1371/journal.pone.0000790 17726520
50. Burki  F, Roger  AJ, Brown  MW  et al.  The new tree of eukaryotes. Trends Ecol Evol  2020;35 :43–55. 10.1016/j.tree.2019.08.008 31606140
51. Burki  F, Inagaki  Y, Bråte  J  et al.  Large-scale phylogenomic analyses reveal that two enigmatic protist lineages, Telonemia and Centroheliozoa, are related to photosynthetic Chromalveolates. Genome Biol Evol  2009;1 :231–8. 10.1093/gbe/evp022 20333193
52. Hampl  V, Hug  L, Leigh  JW  et al.  Phylogenomic analyses support the monophyly of Excavata and resolve relationships among eukaryotic “supergroups”. Proc Natl Acad Sci  2009;106 :3859–64. 10.1073/pnas.0807880106 19237557
53. He  D, Sierra  R, Pawlowski  J  et al.  Reducing long-branch effects in multi-protein data uncovers a close relationship between Alveolata and Rhizaria. Mol Phylogenet Evol  2016;101 :1–7. 10.1016/j.ympev.2016.04.033 27132173
54. Sierra  R, Matz  MV, Aglyamova  G  et al.  Deep relationships of Rhizaria revealed by phylogenomics: a farewell to Haeckel’s Radiolaria. Mol Phylogenet Evol  2013;67 :53–9. 10.1016/j.ympev.2012.12.011 23280368
55. Sinha  SD, Wideman  JG. The persistent homology of mitochondrial ATP synthases. iScience  2023;26 :106700. 10.1016/j.isci.2023.106700 37250340
56. Patterson  DJ . Stramenopiles: Chromophytes from a protistan perspective. In: Green  JC, Leadbeater  BSC, Diver  WL (eds.), The Chromophyte Algae. Oxford University Press, Oxford UK, 1990, 357–80.
57. Derelle  R, López-García  P, Timpano  H  et al.  A phylogenomic framework to study the diversity and evolution of stramenopiles (=heterokonts). Mol Biol Evol  2016;33 :2890–8. 10.1093/molbev/msw168 27512113
58. Riisberg  I, Orr  RJS, Kluge  R  et al.  Seven gene phylogeny of Heterokonts. Protist  2009;160 :191–204. 10.1016/j.protis.2008.11.004 19213601
59. Thakur  R, Shiratori  T, Ishida  K. Taxon-rich multigene phylogenetic analyses resolve the phylogenetic relationship among deep-branching Stramenopiles. Protist  2019;170 :125682. 10.1016/j.protis.2019.125682 31568885
60. Cho  A, Tikhonenkov  DV, Lax  G  et al.  Phylogenomic position of genetically diverse phagotrophic stramenopile flagellates in the sediment-associated MAST-6 lineage and a potentially halotolerant placididean. Mol Phylogenet Evol  2024;190 :107964. 10.1016/j.ympev.2023.107964 37951557
61. Cho  A, Lax  G, Keeling  PJ. Phylogenomic analyses of ochrophytes (stramenopiles) with an emphasis on neglected lineages. Mol Phylogenet Evol  2024;198 :108120. 10.1016/j.ympev.2024.108120 38852907
62. Moriya  M, Nakayama  T, Inouye  I. Ultrastructure and 18S rDNA sequence analysis of Wobblia lunata gen. et sp. nov., a new heterotrophic flagellate (Stramenopiles, Incertae Sedis). Protist  2000;151 :41–55. 10.1078/1434-4610-00006 10896132
63. Sekiguchi  H, Moriya  M, Nakayama  T  et al.  Vestigial chloroplasts in heterotrophic stramenopiles Pteridomonas danica and Ciliophrys infusionum (Dictyochophyceae). Protist  2002;153 :157–67. 10.1078/1434-4610-00094 12125757
64. Simon  M, Jardillier  L, Deschamps  P  et al.  Complex communities of small protists and unexpected occurrence of typical marine lineages in shallow freshwater systems. Environ Microbiol  2015;17 :3610–27. 10.1111/1462-2920.12591 25115943
65. Adl  SM, Bass  D, Lane  CE  et al.  Revisions to the classification, nomenclature, and diversity of eukaryotes. J Eukaryot Microbiol  2019;66 :4–119. 10.1111/jeu.12691 30257078
66. Sibbald  SJ, Archibald  JM. Genomic insights into plastid evolution. Genome Biol Evol  2020;12 :978–90. 10.1093/gbe/evaa096 32402068
67. Kamikawa  R, Moog  D, Zauner  S  et al.  A non-photosynthetic diatom reveals early steps of reductive evolution in plastids. Mol Biol Evol  2017;34 :2355–66. 10.1093/molbev/msx172 28549159
68. Kamikawa  R, Azuma  T, Ishii  K  et al.  Diversity of organellar genomes in non-photosynthetic diatoms. Protist  2018;169 :351–61. 10.1016/j.protis.2018.04.009 29803116
69. Dorrell  RG, Azuma  T, Nomura  M  et al.  Principles of plastid reductive evolution illuminated by nonphotosynthetic chrysophytes. Proc Natl Acad Sci  2019;116 :6914–23. 10.1073/pnas.1819976116 30872488
70. Kayama  M, Maciszewski  K, Yabuki  A  et al.  Highly reduced plastid genomes of the non-photosynthetic dictyochophyceans Pteridomonas spp. (Ochrophyta, SAR) are retained for tRNA-Glu-based organellar heme biosynthesis. Front Plant Sci  2020;11 :602455. 10.3389/fpls.2020.602455 33329672
71. Barcytė  D, Jaške  K, Pánek  T  et al.  The net-like heterotrophic amoeba Leukarachnion salinum sp. nov. (Ochrophyta, Stramenopiles) has a cryptic plastid. bioRxiv  2022;2003–5.
