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Phytomyxea

  • Simon BulmanEmail author
  • Sigrid Neuhauser
Living reference work entry

Abstract

Phytomyxea are endoparasites of Plantae or heterokont hosts. They are distributed between two orders, the Plasmodiophorida and Phagomyxida. Several phytomyxids cause the formation of large galls on their hosts, but many are only visible via light microscopy of roots, hyphae, or phytoplankton. PCR and sequencing from environmental DNA samples is beginning to reveal many new phytomyxid lineages. Phytomyxids persist in the environment as thick-walled, uninucleate resting spores. These germinate to produce zoospores which locate hosts via the propulsion of heterokont flagellae. Cell penetration by encysting zoospores is via a distinctive projectile-like extrusome. Intracellular growth forms are multinucleate, unwalled protoplasts termed plasmodia. A synapomorphy for the class is cruciform mitotic division at metaphase. Plasmodiophorids occupy a phylogenetically distinct position from most other plant parasites. They cause several economically important diseases such as clubroot of Brassicaceae and powdery scab of potatoes. Plasmodiophorids also act as vectors of damaging plant viruses.

Keywords

Plant parasites Heterokont parasites Intracellular Extrusome Virus vector Cruciform divisions Plasmodiophora 

Summary Classification

  • Phytomyxea

  • ••Plasmodiophorida (Plasmodiophora, Spongospora, Woronina, etc.)

  • ••Phagomyxida (Phagomyxa, Maullinia, etc.)

Introduction

General Characteristics

Organisms in the class Phytomyxea are endoparasites of plants or heterokonts. They are distributed between two orders, the Plasmodiophorida, to date found in soil/freshwater, and Phagomyxida, found in marine ecosystems. Phytomyxids persist in the environment as thick-walled, uninucleate resting spores. Upon resting spore germination, the released zoospores locate hosts propelled by smooth heterokont flagellae. Cell penetration is via a distinctive projectile-like extrusome . Intracellular growth forms are multinucleate, unwalled protoplasts termed plasmodia. A synapomorphy for the class is cruciform mitotic division at metaphase. Plasmodiophorids cause several economically important plant diseases – either directly or as vectors of viruses .

Literature and History of Knowledge

The first phytomyxid genus was erected by Woronin in 1877 to accommodate Plasmodiophora brassicae , the organism causing clubroot of cabbage. Literature published in the early 1900s was based largely on light microscopy (Cook 1933). Much of the phytomyxid species discovery occurred during this period, with knowledge summarized in important monographs by Karling (1942, 1968). Ultrastructural and karyological studies predominated into the 1990s, leading to a greatly increased understanding of phytomyxid lifecycles (Dylewski 1990; Braselton 1995; Bulman and Braselton 2014). Considerable research during this period was carried out on the oomycete parasite Woronina and on the Veronica parasite, Sorosphaerula veronicae, whereas little has been published on these organisms in subsequent decades. Continuing research is centered upon phytomyxid species that cause diseases of economically important crops, while reports on other phytomyxids are infrequent. Physical identification of new phytomyxid species, such as the recently described parasite of grapes Sorosphaerula viticola (Kirchmair et al. 2005), is rare. With greater focus on marine ecosystems, a small number of new phagomyxid parasites have been discovered (Maier et al. 2000; Schnepf et al. 2000; Goecke et al. 2012). An accessible and important source of information on the plasmodiophorids remains the Plasmodiophorid Home Page (http://oak.cats.ohiou.edu/~braselto/plasmos/).

