Keywords:Perkinsus, Parvilucifera, X-cell parasite, broad host range parasite, emerging diseases, severe Perkinsea infection, opportunistic parasite

Perkinsus spp., an Emerging Parasite of Mollusks

In 1946, following a mass mortality event (MME) of oyster stocks in Louisiana (Gulf of Mexico, United States),Mackin et al. (1950) identified a protist present in host tissue sample as the causative agent. First affiliated to a fungal lineage and namedDermocystidium marinum (Mackin et al., 1950), electronic microscopy observations then showed morphological characteristics, such as the presence of a subpellicular membrane, micropores, and a conoid-like structure (zoospore stage) suggesting a re-affiliation of this protist into the Apicomplexa phylum (Alveolata) (Perkins and Menzel, 1967;Perkins, 1976). While doubts remained about the accuracy of this affiliation (Siddall et al., 1997), molecular phylogenies based on ribosomal and actin sequences revealed thatPerkinsus species were more closely related to dinoflagellates than apicomplexans (Reece et al., 1997). In 2003, multiple protein phylogeny indicated thatPerkinsus is an early branch in dinoflagellate lineage (see schematic representation inFigure 2A;Saldarriaga et al., 2003).

Nowadays, seven species are described within the genusPerkinsus:P. marinus,P. olseni,P. qugwadi,P. chesapeaki,P. mediterraneus,P. honshuensis, andP. beihaiensis (seeFigures 1B,2B for the phylogenetic classification of these parasites). However, onlyP. marinus andP. olseni, the etiological agents of “Dermo” disease and Perkinsosis, respectively, have significant negative impacts on mollusk populations worldwide (Siddall et al., 1997). These two infectious agents are today within the list of notifiable diseases in the World Organization for Animal Health (OIE) (Oie-Listed Diseases, 2021: OIE—World Organisation for Animal Health).

SomePerkinsus species have a putative BHR among mollusk species as, for exampleP. olseni infecting various clams (e.g.,R. decussatus,R. philippinarum,Austrovenus stutchburyi,Tridacna maxima,T. crocea,and Pitar rostrata), oysters (Crassostrea rhizophorae,Saccostrea sp.), pearl oyster (e.g.,Pinctada imbricata andP. fucata), and abalone (e.g.,Haliotis rubra andH. laevigata) (Lester and Davis, 1981;Cremonte et al., 2005;Dungan et al., 2007;Sheppard and Phillips, 2008;Sanil et al., 2010;Ramilo et al., 2015;Pagenkopp Lohan et al., 2018), orP. chesapeaki, which can infect clams (R. decussatus,R. philippinarum) and oysters (C. rhizophorae,C. virginica) (Coss et al., 2001;Arzul et al., 2012;Dantas Neto et al., 2016). Conversely otherPerkinsus species show an intermediate host range as, for example,P. marinus infecting mostly oysters (e.g.,C. agar,C. rhizophorae,C. virginica,C. agar, andSaccostrea palmula) (e.g.,Enríquez-Espinoza et al., 2010;Cáceres-Martínez et al., 2012;da Silva et al., 2013,2014).

Until now, most of the host species described above are of commercial interest but other bivalve and gastropod species may also be susceptible to infection, especially in the described geographical range. However, thePerkinsus putative wide host range established solely using microscopic and molecular methodology may also be overestimated. Indeed, these results do not allow the identification of dead-end hosts that prevent the parasite transmission to the definite hosts. Furthermore, despite their BHR character, host susceptibility needs to be considered to prevent and protect most sensible bivalve stocks. For example, the two genetically related oystersC. gigas andC. virginica respond with different susceptibility when challenged byP. marinus. Although infection is established in both hosts, only infection inC. virginica is lethal (Barber and Mann, 1994). AmongPerkinsus species, different levels of prevalence depending on hosts species are recorded across the world (Figure 3). In the Chesapeake Bay,P. marinus andP. chesapeaki infect different species of oysters and clams living in sympatry. Among these hosts,P. marinus rarely infects clams compared withP. chesapeaki and exhibits oyster preference (Reece et al., 2008). Variability in pathogenicity across the BHR ofPerkinsus spp. raises the question about the existence of specific strains. For example, such specific strain was recently highlighted forPerkinsus marinus with the emergence between 1983 and 1990 of a new hypervirulent phenotypic strain that includes a shortened life cycle and a trophism shift from deeper connective tissues to digestive epithelia (Carnegie et al., 2021). Authors hypothesized that the development of this new strain may be related to reduced oyster abundance and the rapid establishment of the exotic parasiteH. nelsonii in 1959. Hence, given the increase in epizootic diseases (= disease event in a non-human animal population analogous to an epidemic in humans) worldwide, consideration of the host range is absolutely essential because i) they may be the starting point for a host shift, ii) they represent vector of transmission with no apparent signs of disease, and iii) new native parasitic strains can emerge in response to biotic or abiotic stressors.

Perkinsus infection appears to occur directly by filtration without an intermediate host, with gills and labial palps playing a crucial role as entrance portal for the parasite (Chintala et al., 2002;Allam et al., 2013;Wang et al., 2018). Four different life stages (Figure 4A) are described today occurring inside or outside the host with an exception forP. qugwadi where all stages can be observed in host tissues (Blackbourn et al., 1998). All of them can induce new infections into a healthy host. The trophozoite life stage proliferates by vegetative multiplication (palintomy) in the host tissue. Disruption of the cell wall allows the release of immature spherical trophozoites that will gradually enlarge becoming mature vegetative cells (Perkins, 1996). In case of heavy infection, trophozoites can invade totally host tissues, with occasional production of cutaneous white nodules (inflammatory reaction), and induce a global decrease in host fitness (e.g., lower filtration activity, retard in growth and reproduction) (Dittman et al., 2001;Lee et al., 2001;Choi and Park, 2010). When the host becomes moribund, the parasite enlarges and develops a thick cell wall becoming a hypnospore life stage (Valiulis and Mackin, 1969;Perkins, 1996). When transferred to fresh seawater under favorable conditions of temperature and salinity (Auzoux-Bordenave et al., 1996), hypnospores undergo multiple divisions leading to the formation of free-living biflagellate life stages, the zoospores, into a cellular structure called zoosporangium. At maturity, the zoosporangium releases zoospores in the medium allowing the infection of new hosts.