72. McKie-Krisberg  ZM, Sanders  RW, Gast  RJ. Evaluation of mixotrophy-associated gene expression in two species of polar marine algae. Front Mar Sci  2018;5 :273. 10.3389/fmars.2018.00273
73. Li  M, Chen  Y, Zhang  F  et al.  A three-dimensional mixotrophic model of Karlodinium veneficum blooms for a eutrophic estuary. Harmful Algae  2022;113 :102203. 10.1016/j.hal.2022.102203 35287934
74. Koppelle  S, López-Escardó  D, Brussaard  CPD  et al.  Mixotrophy in the bloom-forming genus Phaeocystis and other haptophytes. Harmful Algae  2022;117 :102292. 10.1016/j.hal.2022.102292 35944956
75. Ptacnik  R, Gomes  A, Royer  S-J  et al.  A light-induced shortcut in the planktonic microbial loop. Sci Rep  2016;6 :29286. 10.1038/srep29286 27404551
76. Wilken  S, Yung  CCM, Hamilton  M  et al.  The need to account for cell biology in characterizing predatory mixotrophs in aquatic environments. Philos Trans R Soc B Biol Sci  2019;374 :20190090. 10.1098/rstb.2019.0090
77. Barbaglia  GS, Paight  C, Honig  M  et al.  Environment-dependent metabolic investments in the mixotrophic chrysophyte Ochromonas. J Phycol  2024;60 :170–84. 10.1111/jpy.13418 38141034
78. Green  BR . After the primary endosymbiosis: an update on the chromalveolate hypothesis and the origins of algae with Chl c. Photosynth Res  2011;107 :103–15. 10.1007/s11120-010-9584-2 20676772
79. Streckaite  S, Gardian  Z, Li  F  et al.  Pigment configuration in the light-harvesting protein of the xanthophyte alga Xanthonema debile. Photosynth Res  2018;138 :139–48. 10.1007/s11120-018-0557-1 30006883
80. Jeffrey  S, Wright  SW, Zapata  M. Microalgal classes and their signature pigments. In: Roy  S, Llewellyn  C, Skarstad  E, Johnsen  G (eds). Phytoplankton Pigments: Characterization, Chemotaxonomy and Applications in Oceanography. 2011. Cambridge University Press, Cambridge, UK. pp. 3–77.
81. Pierella Karlusich  JJ, Bowler  C, Biswas  H. Carbon dioxide concentration mechanisms in natural populations of marine diatoms: insights from Tara Oceans. Front Plant Sci  2021;12 :657821. 10.3389/fpls.2021.657821 33995455
82. Tokushima  H, Inoue-Kashino  N, Nakazato  Y  et al.  Advantageous characteristics of the diatom Chaetoceros gracilis as a sustainable biofuel producer. Biotechnol Biofuels  2016;9 :235. 10.1186/s13068-016-0649-0 27822308
83. Zeni  V, Baliota  GV, Benelli  G  et al.  Diatomaceous earth for arthropod pest control: back to the future. Molecules  2021;26 :7487. 10.3390/molecules26247487 34946567
84. Piotrowski  K, Romanowska-Duda  Z, Messyasz  B. Cultivation of energy crops by ecological methods under the conditions of global climate and environmental changes with the use of diatom extract as a natural source of chemical compounds. Acta Physiol Plant  2020;42 :146. 10.1007/s11738-020-03135-8
85. Kroth  PG, Bones  AM, Daboussi  F  et al.  Genome editing in diatoms: achievements and goals. Plant Cell Rep  2018;37 :1401–8. 10.1007/s00299-018-2334-1 30167805
86. Huang  W, Daboussi  F. Genetic and metabolic engineering in diatoms. Philos Trans R Soc B Biol Sci  2017;372 :20160411. 10.1098/rstb.2016.0411
87. Vergés  A, Campbell  AH. Kelp forests. Curr Biol  2020;30 :R919–20. 10.1016/j.cub.2020.06.053 32810446
88. Smale  DA . Impacts of ocean warming on kelp forest ecosystems. New Phytol  2020;225 :1447–54. 10.1111/nph.16107 31400287
89. Neushul  M . Studies on the giant kelp, macrocystis.II.Reproduction. Am J Bot  1963;50 :354–9. 10.1002/j.1537-2197.1963.tb07203.x
90. B-Béres  V, Stenger-Kovács  C, Buczkó  K  et al.  Ecosystem services provided by freshwater and marine diatoms. Hydrobiologia  2023;850 :2707–33. 10.1007/s10750-022-04984-9
91. Kim  MJ, Yun  HY, Shin  K-H  et al.  Evaluation of food web structure and complexity in the process of kelp bed recovery using stable isotope analysis. Front Mar Sci  2022;9 :885676. 10.3389/fmars.2022.885676
92. Taucher  J, Bach  LT, Prowe  AEF  et al.  Enhanced silica export in a future ocean triggers global diatom decline. Nature  2022;605 :696–700. 10.1038/s41586-022-04687-0 35614245
93. Pernice  MC, Giner  CR, Logares  R  et al.  Large variability of bathypelagic microbial eukaryotic communities across the world’s oceans. ISME J  2016;10 :945–58. 10.1038/ismej.2015.170 26451501
94. Nicholls  KH, Wujek  DE. Chrysophyceae and Phaeothamniophyceae. In: Wehr  JD, Sheath  RG, Kociolek  JP (eds.), Freshwater Algae of North America, 2nd edn. Academic Press Inc Hoboken, NJ, 2015, 537–86.