Beginning with first the DNA-based studies (Mutasa et al. 1993), techniques of molecular biology are now driving large changes to our understanding of phytomyxid biology and diversity. Molecular tools were initially developed for pathogen detection (Buhariwalla et al. 1995), delineation of populations and species within Polymyxa, and assessing the presence of plant viruses associated with phytomyxids (Legreve et al. 2002; Kanyuka et al. 2003; Smith et al. 2013). Molecular phylogenetic analyses succeeded in finding a taxonomic home for the phytomyxids in Rhizaria (CavalierSmith and Chao 1997; Archibald and Keeling 2004; Nikolaev et al. 2004; Burki et al. 2010) and confirmed the close relationship of Phagomyxida with Plasmodiophorida (Bulman et al. 2001; Neuhauser et al. 2014). While identification of new phytomyxid lineages in broadly targeted culture-independent studies is not common, such works do suggest a diversity of phytomyxids not yet seen with the naked eye or by microscopy (Takishita et al. 2005, 2007). Importantly, anonymous DNA sequencing coupled with targeted PCR amplification is revealing many new lineages of plasmodiophorids (Neuhauser et al. 2014). Studies of phytomyxid genomes, which were initially slowed by the obligate biotrophic nature of these protists, began with small collections of DNA sequences (Bulman et al. 2006, 2007; Siemens et al. 2009). Early utilization of next-generation sequencing techniques provided a better understanding of the phylogenomics of plasmodiophorids (Burki et al. 2010). A complete mitochondrial DNA sequence from Spongospora subterranea (Gutierrez et al. 2014) and the first complete phytomyxid genome sequence, from P. brassicae (Schwelm et al. 2015a), have been published. The greatest amount of research continues to be focused on P. brassicae infection of the model plant Arabidopsis thaliana (Devos et al. 2006; Siemens et al. 2006; Malinowski et al. 2012). Biochemical and cellular characterization of this interaction is accelerating based on new genomic data (Feng et al. 2010; Ludwig-Muller et al. 2015). The first technique for the genetic transformation of P. brassicae has recently been published, potentially opening up greater opportunities for characterization of gene function in this parasite (Feng et al. 2013). A node of clubroot research has developed as a response to the large losses caused by the disease in the Canadian canola industry (Hwang et al. 2012).

Practical Importance

The best studied phytomyxids infect important agricultural food plants worldwide. Plasmodiophora brassicae causes clubroot in Brassicaceae (Dixon 2009). Spongospora subterranea causes powdery scab of potato (Merz and Falloon 2009). Spongospora nasturtii causes crook root disease in watercress (Claxton et al. 1996). Polymyxa graminis infects the roots of grasses such as wheat, oats, and rice, and Polymyxa betae grows in the roots of sugar beets. Phytomyxids are known to transmit about 20 plant viruses, all but one of which are non-enveloped, positive polarity, single-stranded RNA viruses (Rochon et al. 2004). Polymyxa species do not directly cause disease but instead transmit a range of damaging viruses (Kanyuka et al. 2003; Rochon et al. 2004). Viruses transmitted by P. graminis include wheat spindle streak mosaic virus, oat golden stripe virus, rice necrosis mosaic virus, and peanut clump virus. Polymyxa betae transmits viruses including beet necrotic yellow vein virus, the cause of sugar beet rhizomania. Spongospora subterranea and S. nasturtii, respectively, transmit potato mop-top virus and watercress yellow spot virus. The resting spores of plasmodiophorids remain viable in soil for many years, and there are few pesticides available for control of diseases caused by plasmodiophorids. Once soils become infested with resting spores, it is difficult to continue cropping of the susceptible host.

The ecological roles of phagomyxids in marine environments are a matter of increasing interest but as yet have been little studied (Neuhauser et al. 2011).