Perkinsus species are one of the most famous examples of successful invasive pests in marine environments (Figure 3A). Three different scenarios are described through three differentPerkinsus species:(1) Expansion: P. marinus has shown a significant increase in its geographic area in the United States along the East coast by more than 500 km in 2 years (1990–1992). This expansion, which is now permanent, has been correlated with an increase in winter surface temperature due to milder winters (Ford, 1996;Ford and Smolowitz, 2007).(2) Introduction: P. chesapeaki appears to have been accidentally introduced in Europevia its vectorsMya arenaria, or the hard clam,Mercenaria mercenaria from the United States (Arzul et al., 2012). The sporadic detection ofP. chesapeaki does not reflect an expansion dynamic. To date, no mortality event affiliated toP. chesapeaki has been recorded.(3) Introduction and expansion: P. olseni has been co-introduced in Europe with its host the Manila clam,Ruditapes philippinarum, in 1972 for the development of clam aquaculture (Ruano et al., 2015). After its first detection in the late 1980s, the first mortalities of clam stocks were attributed to this parasite at Ria de Faro in Portugal (Ruano and Cachola, 1986). Since then,P. olseni has produced mortalities in Spain, Portugal, and Italy, infecting both the exotic Manila clam,Ruditapes philippinarum, and the native European clam,Ruditapes decussatus (Azevedo, 1989;Figueras et al., 1992;Pretto et al., 2014). Following these first mortalities,P. olseni was detected along the French Atlantic coast with infection prevalence up to 100% (Lassalle et al., 2007), in contrast to the non-detection ofPerkinsus spp. in most of (these) previously studied sites (Goggin, 1992). Similarly, histological studies conducted byVilela (1951) onMinchinia tapetis (Haplosporidia) infection in the Formosa lagoon in Portugal, where the prevalence ofP. olseni infection is now very high, ranging from 60 to 90% depending of the season (Leite et al., 2004), did not mention any infection withPerkinsus spp. Finally, in 2011, using microsatellite genetic diversity analysis, Vilas et al. described low genetic diversity of theP. olseni parasite in Spain and Portugal compared with samples collected in Japan and New Zealand (Vilas et al., 2011), which might result in a founder effect testifying an introduction event of this parasite. These examples ofPerkinsus spp. dissemination demonstrates the ubiquitous nature (exploitation of an important range of abiotic and biotic niches) ofPerkinsus species, which is a threat for shellfish farming economy and for ecosystem equilibrium. Bivalves are known to provide important ecosystem services through their status as habitat engineers, as filter feeders, and as a connecting compartment between primary producers and predators (Jones et al., 1996;Coen and Grizzle, 2007;Gallardi, 2014). By cascade effect, parasitic infection can have deleterious effects on the different functions attributed to these populations: (i) by altering their bioturbatory activity and, thus, by modifying biogeochemical cycles at the water–sediment interface (Dairain et al., 2019), (ii) by impacting their filtration activity participating in the nitrogen cycle (= production of NH₄⁺ leading to an increase in primary production) and helping to reduce turbidity (due to the increase in light penetration) (Gallardi, 2014), and (iii) by altering their physiological traits (e.g., growth, reproduction, and survival), which will modify the structure of habitat, biotic interaction, and the associated species richness (e.g.,Thomas and Poulin, 1998). In addition to this infectious threat, benthic biodiversity is threatened by human activities (McCauley et al., 2015). Indeed, anthropogenic eutrophication and temperature increase are all factors that can weaken the immune system of the benthic organisms and lead to a decrease in their resistance to parasitic protists asPerkinsus spp. (Morley, 2010). Furthermore, the host range of these parasitic organisms might be wider than described in the scientific literature with the existence of other “non-commercial” potential hosts belonging to suborders infected byPerkinsus spp. Thus, even though many arguments allow classifying thePerkinsus species as a BHR parasite, we cannot determine the real consequences on stocks of “non-optimal” hosts especially when it is related to sporadic infections with low prevalence. Due to research focus on economically valuable species of mollusks likeC. virginica orR. philippinarum, consequences of these infections on the global host range ofPerkinsus spp. are still mainly undiscovered.

Parviluciferaceae, a Group of Parasites of Microalgae

Norén et al. (1999) identified small round structures abnormally present within the toxic dinoflagellateDinophysis spp. on the west coast of Sweden. Molecular and microscopic analyses revealed that this parasitic organism was affiliated to the Perkinsids group and erected a new phylum, the Perkinsea (Norén et al., 1999). So far, five genera have been described and cultivated (Jeon et al., 2018). The first genus described wasParvilucifera genus encompassingP. infectans (Norén et al., 1999),P. multicavata (Jeon et al., 2018),P. sinerae (Figueroa et al., 2008),P. rostrata (Lepelletier et al., 2014b),P. corolla (Reñé et al., 2017b),P. catillosa, andParvilucifera sp. (Alacid et al., 2020). Recently, three new genera were described, the genusSnorkelia containingP. prorocentri (Leander and Hoppenrath, 2008) renamedSnorkelia prorocentri (Reñé et al., 2017a), the genusDinovorax includingD. pyriformis (Reñé et al., 2017a),Tuberlatum encompassingT. coatsi (Jeon et al., 2018), and finallyMaranthos includingM. nigrum (Reñé et al., 2021a). All these genera seem to have a basal position within the Parviluciferaceae group, with the exception of theMaranthos species, whose phylogenetic position is not yet clear (Jeon et al., 2018;Reñé et al., 2021a;Figures 1B,2).