95. Lie  AAY, Liu  Z, Terrado  R  et al.  A tale of two mixotrophic chrysophytes: insights into the metabolisms of two Ochromonas species (Chrysophyceae) through a comparison of gene expression. PLoS One  2018;13 :e0192439. 10.1371/journal.pone.0192439 29438384
96. Wilken  S, Choi  CJ, Worden  AZ. Contrasting mixotrophic lifestyles reveal different ecological niches in two closely related marine protists. J Phycol  2020;56 :52–67. 10.1111/jpy.12920 31529498
97. Holen  DA, Boraas  ME. The feeding behavior of Spumella sp. as a function of particle size: implications for bacterial size in pelagic systems. Hydrobiologia  1991;220 :73–88. 10.1007/BF00017493
98. Meyer  N, Rydzyk  A, Pohnert  G. Pronounced uptake and metabolism of organic substrates by diatoms revealed by pulse-labeling metabolomics. Front Mar Sci  2022;9 :821167. 10.3389/fmars.2022.821167
99. Mitra  A, Caron  DA, Faure  E  et al.  The Mixoplankton Database (MDB): diversity of photo-phago-trophic plankton in form, function, and distribution across the global ocean. J Eukaryot Microbiol  2023;70 :e12972. 10.1111/jeu.12972 36847544
100. Godrijan  J, Drapeau  D, Balch  WM. Mixotrophic uptake of organic compounds by coccolithophores. Limnol Oceanogr  2020;65 :1410–21. 10.1002/lno.11396
101. Balch  WM, Drapeau  DT, Poulton  N  et al.  Osmotrophy of dissolved organic compounds by coccolithophore populations: fixation into particulate organic and inorganic carbon. Sci Adv  2023;9 :eadf6973. 10.1126/sciadv.adf6973 37224255
102. Kamikawa  R, Mochizuki  T, Sakamoto  M  et al.  Genome evolution of a nonparasitic secondary heterotroph, the diatom Nitzschia putrida. Sci Adv  2022;8 :eabi5075. 10.1126/sciadv.abi5075 35486731
103. Maberly  SC, Ball  LA, Raven  JA  et al.  Inorganic carbon acquisition by chrysophytes. J Phycol  2009;45 :1052–61. 10.1111/j.1529-8817.2009.00734.x 27032350
104. Terrado  R, Pasulka  AL, Lie  AAY  et al.  Autotrophic and heterotrophic acquisition of carbon and nitrogen by a mixotrophic chrysophyte established through stable isotope analysis. ISME J  2017;11 :2022–34. 10.1038/ismej.2017.68 28524870
105. Terrado  R, Monier  A, Edgar  R  et al.  Diversity of nitrogen assimilation pathways among microbial photosynthetic eukaryotes. J Phycol  2015;51 :490–506. 10.1111/jpy.12292 26986665
106. Kamjunke  N, Henrichs  T, Gaedke  U. Phosphorus gain by bacterivory promotes the mixotrophic flagellate Dinobryon spp. during re-oligotrophication. J Plankton Res  2006;29 :39–46. 10.1093/plankt/fbl054
107. Rothhaupt  KO . Utilization of substitutable carbon and phosphorus sources by the mixotrophic chrysophyte Ochromonas sp. Ecology  1996;77 :706–15. 10.2307/2265495
108. Johnson  WM, Alexander  H, Bier  RL  et al.  Auxotrophic interactions: a stabilizing attribute of aquatic microbial communities?  FEMS Microbiol Ecol  2020;96 :fiaa115. 10.1093/femsec/fiaa115 32520336
109. Faure  E, Not  F, Benoiston  A-S  et al.  Mixotrophic protists display contrasted biogeographies in the global ocean. ISME J  2019;13 :1072–83. 10.1038/s41396-018-0340-5 30643201
110. Leles  SG, Mitra  A, Flynn  KJ  et al.  Oceanic protists with different forms of acquired phototrophy display contrasting biogeographies and abundance. Proc R Soc B Biol Sci  2017;284 :20170664. 10.1098/rspb.2017.0664