Habitats and Ecology

As obligate biotrophs, phytomyxid distribution follows that of their hosts (Table 1). Phytomyxids are mostly not available from culture collections. To date, Plasmodiophorida are found in terrestrial and freshwater environments where they parasitize plants and heterokonts such as Phytophthora and Pythium spp. Both P. brassicae and S. subterranea are found worldwide in soils where Brassicaceae and potatoes are grown. Polymyxa, Sorosphaerula, and Ligniera species are common soilborne plant parasites , found widely in arable and natural environments. Several phytomyxids including Spongospora nasturtii, Tetramyxa spp., Sorodiscus callitrichis, and Membranosorus spp. parasitize aquatic vascular plants. Sorodiscus karlingii infects charophyte algae. Phagomyxida so far discovered are marine parasites of heterokonts , including diatoms and brown algae.
Table 1

A selection of the most common and consistently reported phytomyxids together with information on host taxa

Speciesa

Host and location

Citation

Plasmodiophorida

Ligniera verrucosa

Veronica, Beta, Chenopodium, Bromus, and Festuca spp. (P, A)

Miller et al. (1985)

L. junci 7

Especially Juncus spp., but many wild and cultivated plants (P, A)

Neuhauser and Kirchmair (2009)

L. pilorum

Poa annua, Bromus inermis (P, A)

Barr (1979)

Membranosorus heterantherae

Heteranthera dubia (P, A)

Forest et al. (1986)

Octomyxa achlyae

Achlya glomerata (H, O)

Dylewski (1990)

O. brevilegniae

Brevilegnia linearis, Geolegnia inflata. (H, O)

Dylewski (1990)

Plasmodiophora brassicae 10

Brassicaceae plants (P, A)

Dixon (2009)

P. bicaudata

Internodes of Zostera nana (P, A)

den Hartog (1989)

P. halophilae

Halophila spp. (P, A)

Marziano et al. (1995)

Polymyxa graminis 1 and 6

Many cultivated and wild grasses including sorghum, oats, wheat (P, A)

Vaianopoulos et al. (2007)

P. betae 5

Many plants including Beta vulgaris, Chenopodium spp. (P, A)

Barr (1979)

Sorodiscus callitrichis

Stems of Callitriche spp. (P, A)

Robbins and Braselton (1997)

S. karlingii

Chara contraria, C. delicatula (P, C)

Cook (1933)

Sorosphaerula veronicae 4

Roots and stems of Veronica spp. (P, A)

Miller (1958)

S. viticola 3

Grapes (Vitis sp.) (P, A)

Neuhauser et al. (2009)

Spongospora subterranea 9

Roots and tubers of Solanum spp. (P, A)

Merz and Falloon (2009)

S. nasturtii 15

Nasturtium officinale, N. microphyllum. (P, A)

Claxton et al. (1996)

S. campanulae

Campanula rapunculoides. (P, A)

Cook (1933)

S. cotulae

Cotula australis (P, A)

Karling (1968)

Tetramyxa parasitica

Ruppia, Zannichellia, Potamogeton species (P, A)

Braselton (1990)

Woronina pythii 13

Pythium spp. (H, O)

Dylewski (1990)

W. glomerata

Vaucheria spp. (H, YGA)

Dylewski (1990)

W. leptolegniae

Leptolegnia caudata. (H, O)

Karling (1981)

W. cokeri

Pythium spp. (H, O)

Robbins and Braselton (1997)

Env2

Glacier forefield soil, Austria

Neuhauser et al. (2014)

Env8 and 12

Fynbos soil, South Africa

Neuhauser et al. (2014)

Env11, 14, 16

Volga soil, Russia

Neuhauser et al. (2014)

Phagomyxida

Phagomyxa algarum

Ectocarpus mitchellae, Pylaiella fulvescens (H, BA)

Karling (1944)

P. bellerocheae

Bellerochea malleus (H, D)

Schnepf et al. (2000)

P. odontellae

Odontella sinensis (H, D)

Schnepf et al. (2000)

Maullinia ectocarpii

Ectocarpus siliculosus (H, BA)

Maier et al. (2000)

Maullinia sp.