Members of the Parviluciferaceae family share many common traits including development and characteristic of their life cycle, and their infection strategy (Reñé et al., 2017a). The infection begins with the entry of the zoospore into the dinoflagellate host cell (Figure 4B). Once inside, the parasite develops in the cytoplasm forming a spherical cell, the trophont (= trophocyte), which consumes its host for its own growth. At the end of the feeding stage, a free-living non-motile cell, the sporocyte (= immature sporangium), occupying the whole intracellular space, is released by breaking the theca of the host. Next, the cell undergoes morphological transformations into a spherical multinucleated dark structure called the sporangium (= mature sporocyte). The sporangium remains in dormancy until the detection of a host chemical signal (Garcés et al., 2013b). After a short maturation (∼48 h), a sporangium with open aperture(s) releases several hundred new infectious cells (zoospores) (Alacid et al., 2017). To date, all described species are parasites of dinoflagellates including toxic species and possess a BHR (here belonging to a “true” generalist class) (Figueroa et al., 2008;Garcés et al., 2013a;Jeon et al., 2018;Rodríguez and Figueroa, 2020;Reñé et al., 2021b) exceptS. prorocentri, isolated from Boundary Bay (Canada), which was known to only infect the marine benthic dinoflagellateProrocentrum fukuyoi (Leander and Hoppenrath, 2008;Figure 1B). However, in 2017, Reñé et al. isolated a new species ofSnorkelia from a different location (Catalan Coast, NW Mediterranean) infecting the dinoflagellateLevanderina fissa (Reñé et al., 2017a). The host range ofSnorkelia species, therefore, remains enigmatic. For the “true” generalists,P. sinerae,P. corolla, andP. rostrata, the host range was determined mainly byin vitro cross-infection.In vitro experiments could result in artificial linkage between a host species and a parasite, which is not representative of the ecological reality (Råberg et al., 2014;Alacid et al., 2016;Reñé et al., 2021b). Infection of non-preferred hosts could be based on chemical and physiological processes shared by several related hosts (e.g.,Garcés et al., 2013b). Furthermore, when a parasite infects a host but is not able to produce viable parasitic cells, such “host” cannot be considered as a “true” host for the parasite, e.g., infection of a chlorophyte strain ofPyramimonas byP. corolla (Rodríguez and Figueroa, 2020). The infection process is probably the result of plesiomorphic mechanisms amongP. corolla’s hosts, but the parasite cannot achieve its whole life cycle. In this review, we consideredParvilucifera species as generalists when they realize a viable life cycle in a large repertoire of host species. A BHR gives them the full potential of host shifting in new environments, although there are evidences of host preference in nature (Alacid et al., 2016). However, much remains to be done to evaluate the contribution of the plesiomorphic and convergent traits of a parasite to the success of generalists.

Despite a “hot-spot” of detection in Europe, the distribution of these parasites of dinoflagellates is worldwide in marine environments (Figure 3B). As major contributors, with diatoms, to the fixation of inorganic carbon through photosynthesis (Falkowski et al., 1998), dinoflagellates are one of the most important components of marine phytoplankton as primary producers and grazers.Parvilucifera parasites could participate in regulating dinoflagellate populations in the blooming period, as shown in Spain where 5 to 18% of the population of the noxious dinoflagellates,Alexandrium minutum, were killed byParvilucifera sp. during natural bloom (Alacid et al., 2017). For methodological limitation reasons, most studies describingParvilucifera spp. have been carried out when prevalence is the highest on blooming host species, thus, increasing the chances to detect and describe new parasitic species. Unfortunately, this bias hides a possible important part of host and parasite diversity (Reñé et al., 2021b). Indeed, in some studies, the host range of these parasites appears to be much wider, including non-harmful species (e.g.,Figueroa et al., 2008;Garcés et al., 2013a;Rodríguez and Figueroa, 2020), even if thein vitro specificity tests do not take into account the complexity of planktonic interaction network and thein situ parasitic host preferences. This lack of knowledge onParvilucifera distribution, diversity, and trophic niche might become problematic, given its BHR nature and the global trades. Indeed, ballast waters, which are one of the most important vectors of introduction in marine systems (Carlton, 1985;Barry et al., 2008), could contribute to the dispersal ofParvilucifera trophonts or dormant sporangia (e.g.,Darling et al., 2018), thus, increasing the risk of settlement in new areas. Hence, introductions of these BHR parasites could act positively as a top–down control of the toxic dinoflagellate blooming species and, conversely, could play a key role in deregulating the planktonic compartment given the importance of dinoflagellates as primary producers inducing significant shifts in marine food webs.

Xcellidae, a New Group of Fish Parasites

In 1969, Brooks et al. of two flatfish species, the black-tipped flounder,Psettichthys melanostictus, and the Pacific plaice,Platichthys stellatus (Brooks et al., 1969). Thirty-five years later, a protozoan named X-cell was identified as responsible for this pathology inducing also multiple lesions and swelling of the gill filaments (Miwa et al., 2004). Using small and large concatenated subunit (SSU and LSU) rDNA phylogeny reconstruction,Freeman et al. (2017) investigated the phylogenetic position of this enigmatic X-cell protist and showed that it formed two highly distinct clades, one corresponding to the pseudobranchial parasites of Gadiformes, and the other to gill and epidermal X-cells from Perciforms and Pleuronectiformes calledGadixcellia andXcellia, respectively. More recently,Karlsbakk et al. (2021) described a new genus namedSalmoxcellia, a sister group toGadixcellia. Despite an unusually high genetic divergence between the three genera, with a similarity of 74.9% for the SSU rDNA sequences, these two close sister clades form a new family called Xcellidae (or Xcellins) branching within the Perkinsea lineage (Freeman et al., 2017;Karlsbakk et al., 2021) (Figure 2B).

Histological examination and scanning electron microscopy of xenomas of infected fish revealed a large number of clustered round parasitic cells surrounded by host connective tissue. Currently, only one developmental life stage has been described up to today; hence, the life cycle of this parasite still remains unknown (Freeman et al., 2017). However,Freeman et al. (2017) suggested that infection occursvia contact between fish and the benthos. Indeed, X-cell infection occurs in fish species with at least one benthic stage during their life cycle (Fahay, 1983;Lough et al., 1989). An experimental infectionin situ in tanks revealed that only fish exposed, even transiently, to the benthos were infected and showed symptoms of disease (Eydal et al., 2010). The infected fish samples are mainly young and adult individuals. Indeed, the first pathogenic symptoms (pseudobranchial xenomas) appear macroscopically in young wild cod (around 6 months old, 6- to 13-cm length) from Icelandic waters with a prevalence peak of 23% at the age of 22 months, whereas older or larger fish (40- to 76-cm length) have a lower prevalence around 7 and 1%, respectively (Eydal et al., 2010). Similar trends had been observed in Atlantic cod (Morrison et al., 1982) and the Pacific cod (Stich et al., 1976). However, the question remains open whether the mortality induced by the X-cell infectious agent on juveniles could remove an age class from the field sampling and bias the observed prevalence. Although mortality induced by these infectious agents still needs to be demonstrated, the transfer of wild infected individuals ofGadus morhua in tanks has shown an important mortality (Eydal et al., 2010).