111. Tanaka  R, Tanaka  A. Tetrapyrrole biosynthesis in higher plants. Annu Rev Plant Biol  2007;58 :321–46. 10.1146/annurev.arplant.57.032905.105448 17227226
112. López  G, Yate  C, Ramos  FA  et al.  Production of polyunsaturated fatty acids and lipids from autotrophic, mixotrophic and heterotrophic cultivation of Galdieria sp. strain USBA-GBX-832. Sci Rep  2019;9 :10791. 10.1038/s41598-019-46645-3 31346188
113. Boëchat  IG, Weithoff  G, Krüger  A  et al.  A biochemical explanation for the success of mixotrophy in the flagellate Ochromonas sp. Limnol Oceanogr  2007;52 :1624–32. 10.4319/lo.2007.52.4.1624
114. Brown  JW, Sorhannus  U. A molecular genetic timescale for the diversification of autotrophic stramenopiles (Ochrophyta): substantive underestimation of putative fossil ages. PLoS One  2010;5 :e12759. 10.1371/journal.pone.0012759 20862282
115. Burki  F, Kaplan  M, Tikhonenkov  DV  et al.  Untangling the early diversification of eukaryotes: a phylogenomic study of the evolutionary origins of Centrohelida, Haptophyta and Cryptista. Proc R Soc B Biol Sci  2016;283 :20152802. 10.1098/rspb.2015.2802
116. Keeling  PJ . Chromalveolates and the evolution of plastids by secondary endosymbiosis. J Eukaryot Microbiol  2009;56 :1–8. 10.1111/j.1550-7408.2008.00371.x 19335769
117. Ševčíková  T, Horák  A, Klimeš  V  et al.  Updating algal evolutionary relationships through plastid genome sequencing: did alveolate plastids emerge through endosymbiosis of an ochrophyte?  Sci Rep  2015;5 :10134. 10.1038/srep10134 26017773
118. Cavalier-Smith  T . Principles of protein and lipid targeting in secondary symbiogenesis: euglenoid, dinoflagellate, and sporozoan plastid origins and the eukaryote family tree. J Eukaryot Microbiol  1999;46 :347–66. 10.1111/j.1550-7408.1999.tb04614.x 18092388
119. Khan  H, Parks  N, Kozera  C  et al.  Plastid genome sequence of the cryptophyte alga Rhodomonas salina CCMP1319: lateral transfer of putative DNA replication machinery and a test of chromist plastid phylogeny. Mol Biol Evol  2007;24 :1832–42. 10.1093/molbev/msm101 17522086
120. Dorrell  RG, Gile  G, McCallum  G  et al.  Chimeric origins of ochrophytes and haptophytes revealed through an ancient plastid proteome. elife  2017;6 :e23717. 10.7554/eLife.23717 28498102
121. Baurain  D, Brinkmann  H, Petersen  J  et al.  Phylogenomic evidence for separate acquisition of plastids in cryptophytes, haptophytes, and stramenopiles. Mol Biol Evol  2010;27 :1698–709. 10.1093/molbev/msq059 20194427
122. Yoon  HS, Hackett  JD, Pinto  G  et al.  The single, ancient origin of chromist plastids. Proc Natl Acad Sci  2002;99 :15507–12. 10.1073/pnas.242379899 12438651
123. Fast  NM, Kissinger  JC, Roos  DS  et al.  Nuclear-encoded, plastid-targeted genes suggest a single common origin for apicomplexan and dinoflagellate plastids. Mol Biol Evol  2001;18 :418–26. 10.1093/oxfordjournals.molbev.a003818 11230543
124. Tyler  BM, Tripathy  S, Zhang  X  et al.  Phytophthora genome sequences uncover evolutionary origins and mechanisms of pathogenesis. Science  2006;313 :1261–6. 10.1126/science.1128796 16946064
125. Sargent  JR, Bell  M V., Henderson  RJ. Protists as sources of (n-3) polyunsaturated fatty acids for vertebrate development. In: Brugerolle  G, Mignot  JP (eds). Proceedings of the Second European Congress of Protistology. 1995. Clermont-Ferrand, pp. 54–64.