Durvillaea antarctica (H, BA)

Goecke et al. (2012)

Env (2)

Anoxic sediments

Takishita et al. (2005) and Takishita et al. (2007)

Plasmodiophora diplantherae

Stem galls Halodule wrightii (P, A)

Braselton and Short (1985)

H Heterokontophyta, P Plantae, A Angiosperm, C Charophyta, O Oomycota, D diatom, BA brown algae, YGA yellow green algae. Infections of green plants occur in roots unless otherwise stated

aNumbers in superscript indicate phylogenetic clades from Neuhauser et al. (2014). A subset of phytomyxid clades that have only been detected via environmental sequencing (Env) is also shown. A citation specifically focused on the organism in question, or a review article giving such information, is also provided

Since the retirement of prominent researchers in this field, knowledge of sites to collect many phytomyxid species is restricted. Records of non-crop-infecting phytomyxids are increasingly sporadic and often in the realm of citizen science. Species of Tetramyxa, Sorodiscus, Membranosorus, and, especially, Octomyxa are currently little studied. Although the distribution of many phytomyxids is reportedly limited, some studies of herbarium samples suggest a broader geographical distribution than previously recognized (Forest et al. 1986; den Hartog 1989).

Anonymous DNA sequencing techniques have revealed new phagomyxid lineages in anoxic marine/saline environments (Takishita et al. 2005, 2007) and plasmodiophorid lineages from geographically widespread soil and rhizosphere sites (Lesaulnier et al. 2008; Bass et al. 2009; Neuhauser et al. 2014). Although anonymous phytomyxid sequences have not so far been associated with specific hosts, the low diversity of sequences in any one sample, and greater abundance in rhizosphere versus bulk soil, implies a close relationship with plant hosts (Neuhauser et al. 2014). Cloning of anonymous sequences also suggests that oomycete-infecting phytomyxids are diverse and widely distributed in soils (Neuhauser et al. 2014).

Characterization and Recognition

By far the majority of phytomyxid life-cycle research has been carried out on plasmodiophorids such as P. brassicae (Kageyama and Asano 2009), S. veronicae (Miller 1958), and W. pythii (Dylewski 1990), rather than phagomyxids. The most recognized phytomyxid life cycle has a bipartite format; a composite life-cycle scheme, most strongly drawn from plant-infecting Plasmodiophorida, is presented here (Fig. 1).
Fig. 1

A Phytomyxid life-cycle scheme drawn mostly from plasmodiophorid infection of crop plants. Variations to this life cycle may occur in some species such as among the marine phagomyxids. 1, environmentally resistant resting spore; 2, biflagellate primary zoospore; 3, location of host cell by zoospore and commencement of encystment – for many plasmodiophorids, primary infection occurs in root hairs; 4, cell penetration by Stachel followed by zoospore contents; 5, development of multinucleate plasmodium; 6, multilobed structure containing zoosporangia; 7, secondary zoospore – reinfection of host cells occurs by encystment as in 3 and 4, or via a myxamoeboid phase (dashed lines to 8); 8, secondary plasmodium; 9, resting spores or aggregates thereof. Dashed lines indicate uncertainty about direct progression to secondary infection mediated via primary zoospores or generation of new cycles of primary infection by secondary zoospores

Penetration of Host

Phytomyxids persist over time through environmentally resistant resting spores. These germinate to produce heterokont primary zoospores that exhibit a cyclotic swimming motion. On encountering the host, zoospores retract their flagellae and begin a characteristic infection process termed encystment. An infection apparatus develops within the zoospore, consisting of a tubular cavity (Rohr) containing a bullet-like structure (Stachel), with one end oriented in the direction of the host wall (Keskin and Fuchs 1969; Aist and Williams 1971). The Rohr rapidly contracts, and the Stachel penetrates the host wall followed by the unwalled, uninucleate protoplast of the parasite, which is presumably forced out by turgor pressure created by the expansion of a large vacuole in the encysted zoospore (Fig. 1).