Parasites belonging to these three genera have been detected in more than 20 fish species belonging to five orders of teleost: Pleuronectiformes (flatfishes) (Freeman, 2009;Freeman et al., 2011), Perciformes (perch-like fish) (Katsura et al., 1984), Gadiformes (cods) (Freeman et al., 2011), Siluriformes (catfishes) (Diamant et al., 1994), and Salmoniformes (salmonids) (Dyková et al., 1993;Karlsbakk et al., 2021). Until now, five species have been described (Figure 1B). The parasiteXcellia pleuronecti infection has been confirmed inHippoglossoides dubius andPseudopleuronectes obscurus but also suspected in many other species from the same area:Cleisthenes herzensteini,C. pinetorum,Glyptocephalus stelleri,Kareius bicoloratus,Hippoglossoides elassodon,Liopsetta pinnifasciata,Platichthys stellatus,Parophrys vetulus,Pseudopleuronectes schrenki, andVerasper moseri. Equally,Xcellia lamelliphila infection is detected inLimanda limanda,Lycodes spp.,Macruronus novaezelandiae,Merluccius gayi gayi, andTrematomus spp. (Freeman et al., 2017). Finally, the newly described parasiteSalmoxcellia vastator is detected inOncorhynchus mykiss and inSalmo salar (Karlsbakk et al., 2021). These three parasites are, therefore, BHR parasites (Figure 1B). Conversely,Xcellia gobii andGadixcellia gadi infecting, respectively,Acanthogobius flavimanus andGadus morhua are NHR parasites and classified as specialists (Freeman et al., 2017;Figure 1B). This classification will certainly evolve as new knowledge on Xcellidae group becomes available.

In addition, Xcellidae protists have been detected in a restricted geographical area (Northern Europe and Japan) (Figure 3C), but the monitoring of these X-cell infections is hard to carry out and stay largely incomplete because of migratory fish hosts in the worldwide ocean. Their host range is mainly unknown, but we can hypothesize that it may be broader than that currently identified. An extended host range would not be surprising considering the Perkinsea phylum constituted of a majority of BHR parasites closely related phylogenetically (e.g., the Perkinsidae) (Figure 2B). Overall, the consequences of the X-cell infection on fish populations are not well known. However, as it affects important halieutic resources as described for the salmonid parasite,Salmoxcellia vastator, it makes farmed salmonid fillets unsuitable for sale (Karlsbakk et al., 2021). Moreover, these parasites could infect vulnerable species, such as Atlantic cod fishG. morhua, extensively farmed, but mainly in the north of Europe (seeFAO, 2021 Programme—Gadus morhua). Commercially important fish, a concern of the global trades, which could act as reservoirs allowing the dissemination in farmed and wild fish populations.

Severe Perkinsea Infection, an Infectious Agent of Tadpoles Populations

With up to 50% of all threatened species, amphibian populations are emblematic representatives of the sixth mass extinction event to which highly virulent wildlife diseases contribute (Stuart et al., 2004;Gewin, 2008;Chambouvet et al., 2020). Recent works have mainly identified two pathogens as drivers of this decline, the fungusBatrachochytrium dendrobatidis and the Ranavirus (e.g.,Bosch et al., 2001;Fisher et al., 2009;Gray et al., 2009;Olson et al., 2013).

However, in April 2006, in a pond in northeast Georgia (United States),Davis et al. (2007) observed a massive mortality event of southern leopard tadpoles,Rana sphenocephala (syn.Lithobates sphenocephalus), attributed to an unknown protist affiliated by phylogenetic analysis to the Perkinsea lineage. Using phylogenetic analysis, it has been shown that this organism belongs to a discrete clade named pathogenic Perkinsea clade (PPC) (Isidoro-Ayza et al., 2017) within the wider monophylic cluster of the Novel Alveolate Group 01 (NAG01) (Chambouvet et al., 2015) obtained from disparate freshwater environments or internal organs (e.g., liver tissues) of a wide variety of Neobatrachia suborder (Chambouvet et al., 2015;Isidoro-Ayza et al., 2017;Figures 1B,2B). Infectious agents of the PPC clade have been identified as responsible for the die-offs of tadpole throughout the United States (Isidoro-Ayza et al., 2017). Infected tadpoles with pathological symptoms, named “severe Perkinsea infection” (SPI), showed lethargic swimming with enlarged and histopathologic lesions of the liver, mesonephros, spleen, pancreas, gills, gastrointestinal tract, skeletal muscle, dermis, and peritoneum (Green et al., 2002;Davis et al., 2007;Jones et al., 2012;Isidoro-Ayza et al., 2017). Histological examination of infected liver tissues revealed a massive number of round infective cells replacing the normal hepatic parenchyma. Two distinct putative life stages were identified invading the hepatocyte cells corresponding to hypnospore-like and trophozoite-like life stages (Isidoro-Ayza et al., 2017,2019). SPI symptoms have been reported from summer to early autumn in boreal and temperate regions, and from late winter to early spring in subtropical areas (Isidoro-Ayza et al., 2017;Figure 3D). Although this parasite is mainly described as infecting tadpole life stage, one report highlights infection of the adult populations with granulomatous lesion in the legs (Jones et al., 2012). These results suggest that either most of the tadpoles die before the metamorphosis or that mature immune systems acquired after the metamorphosis may drive back the infection (Isidoro-Ayza et al., 2017). Although the relationship between the infectious agent and the disease is not yet well established, symptoms of the disease could result from co-infection between the Perkinsea parasite and others infectious agents, such as the FV3-like virus, responsible for frog population declines, which infects both larval and adult amphibians (Gray et al., 2009;Lesbarrères et al., 2012), and/or alteration of tadpole immune systems (Isidoro-Ayza et al., 2017).

During monitored SPI outbreaks, mortality rate can reach up to 95% of the population leading to the loss of an entire age class or, in case of chronic infection, to a reduced recruitment (Green et al., 2002;Isidoro-Ayza et al., 2017,2019). Now recognized as the third most common infectious disease of anuran species, SPI infectious agent is described as an emerging pathogen. It is therefore now fundamental to understand the relationship between this infectious agent and the outcome of the disease and how other pathogen communities and/or environmental factors could affect disease susceptibility. Furthermore, as the globalization of trade where amphibian species, generally involved in meat or pet trade, could spread the parasitic invadersvia reservoir species into native and/or naive amphibian populations, the study of this pathogen is crucial.

Environmental Diversity: Unveiling Putative Pathogens?

Molecular methodologies have recently highlighted that our vision of Perkinsea diversity, mainly assessed by culture-based methods, is still only the tip of the iceberg. In both marine and freshwater environments, analysis of genetic diversity based on SSU rDNA sequences of the smallest sizes of plankton (<5 μm), revealed a previously unknown diversity of these organisms (Moon-van der Staay et al., 2001;López-García et al., 2001,2003;Lefranc et al., 2005;Mangot et al., 2009).