126. Harper  JT, Keeling  PJ. Nucleus-encoded, plastid-targeted glyceraldehyde-3-phosphate dehydrogenase (GAPDH) indicates a single origin for chromalveolate plastids. Mol Biol Evol  2003;20 :1730–5. 10.1093/molbev/msg195 12885964
127. Wang  Q, Sun  H, Huang  J. Re-analyses of “algal” genes suggest a complex evolutionary history of oomycetes. Front Plant Sci  2017;8 :1540. 10.3389/fpls.2017.01540 28932232
128. Santos  HJ, Nozaki  T. The mitosome of the anaerobic parasitic protist Entamoeba histolytica: a peculiar and minimalist mitochondrion-related organelle. J Eukaryot Microbiol  2022;69 :e12923. 10.1111/jeu.12923 35588086
129. Muñoz-Gómez  SA . Energetics and evolution of anaerobic microbial eukaryotes. Nat Microbiol  2023;8 :197–203. 10.1038/s41564-022-01299-2 36646908
130. Karnkowska  A, Vacek  V, Zubáčová  Z  et al.  A eukaryote without a mitochondrial organelle. Curr Biol  2016;26 :1274–84. 10.1016/j.cub.2016.03.053 27185558
131. Hjort  K, Goldberg  AV, Tsaousis  AD  et al.  Diversity and reductive evolution of mitochondria among microbial eukaryotes. Philos Trans R Soc B Biol Sci  2010;365 :713–27. 10.1098/rstb.2009.0224
132. Salomaki  E, Kolisko  M. There is treasure everywhere: reductive plastid evolution in apicomplexa in light of their close relatives. Biomol Ther  2019;9 :378. 10.3390/biom9080378
133. Makiuchi  T, Nozaki  T. Highly divergent mitochondrion-related organelles in anaerobic parasitic protozoa. Biochimie  2014;100 :3–17. 10.1016/j.biochi.2013.11.018 24316280
134. Zhu  G, Marchewka  MJ, Keithly  JS. Cryptosporidium parvum appears to lack a plastid genome. Microbiology  2000;146 :315–21. 10.1099/00221287-146-2-315 10708370
135. Gornik  SG, Febrimarsa  CAM, MacRae  JI  et al.  Endosymbiosis undone by stepwise elimination of the plastid in a parasitic dinoflagellate. Proc Natl Acad Sci USA  2015;112 :5767–72. 10.1073/pnas.1423400112 25902514
136. Schön  ME, Zlatogursky  VV, Singh  RP  et al.  Single cell genomics reveals plastid-lacking Picozoa are close relatives of red algae. Nat Commun  2021;12 :6651. 10.1038/s41467-021-26918-0 34789758
137. Burki  F . The eukaryotic tree of life from a global phylogenomic perspective. Cold Spring Harb Perspect Biol  2014;6 :a016147–7. 10.1101/cshperspect.a016147 24789819
138. Adl  SM, Simpson  AGB, Lane  CE  et al.  The revised classification of eukaryotes. J Eukaryot Microbiol  2012;59 :429–514. 10.1111/j.1550-7408.2012.00644.x 23020233
139. Petersen  J, Ludewig  A-K, Michael  V  et al.  Chromera velia, endosymbioses and the rhodoplex hypothesis—plastid evolution in cryptophytes, alveolates, stramenopiles, and haptophytes (CASH lineages). Genome Biol Evol  2014;6 :666–84. 10.1093/gbe/evu043 24572015
140. Hempel  F, Bullmann  L, Lau  J  et al.  ERAD-derived preprotein transport across the second outermost plastid membrane of diatoms. Mol Biol Evol  2009;26 :1781–90. 10.1093/molbev/msp079 19377060
141. Felsner  G, Sommer  MS, Gruenheit  N  et al.  ERAD components in organisms with complex red plastids suggest recruitment of a preexisting protein transport pathway for the periplastid membrane. Genome Biol Evol  2011;3 :140–50. 10.1093/gbe/evq074 21081314
142. Kienle  N, Kloepper  TH, Fasshauer  D. Shedding light on the expansion and diversification of the Cdc48 protein family during the rise of the eukaryotic cell. BMC Evol Biol  2016;16 :215. 10.1186/s12862-016-0790-1 27756227
143. Sommer  MS, Gould  SB, Lehmann  P  et al.  Der1-mediated preprotein import into the periplastid compartment of chromalveolates?  Mol Biol Evol  2007;24 :918–28. 10.1093/molbev/msm008 17244602
144. Bolte  K, Gruenheit  N, Felsner  G  et al.  Making new out of old: recycling and modification of an ancient protein translocation system during eukaryotic evolution. BioEssays  2011;33 :368–76. 10.1002/bies.201100007 21425305
145. Stiller  JW, Schreiber  J, Yue  J  et al.  The evolution of photosynthesis in chromist algae through serial endosymbioses. Nat Commun  2014;5 :5764. 10.1038/ncomms6764 25493338
146. Bodył  A, Stiller  JW, Mackiewicz  P. Chromalveolate plastids: direct descent or multiple endosymbioses?  Trends Ecol Evol  2009;24 :119–21.19200617
147. Strassert  JFH, Irisarri  I, Williams  TA  et al.  A molecular timescale for eukaryote evolution with implications for the origin of red algal-derived plastids. Nat Commun  2021;12 :1879. 10.1038/s41467-021-22044-z 33767194
148. Kim  JI, Moore  CE, Archibald  JM  et al.  Evolutionary dynamics of cryptophyte plastid genomes. Genome Biol Evol  2017;9 :1859–72. 10.1093/gbe/evx123 28854597