Sporangial Plasmodia

Inside infected cells, the uninucleate protoplast matures into a zoosporangial plasmodium, with a 9–24 nm host-plasmodiophorid interface (Aist and Williams 1971; Braselton and Miller 1975; Miller and Dylewski 1983a). Synchronous mitotic divisions yield a multinucleate plasmodium (Dylewski and Miller 1983). These cruciform divisions are the major synapomorphy defining phytomyxids (Braselton and Miller 1975). At metaphase, chromatin aligns at the equator of the nucleus, perpendicular to the elongating, persistent nucleolus. A symmetrical cross is formed that can be seen by light microscopy (Fig. 2) (Dylewski et al. 1978; Garber and Aist 1979).
Fig. 2

Transmission electron micrograph of cruciform division in sporogenic plasmodium of Plasmodiophora brassicae on Chinese cabbage (Brassica rapa). N nucleolus, Ce centriole, Ch chromatin. Scale bar = 0.5 μM. Photograph James Braselton, Plasmodiophorid Homepage

After the mitotic divisions, the plasmodium cleaves into a thin-walled multicelled structure in the infected cell (Fig. 3). Zoospore formation occurs as the protoplasm within each zoosporangium cleaves. Secondary zoospores from zoosporangia may be released outside of the host, into adjacent cells, or into the same cell. Conspicuous exit tubes may be formed between the zoosporangia and adjacent host cells (Littlefield et al. 1998).
Fig. 3

Spongospora subterranea zoosporangia in trypan blue stained potato roots. (a) root with heavily infected root hairs; (b) infected root epidermal cells; (c) root hair with zoosporangia; (d) and (e) root hair and epidermal cells containing empty zoosporangia following zoospore release. Photographs Richard Falloon

Sporogenic Plasmodia

The sporogenic phase culminates in the formation of thick-walled resting spores (Fig. 4). At the cessation of sporogenic division, cleavage furrows appear and meiosis begins (Dylewski and Miller 1984). Nucleoli begin to disperse during prophase of meiosis I, rendering the nuclei less obvious in the plasmodium. It is believed that chromosome number is halved as a result of meiosis during resting spore formation (Dylewski 1990). Eventually, each nucleus is partitioned, forming binucleate resting spores. One nucleus presumably undergoes degeneration because all mature resting spores become uninucleate (Dylewski and Miller 1984).
Fig. 4

Morphology of resting spores from selected phytomyxids: left column, Plasmodiophorida; right column, Phagomyxida. (a) Sorosphaerula viticola: hollow sporosori in the roots of Vitis sp. (b) Woronina pythii: resting spores in Pythium sp. (c) W. pythii in Pythium sp.: lobose plasmodium, just starting to develop into resting spores (arrow); right mature resting spores. (d) Ligniera junci: resting spores in the root hairs of Juncus effusus. (e) Maullinia sp. resting spores in Durvillaea antarctica. (f) Plasmodiophora diplantherae: resting spores in enlarged cells of Halodule sp. Arrow: starch grains. (g) Maullinia ectocarpii: hatching zoospores (arrow) from an enlarged infected cell of the host Ectocarpus fasciculatus. *Plasmodia in enlarged host cells. Scale bar = 10 μM. Photographs Sigrid Neuhauser

Many phytomyxids have resting spores arranged in aggregate bodies called sporosori. For example, Sorodiscus sporosori are usually composed of two closely pressed layers of resting spores, whereas those of Membranosorus occur in a single layer usually lining the inner periphery of the host cell (Fig. 5). Spongospora subterranea sporosori are particularly large and distinctive; approximately 200–700 resting spores are aggregated into spongelike structures of variable size (Falloon et al. 2011) (Fig. 5).
Fig 5

(a) Scanning electron micrograph of Spongospora subterranea sporosorus showing individual spores with punctate outer surface ornamentation. Scale bar = 10 μM. Photograph Ueli Merz. (b) Transmission electron micrograph of S. subterranea secondary zoospores in zoosporangia. Scale bar = 3 μM; (c) Scanning electron micrograph of Membranosorus heterantherae sporosori. Scale bar = 8 μM. (d) Transmission electron micrograph of a sporosorus of M. heterantherae. Scale bar = 1 μM. (bd) Photographs James Braselton, Plasmodiophorid Home Page