A recent overview of freshwater Perkinsea genetic diversity, especially in lakes, revealed that this group appears to be diverse and abundant, with an observation of 2,927 operational taxonomic units (OTUs) defined at 95% of sequence similarity, which represented 3% of the eukaryotic diversity analyzed inDebroas et al. (2017), suggesting an important role in the trophic web (e.g.,Lepère et al., 2008,2011;Jobard et al., 2020). Recent studies have highlighted that freshwater clades could be a parasitic protist of the colonial green algaeSphaerocystis sp. (Jobard et al., 2020). These results are consistent with the description ofRastrimonas gen. nov. (Brugerolle, 2003), previously described asCryptophagus subtilis, infecting the free-living CryptophyteChilomonas paramaecium (Brugerolle, 2002). However, apart from these two descriptive publications, no molecular analysis has yet been carried out to definitively affiliateRastrimonas sp. within the Perkinsea lineage.

In marine environments, molecular signatures of Perkinsea were also found in extreme environments, such as hydrothermal vents or anoxic fjords (López-García et al., 2003;Zuendorf et al., 2006), but most of environmental genomics studies target water column samples [e.g., surface or deep chlorophyll maximum (DCM) depths] (e.g.,Moon-van der Staay et al., 2001;López-García et al., 2003;De Vargas et al., 2015). This lack of genetic signatures in marine environmental surveys represent a real paradox, since the two main cultivable groups described,Perkinsus spp. andParvilucifera spp., are marine. In 2014, targeting the V4 hypervariable region of the rDNA and rRNA templates using 454 sequencing technology,Chambouvet et al. (2014) evaluated the genetic diversity of the eukaryotic microbial community in two sampling depths (surface and DCM) and in the sediment across four different locations across Europe (Oslo, Norway; Naples, Italy; Barcelona, Spain; Roscoff and France). The analysis revealed an unexpected genetic diversity of ribosomally active organisms belonging to Perkinsea other than thanParviluciferaceae andPerkinsus clusters mainly detected in the sediment with a total of 265 sequences clustered in 150 OTUs defined at 99% similarity. The ribosomal RNA sequences present in these clades belong to metabolically active organisms, which certainly play an active role in the ecosystem functioning. This cryptic diversity, only detected by their genetic signatures, raises new scientific questions about the putative existence of non-parasitic heterotrophic organisms, the host range of parasitic Perkinsea, and their role and impact on the aquatic food web. Finally, these results suggested that sediments can act as parasite reservoirs, as suggested forAmoebophrya parasitoids (Chambouvet et al., 2011) and for marine Perkinsea (Chambouvet et al., 2014;Freeman et al., 2017;Reñé et al., 2021b). However, studies on diversity and distribution are scarce and restricted to specific environments (e.g., coastal water or few freshwater environments). A wider study of the Perkinsea distribution could give a more complete picture of environmental distribution unveiling potential reservoirs of pathogens. A phylogenetic analysis of Perkinsea illustrates two main evolutionary divisions between clades associated to Perkinsidae and Xcellidae, and Parviluciferaceae prior to marine/freshwater specialization (Jobard et al., 2020). This also suggests that transitions between marine and freshwater were few in the life history of Perkinsea, as differences between these environments constitute a barrier for cross-colonization for most organisms (Logares et al., 2009;Bråte et al., 2010). However, a deep analysis of environmental DNA may reveal new information on the evolution and environmental colonization of Perkinsea.