149. Karnkowska  A, Yubuki  N, Maruyama  M  et al.  Euglenozoan kleptoplasty illuminates the early evolution of photoendosymbiosis. Proc Natl Acad Sci  2023;120 :e2220100120. 10.1073/pnas.2220100120 36927158
150. Shiratori  T, Nakayama  T, Ishida  K. A new deep-branching stramenopile, Platysulcus tardus gen. Nov., sp. nov. Protist  2015;166 :337–48. 10.1016/j.protis.2015.05.001 26070192
151. Leonard  G, Labarre  A, Milner  DS  et al.  Comparative genomic analysis of the ‘pseudofungus’ Hyphochytrium catenoides. Open Biol  2018;8 :170184. 10.1098/rsob.170184 29321239
152. Spanu  PD, Panstruga  R. Editorial: biotrophic plant-microbe interactions. Front Plant Sci  2017;8 :192.28243250
153. Martin  F, Kohler  A, Murat  C  et al.  Unearthing the roots of ectomycorrhizal symbioses. Nat Rev Microbiol  2016;14 :760–73. 10.1038/nrmicro.2016.149 27795567
154. Van der Auwera  G, De Baere  R, Van de Peer  Y  et al.  The phylogeny of the Hyphochytriomycota as deduced from ribosomal RNA sequences of Hyphochytrium catenoides. Mol Biol Evol  1995;12 :671–8.7659021
155. Lévesque  CA . Fifty years of oomycetes—from consolidation to evolutionary and genomic exploration. Fungal Divers  2011;50 :35–46. 10.1007/s13225-011-0128-7
156. Lévesque  CA, Brouwer  H, Cano  L  et al.  Genome sequence of the necrotrophic plant pathogen Pythium ultimum reveals original pathogenicity mechanisms and effector repertoire. Genome Biol  2010;11 :R73. 10.1186/gb-2010-11-7-r73 20626842
157. Richards  TA, Soanes  DM, Jones  MDM  et al.  Horizontal gene transfer facilitated the evolution of plant parasitic mechanisms in the oomycetes. Proc Natl Acad Sci  2011;108 :15258–63. 10.1073/pnas.1105100108 21878562
158. Savory  F, Leonard  G, Richards  TA. The role of horizontal gene transfer in the evolution of the oomycetes. PLoS Pathog  2015;11 :e1004805. 10.1371/journal.ppat.1004805 26020232
159. Aleoshin  VV, Mylnikov  AP, Mirzaeva  GS  et al.  Heterokont predator Develorapax marinus gen. et sp. nov. – a model of the ochrophyte ancestor. Front Microbiol  2016;7 :1194.27536283
160. Leipe  DD, Tong  SM, Goggin  CL  et al.  16S-like rDNA sequences from Developayella elegans, Labyrinthuloides haliotidis, and Proteromonas lacertae confirm that the stramenopiles are a primarily heterotrophic group. Eur J Protistol  1996;32 :449–58. 10.1016/S0932-4739(96)80004-6
161. Weiler  BA, Elisabet  LS, Sieracki  ME  et al.  Mediocremonas mediterraneus, a new member within the developea. J Eukaryot Microbiol  2021;68 :12825. 10.1111/jeu.12825
162. Silberman  JD, Sogin  ML, Leipe  DD  et al.  Human parasite finds taxonomic home. Nature  1996;380 :398–8. 10.1038/380398a0 8602239
163. Bahnweg  G, Sparrow  FK. Aplanochytrium kerguelensis gen. nov. spec. nov., a new phycomycete from subantarctic marine waters. Arch Mikrobiol  1972;81 :45–9. 10.1007/BF00715023 5058877
164. Yubuki  N, Pánek  T, Yabuki  A  et al.  Morphological identities of two different marine stramenopile environmental sequence clades: Bicosoeca kenaiensis (Hilliard, 1971) and Cantina marsupialis (Larsen and Patterson, 1990) gen. nov., comb. nov. J Eukaryot Microbiol  2015;62 :532–42. 10.1111/jeu.12207 25594562
165. Labarre  A, López-Escardó  D, Latorre  F  et al.  Comparative genomics reveals new functional insights in uncultured MAST species. ISME J  2021;15 :1767–81. 10.1038/s41396-020-00885-8 33452482
166. de Vargas  C, Audic  S, Henry  N  et al.  Eukaryotic plankton diversity in the sunlit ocean. Science  2015;348 :1261605. 10.1126/science.1261605 25999516
167. Wideman  JG, Monier  A, Rodríguez-Martínez  R  et al.  Unexpected mitochondrial genome diversity revealed by targeted single-cell genomics of heterotrophic flagellated protists. Nat Microbiol  2019;5 :154–65. 10.1038/s41564-019-0605-4 31768028
168. Cavalier-Smith  T, Scoble  JM. Phylogeny of Heterokonta: Incisomonas marina, a uniciliate gliding opalozoan related to Solenicola (Nanomonadea), and evidence that Actinophryida evolved from raphidophytes. Eur J Protistol  2013;49 :328–53. 10.1016/j.ejop.2012.09.002 23219323
169. Shiratori  T, Thakur  R, Ishida  K. Pseudophyllomitus vesiculosus (Larsen and Patterson 1990) lee, 2002, a poorly studied phagotrophic biflagellate is the first characterized member of Stramenopile environmental clade MAST-6. Protist  2017;168 :439–51. 10.1016/j.protis.2017.06.004 28822908
170. Tsui  CKM, Marshall  W, Yokoyama  R  et al.  Labyrinthulomycetes phylogeny and its implications for the evolutionary loss of chloroplasts and gain of ectoplasmic gliding. Mol Phylogenet Evol  2009;50 :129–40. 10.1016/j.ympev.2008.09.027 18977305