Sporogenic plasmodia become more abundant as the host ages. In several plasmodiophorid species, sporangial and sporogenic plasmodia are distinguishable by their occurrence in separate tissues. In P. brassicae, sporangial plasmodia develop in root hairs and epidermal cells, whereas sporogenic plasmodia are found in the root cortex and stele. In S. veronicae, sporosori are produced only in galls on shoots and not during root infections. In S. subterranea, zoosporangia occur soon after infection in root epidermal cells, while sporosori are formed later in root galls and tuber lesions. Although sporangial development typically precedes sporogenic development, both stages can be seen in the same tissue early in Polymyxa infection (Ledingham 1939). Primary zoospores may be capable of initiating both sporangial and sporogenic plasmodia in P. brassicae (Mithen and Magrath 1992; McDonald et al. 2014). The biochemical and developmental factors that determine the transition to sporogenic growth are not known. In cultures, the state of the culture medium appears to have an influence on the development path of Woronina plasmodia (Miller and Dylewski 1983b).

Sporogenic development is associated with growth of hypertrophic plant galls characteristic of several plasmodiophorid diseases. While sporogenic development is considered to be initiated via secondary zoospore infection, there are persistent reports of direct penetration of the root cortex. Indeed, a myxamoeboid stage is nearly an accepted part of the plasmodiophorid life cycle, despite the exact nature of this stage remaining unclear (Mithen and Magrath 1992; Claxton et al. 1996; Kobelt et al. 2000; Asano and Kageyama 2006). Few genes encoding proteins with cellulose-binding domains, which might be involved in the cell wall modification needed to penetrate into new plant cells, were detected in the P. brassicae or S. subterranea transcriptomes (Schwelm et al. 2015b).

Nutrition

There are some unanswered questions about the degree of phagotrophy in phytomyxids. As the genus name suggests, ingestion of host material has been reported as a feature of nutrition in Phagomyxa (Karling 1944; Schnepf et al. 2000). On the other hand, during early sporangial growth of plasmodiophorids, pseudopodial-like extensions of protoplasm grow outward and partially surround host organelles and cytoplasm, but it has generally been agreed that these fail to completely surround host cytoplasm and that there is consequently no phagotrophic nutrition (Dylewski 1990).

Karyogamy

The occurrence of karyogamy in phytomyxids is not well understood. Protoplasm fusion was claimed to occur between haploid secondary zoospores or between the nuclei in plasmodia, prior to resting spore formation and the onset of meiosis (Ingram and Tommerup 1972). Potential karyogamy in sporogenic P. brassicae plasmodia has also been reported (Buczacki and Moxham 1980).

Atypical-Host Infection

An unusual feature of plasmodiophorids is their appearance in a wide range of hosts beyond those in which they complete a full life cycle. Primary plasmodia have been observed in the roots of such atypical plant hosts, with little or no evidence for progression to secondary plasmodia. For example, S. subterranea and P. brassicae have been observed in many plant species other than their respective Solanum and Brassicaceae hosts (Ludwig-Muller et al. 1999; Qu and Christ 2006). A model for atypical host infection is provided by the Polymyxa-Arabidopsis thaliana interaction (Desoignies et al. 2010). Frequent host shifts have occurred during the evolution of phytomyxids; whether these host shifts are related to promiscuous host infections at primary stages remains to be investigated (Neuhauser et al. 2014).