1. Agosta S. J., Janz N., Brooks D. R. (2010). How specialists can be generalists: resolving the “parasite paradox” and implications for emerging infectious disease.Zoologia27151–162. 10.1590/S1984-46702010000200001 [DOI] [Google Scholar]
2. Alacid E., Park M. G., Turon M., Petrou K., Garcés E. (2016). A game of russian roulette for a generalist dinoflagellate parasitoid: host susceptibility is the key to success.Front. Microbiol.7:769. 10.3389/fmicb.2016.00769 [DOI] [PMC free article] [PubMed] [Google Scholar]
3. Alacid E., Reñé A., Camp J., Garcés E. (2017).In situ occurrence, prevalence and dynamics ofParvilucifera parasitoids during recurrent blooms of the toxic dinoflagellateAlexandrium minutum.Front. Microbiol.8:1624. 10.3389/fmicb.2017.01624 [DOI] [PMC free article] [PubMed] [Google Scholar]
4. Alacid E., Reñé A., Gallisai R., Paloheimo A., Garcés E., Kremp A. (2020). Description of two new coexisting parasitoids of blooming dinoflagellates in the Baltic sea:Parvilucifera catillosa sp. nov. andParvilucifera sp. (Perkinsea, Alveolata).Harmful Algae100:101944. 10.1016/j.hal.2020.101944 [DOI] [PubMed] [Google Scholar]
5. Alacid E., Reñé A., Garcés E. (2015). New insights into the parasitoidParvilucifera sinerae life cycle: the development and kinetics of infection of a bloom-forming dinoflagellate host.Protist166, 677–699. 10.1016/j.protis.2015.09.001 [DOI] [PubMed] [Google Scholar]
6. Allam B., Carden W. E., Ward J. E., Ralph G., Winnicki S., Pales Espinosa E. (2013). Early host-pathogen interactions in marine bivalves: evidence that the alveolate parasitePerkinsus marinus infects through the oyster mantle during rejection of pseudofeces.J. Invertebr. Pathol.11326–34. 10.1016/j.jip.2012.12.011 [DOI] [PubMed] [Google Scholar]
7. Anderson R. S., Unger M. A., Burreson E. M. (1996). Enhancement ofPerkinsus marinus disease progression in TBT-exposed oysters (Crassostrea virginica).Mar. Environ. Res.42177–180. 10.1016/0141-1136(95)00066-6 [DOI] [Google Scholar]
8. Andrews J. (1996). History ofPerkinsus marinus, A pathogen of oysters in Chesapeake Bay 1950-1984.VIMS Articles503:5. [Google Scholar]
9. Arzul I., Carnegie R. B. (2015). New perspective on the haplosporidian parasites of molluscs.J. Invertebr. Pathol.13132–42. 10.1016/j.jip.2015.07.014 [DOI] [PubMed] [Google Scholar]
10. Arzul I., Chollet B., Boyer S., Bonnet D., Gaillard J., Baldi Y., et al. (2014). Contribution to the understanding of the cycle of the protozoan parasiteMarteilia refringens.Parasitology141227–240. 10.1017/S0031182013001418 [DOI] [PubMed] [Google Scholar]
11. Arzul I., Chollet B., Michel J., Robert M., Garcia C., Joly J.-P., et al. (2012). OnePerkinsus species may hide another: characterization ofPerkinsus species present in clam production areas of France.Parasitology1391757–1771. 10.1017/S0031182012001047 [DOI] [PubMed] [Google Scholar]
12. Auzoux-Bordenave S., Vigario A. M., Ruano F., Domart-Coulon I., Doumenc D. (1996).In vitro sporulation of the clam pathogenPerkinsus atlanticus (Apicomplexa, Perkinsea) under various environmental conditions.Oceanogr. Lit. Rev.9:926. [Google Scholar]
13. Azevedo C. (1989). Fine structure ofPerkinsus atlanticus n. sp. (Apicomplexa, Perkinsea) parasite of the clamRuditapes decussatus from Portugal.J. Parasitol.75627–635. 10.2307/3282915 [DOI] [PubMed] [Google Scholar]
14. Barber B. J., Mann R. L. (1994). Growth and mortality of Eastern oysters,Crassostrea virginica (Gmelin, 1791), and Pacific oysters, Crassostrea gigas (Thunberg, 1793) under challenge from the parasite,Perkinsus marinus.J. Shell. Res.13:7. [Google Scholar]
15. Barry S. C., Hayes K. R., Hewitt C. L., Behrens H. L., Dragsund E., Bakke S. M. (2008). Ballast water risk assessment: principles, processes, and methods.ICES J. Mar. Sci.65121–131. 10.1093/icesjms/fsn004 [DOI] [Google Scholar]
16. Bass D., Stentiford G. D., Littlewood D. T. J., Hartikainen H. (2015). Diverse applications of environmental DNA methods in parasitology.Trends Parasitol.31499–513. 10.1016/j.pt.2015.06.013 [DOI] [PubMed] [Google Scholar]
17. Bass D., Stentiford G. D., Wang H.-C., Koskella B., Tyler C. R. (2019). The pathobiome in animal and plant diseases.Trends Ecol. Evol.34996–1008. 10.1016/j.tree.2019.07.012 [DOI] [PMC free article] [PubMed] [Google Scholar]
18. Bjorbækmo M. F. M., Evenstad A., Røsæg L. L., Krabberød A. K., Logares R. (2020). The planktonic protist interactome: where do we stand after a century of research?ISME J.14544–559. 10.1038/s41396-019-0542-5 [DOI] [PMC free article] [PubMed] [Google Scholar]
19. Blackbourn J., Bower S. M., Meyer G. R. (1998).Perkinsus qugwadi sp.nov. (incertae sedis), a pathogenic protozoan parasite of Japanese scallops,Patinopecten yessoensis, cultured in British Columbia, Canada.Can. J. Zool.76942–953. 10.1139/z98-015 [DOI] [Google Scholar]
20. Bosch J., Martıìnez-Solano I., Garcıìa-Parıìs M. (2001). Evidence of a chytrid fungus infection involved in the decline of the common midwife toad (Alytes obstetricans) in protected areas of central Spain.Biol. Conserv.97331–337. 10.1016/S0006-3207(00)00132-4 [DOI] [Google Scholar]
21. Bråte J., Logares R., Berney C., Ree D. K., Klaveness D., Jakobsen K. S., et al. (2010). FreshwaterPerkinsea and marine-freshwater colonizations revealed by pyrosequencing and phylogeny of environmental rDNA.ISME J.41144–1153. 10.1038/ismej.2010.39 [DOI] [PubMed] [Google Scholar]
22. Brooks R. E., McArn G. E., Wellings S. R. (1969). Ultrastructural observations on an unidentified cell type found in epidermal tumors of flounders.J. Natl. Cancer Inst.4397–109. 10.1093/jnci/43.1.97 [DOI] [PubMed] [Google Scholar]
23. Brugerolle G. (2002).Cryptophagus subtilis: a new parasite of cryptophytes affiliated with thePerkinsozoa lineage.Eur. J. Protistol.37379–390. 10.1078/0932-4739-00837 [DOI] [Google Scholar]
24. Brugerolle G. (2003). Apicomplexan parasiteCryptophagus renamedRastrimonas gen. nov.Eur. J. Protistol.39:101. 10.1078/0932-4739-00910 [DOI] [Google Scholar]
25. Burreson E., Alvarez R., Martinez V., Macedo L. (1994).Perkinsus marinus (Apicomplexa) as a potential source of oysterCrassostrea virginica mortality in coastal lagoons of Tabasco, Mexico.Dis. Aquat. Org.2077–82. 10.3354/dao020077 [DOI] [Google Scholar]