171. Raghukumar  S, Damare  VS. Increasing evidence for the important role of Labyrinthulomycetes in marine ecosystems. Bot Mar  2011;54 :3–11. 10.1515/bot.2011.008
172. Xie  N, Hunt  DE, Johnson  ZI  et al.  Annual partitioning patterns of labyrinthulomycetes protists reveal their multifaceted role in marine microbial food webs. Appl Environ Microbiol  2021;87 :e01652–20. 10.1128/AEM.01652-20
173. Rubin  E, Tanguy  A, Pales Espinosa  E  et al.  Differential gene expression in five isolates of the clam pathogen, quahog parasite unknown (QPX). J Eukaryot Microbiol  2017;64 :647–54. 10.1111/jeu.12400 28171698
174. Xie  N, Wang  Z, Hunt  DE  et al.  Niche partitioning of Labyrinthulomycete protists across sharp coastal gradients and their putative relationships with bacteria and fungi. Front Microbiol  2022;13 :906864. 10.3389/fmicb.2022.906864 35685928
175. Hamamoto  Y, Honda  D. Nutritional intake of Aplanochytrium (Labyrinthulea, Stramenopiles) from living diatoms revealed by culture experiments suggesting the new prey–predator interactions in the grazing food web of the marine ecosystem. PLoS One  2019;14 :e0208941. 10.1371/journal.pone.0208941 30625142
176. Gomaa  F, Mitchell  EAD, Lara  E. Amphitremida (Poche, 1913) is a new major, ubiquitous labyrinthulomycete clade. PLoS One  2013;8 :e53046. 10.1371/journal.pone.0053046 23341921
177. Takahashi  Y, Yoshida  M, Inouye  I  et al.  Fibrophrys columna gen. nov., sp. nov: a member of the family Amphifilidae. Eur J Protistol  2016;56 :41–50. 10.1016/j.ejop.2016.06.003 27468745
178. Pan  J, del Campo  J, Keeling  PJ. Reference tree and environmental sequence diversity of Labyrinthulomycetes. J Eukaryot Microbiol  2017;64 :88–96. 10.1111/jeu.12342 27329779
179. Takahashi  Y, Yoshida  M, Inouye  I  et al.  Diplophrys mutabilis sp. nov., a new member of Labyrinthulomycetes from freshwater habitats. Protist  2014;165 :50–65. 10.1016/j.protis.2013.10.001 24334197
180. Qiu  X, Xie  X, Meesapyodsuk  D. Molecular mechanisms for biosynthesis and assembly of nutritionally important very long chain polyunsaturated fatty acids in microorganisms. Prog Lipid Res  2020;79 :101047. 10.1016/j.plipres.2020.101047 32540152
181. Ishibashi  Y, Goda  H, Hamaguchi  R  et al.  PUFA synthase-independent DHA synthesis pathway in Parietichytrium sp. and its modification to produce EPA and n-3DPA. Commun Biol  2021;4 :1378. 10.1038/s42003-021-02857-w 34887503
182. Sanders  RW, Porter  KG, Bennett  SJ  et al.  Seasonal patterns of bacterivory by flagellates, ciliates, rotifers, and cladocerans in a freshwater planktonic community. Limnol Oceanogr  1989;34 :673–87. 10.4319/lo.1989.34.4.0673
183. Hahn  MW, Höfle  MG. Grazing of protozoa and its effect on populations of aquatic bacteria. FEMS Microbiol Ecol  2001;35 :113–21. 10.1111/j.1574-6941.2001.tb00794.x 11295449
184. Harder  CB, Ekelund  F, Karpov  SA. Ultrastructure and phylogenetic position of regin rotiferus and Otto terricolus genera et species novae (Bicosoecida, Heterokonta/Stramenopiles). Protist  2014;165 :144–60. 10.1016/j.protis.2014.01.004 24637333
185. Moestrup  Ø.  Current status of chtysophyte ‘splinter groups’: Synurophytes, pedinellis, silicoflagellates. In: Sandgren  C, Smol  JP, Kristiansen  J (eds). Chrysophyte Algae: Ecology, Phylogeny and Development, 1st ed.  1995. Cambridge University Press, Cambridge, pp. 1535–5, 10.1017/CBO9780511752292.005.
186. Preisig  HR . A modern concept of chrysophyte classification. In: Sandgren  C, Smol  JP, Kristiansen  J (eds). Chrysophyte Algae: Ecology, Phylogeny, Development, 1st ed.  1995. Cambridge University Press, Cambridge, pp. 47–74, 10.1017/CBO9780511752292.004.
187. O’Kelly  CJ, Patterson  DJ. The flagellar apparatus of cafeteria roenbergensis Fenchel & Patterson, 1988 (Bicosoecales = Bicosoecida). Eur J Protistol  1996;32 :216–26. 10.1016/S0932-4739(96)80021-6
188. Millette  NC, Gast  RJ, Luo  JY  et al.  Mixoplankton and mixotrophy: future research priorities. J Plankton Res  2023;45 :576–96. 10.1093/plankt/fbad020 37483910
189. Fenchel  T, Patterson  DJ. Cafeteria roenbergensis nov. gen., nov. sp., a heterotrophic microflagellate from marine plankton. Mar Microb Food Webs  1988;3 :9–19.
190. Baricevic  A, Maric Pfannkuchen  D, Smodlaka Tankovic  M  et al.  Identification of the heterotrophic nanoflagellate Bilabrum latius in the southern Adriatic (Mediterranean Sea). Eur J Protistol  2023;90 :125999. 10.1016/j.ejop.2023.125999 37352685
191. Burger  G, O’Kelly  C, Gray  MW  et al.  Cafeteria Roenbergensis Mitochondrial DNA Complete Sequence OGMP Accession no. AF193903, 1999.