Maintenance and Cultivation

Collection and Isolation

Phytomyxids are “isolated” from soil or water samples through infection of their hosts. Plasmodiophorids may be collected from wild sources or from hosts deliberately planted in infested potted soils. Plant-infecting species of Plasmodiophora (Fig. 6), Spongospora, Sorosphaerula, Sorodiscus, and Tetramyxa produce obvious galls or hypertrophies that are easily collected from infected plants. Spongospora subterranea is most frequently collected from sporosori-filled scabs on potato tubers (Fig. 6). Ligniera and Polymyxa species must be found by microscopically examining the roots of hosts, which is time-consuming due to the lack of external symptoms (Fig. 4).
Fig. 6

Plasmodiophorid plant infections. (a) Chinese cabbage (Brassica rapa) plant showing heavy clubroot symptoms (Plasmodiophora brassicae infection). Scale bar = 10 cm; (b) Arabidopsis thaliana Columbia-0 plants with (left) and without clubroot infection. Scale bars = 1 cm. Photograph Robert Lamberts/Simon Bulman; (c) Potato tuber with severe symptoms of powdery scab (Spongospora subterranea). Scale bar 3 cm (Photograph Robert Lamberts/Richard Falloon)

Woronina and Octomyxa spp. are typically attracted to hosts growing on seeds added to water or water amended with soil. Samples may be baited with specific oomycetes if available. Oomycete-infecting plasmodiophorids are then detected by light microscopy (Fig. 4).

Phagomyxid species are identified through microscopic surveys of marine heterokont hosts. Phagomyxa odontellae and P. bellerocheae are found infecting diatoms in marine phytoplankton samples (Fig. 7) (Schnepf 1994; Schnepf et al. 2000). Maullinia spp. can be collected from galls on marine brown algae macrophytes (Fig. 8). Resting spores were observed for Maullinia infecting Durvillaea antarctica, raising the possibility that this species may be maintained in a viable form within collections (Goecke et al. 2012).
Fig. 7.

Phagomyxa bellerocheae infecting the diatom Bellerochea malleus. (a) Plasmodia containing secondary zoospores. (b) Released zoospores with whiplash flagellae (arrowed). Scale bars 10 μM. Photographs Eberhard Schnepf

Fig. 8

Maullinia. (a) Gall-like structures on infected Durvillaea antarctica fronds from central Chile. Scale bar = 1 cm. Photograph Franz Goecke; (b) Type slide at the NHM London (registration number: 2000:2:29:1) showing zoosporangia of Maullinia ectocarpi. Scale bar = 10 μm. Photograph Sigrid Neuhauser

Cultivation

Studies of the interactions between phytomyxids and their hosts are most tractable for P. brassicae which infects the model plant Arabidopsis thaliana (Fig. 6). For manipulating P. brassicae, spore suspensions are prepared by maceration and filtering of decayed galls (Castlebury et al. 1994). For new plant infection, the suspension is applied to soil surrounding plant seedlings. Temperature and pH are important for disease progression, with 20 °C and pH <7 being typical conditions for maximal P. brassicae growth. Clubroot galls or P. brassicae spore suspensions may be kept frozen in a viable state for at least 3 years.

Sporosori samples from S. subterranea are prepared by scraping scabs from potato tubers followed by air drying and sieving. A solution-culture assay (Merz 1989) has been adopted for studying the potato-S. subterranea interaction.

Although phytomyxids cannot be cultured in the absence of their hosts, several publications have detailed the establishment of dual cultures of plasmodiophorids with plant cells. Plasmodiophora brassicae and S. subterranea have been grown in hormone-induced callus/cell cultures (Buczacki 1983; Asano and Kageyama 2006; Bulman et al. 2011). Plasmodiophora brassicae , S. subterranea, and P. betae have been grown with Agrobacterium-induced in vitro hairy root cultures (Mugnier 1987; Qu and Christ 2007).

Dual cultures of plasmodiophorids and oomycete hosts in water and “soft” agar media have been established for W. pythii and W. cokeri (Miller and Dylewski 1983a). Resting spores of W. pythii may be dried on filter paper and germinated by rehydration after up to 14 months at 6 °C (Miller and Dylewski 1983a). Laboratory co-cultures of Maullinia ectocarpii with a range of brown algae macrophytes have been established under controlled conditions (Maier et al. 2000).