26. Burreson E., Stokes N., Friedman C. (2000). Increased virulence in an introduced pathogen:Haplosporidium nelsoni (MSX) in the eastern oysterCrassostrea virginica.J. Aquat. Anim. Health121–8. [DOI] [PubMed] [Google Scholar]
27. Cáceres-Martínez J., Ortega M. G., Vásquez-Yeomans R., García T., de J. P., Stokes N. A., et al. (2012). Natural and cultured populations of the mangrove oysterSaccostrea palmula from Sinaloa, Mexico, infected byPerkinsus marinus.J. Invertebr. Pathol.110321–325. 10.1016/j.jip.2012.03.019 [DOI] [PubMed] [Google Scholar]
28. Calvo G. W., Luckenbach M., Burreson E. M. (1999).A Comparative Field Study of Crassostrea gigas and Crassostrea virginica in Relation to Salinity in Virginia. Special Reports in Applied Marine Science and Ocean Engineering (SRAMSOE) No. 349.Gloucester Point, VA: Virginia Institute of Marine Science, William & Mary. 10.21220/V5PT7H [DOI] [Google Scholar]
29. Cariton J. T., Geller J. B. (1993). Ecological roulette: the global transport of nonindigenous marine organisms.Science26178–82. 10.1126/science.261.5117.78 [DOI] [PubMed] [Google Scholar]
30. Carlton J. T. (1985). Transoceanic and interoceanic dispersal of coastal marine organisms: the biology of ballast water.Oceanogr. Mar. Biol.23313–371. [Google Scholar]
31. Carnegie R. B., Arzul I., Bushek D. (2016). Managing marine mollusc diseases in the context of regional and international commerce: policy issues and emerging concerns.Phil. Trans. R. Soc. B371:20150215. 10.1098/rstb.2015.0215 [DOI] [PMC free article] [PubMed] [Google Scholar]
32. Carnegie R. B., Ford S. E., Crockett R. K., Kingsley-Smith P. R., Bienlien L. M., Safi L. S. L., et al. (2021). A rapid phenotype change in the pathogenPerkinsus marinus was associated with a historically significant marine disease emergence in the eastern oyster.Sci. Rep.11:12872. 10.1038/s41598-021-92379-6 [DOI] [PMC free article] [PubMed] [Google Scholar]
33. Carrasco N., Andree K. B., Lacuesta B., Roque A., Rodgers C., Furones M. D. (2012). Molecular characterization of theMarteilia parasite infecting the common edible cockleCerastoderma edule in the Spanish Mediterranean coast.Aquaculture324–32520–26. 10.1016/j.aquaculture.2011.10.017 [DOI] [Google Scholar]
34. Chambouvet A., Alves-de-Souza C., Cueff V., Marie D., Karpov S., Guillou L. (2011). Interplay between the parasiteAmoebophrya sp.(Alveolata) and the cyst formation of the red tide dinoflagellateScrippsiella trochoidea.Protist162637–649. [DOI] [PubMed] [Google Scholar]
35. Chambouvet A., Berney C., Romac S., Audic S., Maguire F., De Vargas C., et al. (2014). Diverse molecular signatures for ribosomally ‘active’Perkinsea in marine sediments.BMC Microbiol.14:110. 10.1186/1471-2180-14-110 [DOI] [PMC free article] [PubMed] [Google Scholar]
36. Chambouvet A., Gower D. J., Jirků M., Yabsley M. J., Davis A. K., Leonard G., et al. (2015). Cryptic infection of a broad taxonomic and geographic diversity of tadpoles byPerkinsea protists.Proc. Natl. Acad. Sci. U.S.A.112E4743–E4751. 10.1073/pnas.1500163112 [DOI] [PMC free article] [PubMed] [Google Scholar]
37. Chambouvet A., Morin P., Marie D., Guillou L. (2008). Control of toxic marine dinoflagellate blooms by serial parasitic killers.Science3221254–1257. [DOI] [PubMed] [Google Scholar]
38. Chambouvet A., Smilansky V., Jirků M., Isidoro-Ayza M., Itoïz S., Derelle E., et al. (2020). Diverse alveolate infections of tadpoles, a new threat to frogs?PLoS Path.16:e1008107. 10.1371/journal.ppat.1008107 [DOI] [PMC free article] [PubMed] [Google Scholar]
39. Chintala M., Bushek D., Ford S. (2002). Comparison ofin vitro-cultured and wild-typePerkinsus marinus. II. Dosing methods and host response.Dis. Aquat. Org.51203–216. 10.3354/dao051203 [DOI] [PubMed] [Google Scholar]
40. Choi K.-S., Park K.-I. (2005).Current Status of Perkinsus Infection in Korean Waters Diseases in Asian Aquaculture V. Manila: Fish Health Section, Asian Fisheries Society, 263–274. [Google Scholar]
41. Choi K.-S., Park K.-I. (2010). “Review on the protozoan parasitePerkinsus olseni (Lester and Davis, 1981) infection in Asian waters,” inCoastal Environmental and Ecosystem Issues of the East China Sea, edsIsimatsu A., Lie H. J. (Nagasaki: Nagasaki University; ), 269–281. [Google Scholar]
42. Cleaveland S., Laurenson M. K., Taylor L. H. (2001). Diseases of humans and their domestic mammals: pathogen characteristics, host range and the risk of emergence.Phil. Trans. R. Soc. B356991–999. 10.1098/rstb.2001.0889 [DOI] [PMC free article] [PubMed] [Google Scholar]
43. Coats D. W., Park M. G. (2002). Parasitism of photosynthetic dinoflagellates by three strains ofAmoebophrya (Dinophyta): Parasite survival, infectivity, generation time, and host specificity.J. Phycol.38520–528. 10.1046/j.1529-8817.2002.01200.x [DOI] [Google Scholar]
44. Coen L., Grizzle R. (2007). The importance of habitat created by molluscan shellfish to managed species along the Atlantic Coast of the United States.Habitat. Manag. Ser.81–108. [Google Scholar]
45. Cohen A. N. (1998). Accelerating invasion rate in a highly invaded estuary.Science279555–558. 10.1126/science.279.5350.555 [DOI] [PubMed] [Google Scholar]
46. Coss C. A., Robledo J. A. F., Ruiz G. M., Vasta G. R. (2001). Description ofPerkinsus andrewsi n. sp. isolated from the Baltic clam (Macoma balthica) by characterization of the ribosomal RNA locus, and development of a species-specific PCR-based diagnostic assay.J. Eukaryot. Microbiol.4852–61. 10.1111/j.1550-7408.2001.tb00415.x [DOI] [PubMed] [Google Scholar]
47. Cremonte F., Balseiro P., Figueras A. (2005). Occurrence ofPerkinsus olseni (Protozoa: Apicomplexa) and other parasites in the venerid commercial clamPitar rostrata from Uruguay, southwestern Atlantic coast.Dis. Aquat. Org.6485–90. 10.3354/dao064085 [DOI] [PubMed] [Google Scholar]
48. da Silva P. M., Scardua M. P., Vianna R. T., Mendonça R. C., Vieira C. B., Dungan C. F., et al. (2014). TwoPerkinsus spp. infectCrassostrea gasar oysters from cultured and wild populations of the Rio São Francisco estuary, Sergipe, northeastern Brazil.J. Invertebr. Pathol.11962–71. 10.1016/j.jip.2014.04.005 [DOI] [PubMed] [Google Scholar]