192. Hackl  T, Martin  R, Barenhoff  K  et al.  Four high-quality draft genome assemblies of the marine heterotrophic nanoflagellate cafeteria roenbergensis. Sci Data  2020;7 :29. 10.1038/s41597-020-0363-4 31964893
193. Boegnik  J, Matz  C, Jürgens  K  et al.  Confusing selective feeding with differential digestion in bacterivorous nanoflagellates. J Eukaryot Microbiol  2001;48 :425–32. 10.1111/j.1550-7408.2001.tb00175.x 11456318
194. Ishigaki  T, Terazaki  M. Grazing behavior of heterotrophic nanoflagellates observed with a high speed VTR system. J Eukaryot Microbiol  1998;45 :484–7. 10.1111/j.1550-7408.1998.tb05104.x
195. Jürgens  K, Massana  R. Protistan grazing on marine bacterioplankton. In: Kirchman  D.L. (ed.), Microbial Ecology of the Oceans, 2nd edn. John Wiley & Sons, Inc., Hoboken, NJ, 2008, 383–441.
196. Fischer  MG, Allen  MJ, Wilson  WH  et al.  Giant virus with a remarkable complement of genes infects marine zooplankton. Proc Natl Acad Sci USA  2010;107 :19508–13. 10.1073/pnas.1007615107 20974979
197. Massana  R, Del Campo  J, Dinter  C  et al.  Crash of a population of the marine heterotrophic flagellate cafeteria roenbergensis by viral infection. Environ Microbiol  2007;9 :2660–9. 10.1111/j.1462-2920.2007.01378.x 17922751
198. Gómez  F, Moreira  D, Benzerara  K  et al.  Solenicola setigera is the first characterized member of the abundant and cosmopolitan uncultured marine stramenopile group MAST-3. Environ Microbiol  2011;13 :193–202. 10.1111/j.1462-2920.2010.02320.x 20722698
199. Moriya  M, Nakayama  T, Inouye  I. A new class of the stramenopiles, placididea classis nova: description of Placidia cafeteriopsis gen. Et sp. nov. Protist  2002;153 :143–56. 10.1078/1434-4610-00093 12125756
200. Okamura  T, Kondo  R. Suigetsumonas clinomigrationis gen. Et sp. nov., a novel facultative anaerobic nanoflagellate isolated from the meromictic Lake Suigetsu, Japan. Protist  2015;166 :409–21. 10.1016/j.protis.2015.06.003 26202992
201. Park  JS, Simpson  AGB. Characterization of halotolerant Bicosoecida and Placididea (Stramenopila) that are distinct from marine forms, and the phylogenetic pattern of salinity preference in heterotrophic stramenopiles. Environ Microbiol  2010;12 :1173–84. 10.1111/j.1462-2920.2010.02158.x 20132281
202. Rybarski  AE, Nitsche  F, Soo Park  J  et al.  Revision of the phylogeny of Placididea (Stramenopiles): molecular and morphological diversity of novel placidid protists from extreme aquatic environments. Eur J Protistol  2021;81 :125809. 10.1016/j.ejop.2021.125809 34673437
203. Kostka  M . Opalinata. In: Archibald  J.M., AGB  S., Slamovits  C.H.  et al. (eds.), Handbook of the Protists, 2nd edn. Cham: Springer International Publishing, 2016, 1–23.
204. Corliss  JO . The opalinid infusorians: flagellates or ciliates?  J Protozool  1955;2 :107–14. 10.1111/j.1550-7408.1955.tb02410.x
205. Zhao  W, Hu  G, Ponce-Gordo  F  et al.  Morphological description of Opalina obtrigonoidea Metcalf, 1923 (Heterokonta, Opalinea) from Duttaphrynus melanostictus and evaluation of the ITS region as a suitable genetic marker for inter-species identification in Opalina. Parasitol Int  2020;76 :102103. 10.1016/j.parint.2020.102103 32169658
206. Wang  R, Zhao  W, Hu  G  et al.  Redescription of Opalina triangulata (Heterokonta, Opalinea) from Fejervarya limnocharis based on morphological and molecular data. Eur J Protistol  2019;71 :125639. 10.1016/j.ejop.2019.125639 31550629
207. Li  M, Hu  G, Zhao  W  et al.  A revised taxonomy and phylogeny of opalinids (Stramenopiles: Opalinata) inferred from the analysis of complete nuclear ribosomal DNA genes. Zool J Linnean Soc  2023;201 :269–89. 10.1093/zoolinnean/zlad150
208. Tan  KSW . New insights on classification, identification, and clinical relevance of Blastocystis spp. Clin Microbiol Rev  2008;21 :639–65. 10.1128/CMR.00022-08 18854485
209. Burki  F . The convoluted evolution of eukaryotes with complex plastids. Advances in Botanical Research, 2017;84 :1–30. 10.1016/bs.abr.2017.06.001
210. Guillou  L, Chrétiennot-Dinet  M-J, Boulben  S  et al.  Symbiomonas scintillans gen. et sp. nov. and Picophagus flagellatus gen. et sp. nov. (Heterokonta): two new heterotrophic flagellates of picoplanktonic size. Protist  1999;150 :383–98. 10.1016/S1434-4610(99)70040-4 10714773