Evolutionary History

Classification

Phytomyxids are likely to be at least 400 million years old based on fossil records (Taylor et al. 1992). For a long period, their taxonomic position was unstable, oscillating between fungi, slime molds, and protozoa (Barr 1981). The first ribosomal DNA sequence from P. brassicae provided evidence for a relationship between plasmodiophorids and Cercozoa (Cavalier-Smith and Chao 1997; Castlebury and Domier 1998). Assembly of sequences from a greater diversity of protists has confirmed that this grouping with Rhizaria (Nikolaev et al. 2004; Bass et al. 2005, 2009) and that Phagomyxida belong in Phytomyxea (Bulman et al. 2001). Plasmodiophorid polyubiquitin sequences were shown to have an unusual amino acid insertion, as do those from Cercozoa and Foraminifera (Archibald et al. 2003; Archibald and Keeling 2004). Phytomyxids fall within the subphylum Endomyxa that includes a mixture of free-living and parasitic organisms including vampyrellid amoebae (predators), Filoreta (bacterivores), Ascetosporea (parasites of marine invertebrates), and Gromia (Bass et al. 2009). Phylogenomic studies have mostly indicated that Endomyxa is a distinct clade (Burki et al. 2010; Sierra et al. 2013; Cavalier-Smith et al. 2015). The exact phylogenetic position of Phytomyxea relative to other endomyxans remains to be finalized, although ribosomal phylogenies point to vampyrellids as close relatives (Bass et al. 2009).

Phytomyxea genera were historically designated by the aggregation of resting spores in sporosori and by ultrastructure, with less emphasis placed on host affiliations. However, S. nasturtii was raised to species rank partly on the basis of its significantly different host to S. subterranea (Dick 2001), while W. cokeri was moved to the genus Woronina harboring other oomycete parasites (Robbins and Braselton 1997). Across the last century, a large number of phytomyxids were described; many of the reported species appear to have been synonyms or were doubtful taxa, as reviewed in Karling (1942; 1968). A summary of some of these taxa is presented in Table 1. Although each phytomyxid genus could once be uniquely identified by spore arrangement and ultrastructure (Dylewski 1990), it is now clear that neither feature provides a firm framework for understanding within-group relatedness. As with the overall phylogenetic position of Phytomyxea, the internal relationships of the group have been radically altered by the advent of DNA techniques. Ribosomal small subunit RNA phylogenies showed large evolutionary distances between plasmodiophorid species that were largely indistinguishable by morphology. For example, Spongospora subterranea and S. nasturtii were found to be phylogenetically remote from one another (Bulman et al. 2001). Even more strikingly, Plasmodiophora diplantherae was shown to be a phagomyxid rather than plasmodiophorid (Neuhauser et al. 2014). Anonymous DNA sequencing coupled with specific PCR has now revealed many new distinct lineages, especially within Plasmodiophorida (Neuhauser et al. 2014); a selection of these environmental lineages is listed in Table 1. An intermixed cluster of Polymyxa, Sorosphaerula, and, to a lesser degree, Ligniera species was confirmed (Neuhauser et al. 2014). This group of genera appears ripe for taxonomic revision based on a combination of ecological and DNA data. Anonymous DNA sequences also indicated a significant diversity of lineages in the Woronina clade (Neuhauser et al. 2014). It will be highly informative to use molecular techniques to link these Woronina-like lineages with their, presumably, oomycete hosts. Phagomyxid lineages in marine ecosystems await exploration with the techniques of molecular ecology.

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Copyright information

© Springer International Publishing AG 2016

Authors and Affiliations

  1. 1.The New Zealand Institute for Plant & Food Research LimitedChristchurchNew Zealand
  2. 2.Institute of MicrobiologyUniversity of InnsbruckInnsbruckAustria

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