49. da Silva P. M., Vianna R. T., Guertler C., Ferreira L. P., Santana L. N., Fernández-Boo S., et al. (2013). First report of the protozoan parasitePerkinsus marinus in South America, infecting mangrove oystersCrassostrea rhizophorae from the Paraíba River (NE, Brazil).J. Invertebr. Pathol.11396–103. 10.1016/j.jip.2013.02.002 [DOI] [PubMed] [Google Scholar]
50. Dairain A., Legeay A., de Montaudouin X. (2019). Influence of parasitism on bioturbation: from host to ecosystem functioning.Mar. Ecol. Prog. Ser.619201–214. 10.3354/meps12967 [DOI] [Google Scholar]
51. Dantas Neto M. P., Gesteira T. C. V., Sabry R. C., Feijó R. G., Forte J. M., Boehs G., et al. (2016). First record of Perkinsus chesapeaki infectingCrassostrea rhizophorae in South America.J. Invertebr. Pathol.14153–56. 10.1016/j.jip.2016.10.007 [DOI] [PubMed] [Google Scholar]
52. Darling J. A., Martinson J., Gong Y., Okum S., Pilgrim E., Lohan K. M. P., et al. (2018). Ballast water exchange and invasion risk posed by intracoastal vessel traffic: an evaluation using high throughput sequencing.Environ. Sci. Technol.529926–9936. 10.1021/acs.est.8b02108 [DOI] [PMC free article] [PubMed] [Google Scholar]
53. Daszak P. (2000). Emerging infectious diseases of wildlife – Threats to biodiversity and human health.Science287443–449. 10.1126/science.287.5452.443 [DOI] [PubMed] [Google Scholar]
54. Davis A. K., Yabsley M. J., Kevin Keel M., Maerz J. C. (2007). Discovery of a novel alveolate pathogen affecting southern leopard frogs in Georgia: description of the disease and host effects.EcoHealth4310–317. 10.1007/s10393-007-0115-3 [DOI] [Google Scholar]
55. de Montaudouin X., Labarraque D., Giraud K., Bachelet G. (2001). Why does the introduced gastropodCrepidula fornicata fail to invade Arcachon Bay (France)?J. Mar. Biol. Ass.8197–104. 10.1017/S0025315401003447 [DOI] [Google Scholar]
56. De Vargas C., Audic S., Henry N., Decelle J., Mahe F., Logares R., et al. (2015). Eukaryotic plankton diversity in the sunlit ocean.Science348:1261605. 10.1126/science.1261605 [DOI] [PubMed] [Google Scholar]
57. Debroas D., Domaizon I., Humbert J.-F., Jardillier L., Lepère C., Oudart A., et al. (2017). Overview of freshwater microbial eukaryotes diversity: a first analysis of publicly available metabarcoding data.FEMS Microbiol. Ecol.93:fix023. 10.1093/femsec/fix023 [DOI] [PubMed] [Google Scholar]
58. Desdevises Y., Morand S., Legendre P. (2002). Evolution and determinants of host specificity in the genusLamellodiscus (Monogenea): Determinants of specificity inLamellodiscus.Biol. J. Linn. Soc.77431–443. 10.1046/j.1095-8312.2002.00114.x [DOI] [Google Scholar]
59. Diamant A., Fournie J., Courtney L. (1994). X-cell pseudotumors in a hardhead catfish Arius fells (Ariidae) from Lake Pontchartrain, Louisiana, USA.Dis. Aquat. Org.18181–185. 10.3354/dao018181 [DOI] [Google Scholar]
60. Dittman D. E., Ford S. E., Padillai D. K. (2001). Effects ofPerkinsus marinus on reproduction and condition of the eastern oyster,Crassostrea virginica, depend on timing.J. Shell. Res.201025–1034. [Google Scholar]
61. Dungan C. F., Reece K. S., Moss J. A., Hamilton R. M., Diggles B. K. (2007). Perkinsus olseniin vitro isolates from the New Zealand clamAustrovenus stutchburyi.J. Eukaryot. Microbiol.54263–270. 10.1111/j.1550-7408.2007.00265.x [DOI] [PubMed] [Google Scholar]
62. Dyková I., Figueras A., Novoa B. (1993). X-cell lesions in the liver of coho salmon Oncorhynchus kisutch.Dis. Aquat. Org.15171–174. [Google Scholar]
63. Elston R. A., Farley C., Kent M. (1986). Occurence and significance of bonamiasis in European flat oystersOstrea edulis in North America.Dis. Aquat. Org.249–54. 10.3354/dao002049 [DOI] [Google Scholar]
64. Engelsma M., Culloty S., Lynch S., Arzul I., Carnegie R. (2014). Bonamia parasites: a rapidly changing perspective on a genus of important mollusc pathogens.Dis. Aquat. Org.1105–23. 10.3354/dao02741 [DOI] [PubMed] [Google Scholar]
65. Enríquez-Espinoza T., Grijalva-Chon J., Castro-Longoria R., Ramos-Paredes J. (2010).Perkinsus marinus inCrassostrea gigas in the Gulf of California.Dis. Aquat. Org.89269–273. 10.3354/dao02199 [DOI] [PubMed] [Google Scholar]
66. Eydal M., Kristmundsson Á, Bambir S. (2010).Pseudobranchial X-cell pseudotumors in young wild and farmed Atlantic codGadus morhua in Iceland.Dis. Aquat. Org.9183–88. 10.3354/dao02248 [DOI] [PubMed] [Google Scholar]
67. Fahay M. P. (1983). Guide to the early stages of marine fishes occurring in the western North Atlantic Ocean, Cape Hatteras to the southern Scotian Shelf.J. Northwest Atl. Fish. Sci.4:423. [Google Scholar]
68. Falkowski P. G., Barber R. T., Smetacek V. (1998). Biogeochemical controls and feedbacks on ocean primary production.Science281200–206. [DOI] [PubMed] [Google Scholar]
69. FAO (2021).FAO Fisheries & Aquaculture – Cultured Aquatic Species Information Programme - Gadus morhua (Linnaeus, 1758). Available online at:http://www.fao.org/fishery/culturedspecies/Gadus_morhua/en(accessed September 8, 2021). [Google Scholar]
70. Farhat S., Florent I., Noel B., Kayal E., Da Silva C., Bigeard E., et al. (2018). Comparative time-scale gene expression analysis highlights the infection processes of twoAmoebophrya strains.Front. Microbiol.9:2251. 10.3389/fmicb.2018.02251 [DOI] [PMC free article] [PubMed] [Google Scholar]
71. Figueras A., Fernández Robledo J., Novoa B. (1992). Occurrence of haplosporidian andPerkinsus-like infections in carpet-shell clams,Ruditapes decussatus (Linnaeus 1758), of the Ria de Vigo (Galicia, NW Spain).J. Shell. Res.11377–382. [Google Scholar]
72. Figueroa R. I., Garcés E., Massana R., Camp J. (2008). Description, host-specificity, and strain selectivity of the dinoflagellate parasiteParvilucifera sinerae sp. nov. (Perkinsozoa).Protist159563–578. 10.1016/j.protis.2008.05.003 [DOI] [PubMed] [Google Scholar]
73. Fisher M. C., Garner T. W. J., Walker S. F. (2009). Global emergence ofBatrachochytrium dendrobatidis and amphibian chytridiomycosis in space, time, and host.Annu. Rev. Microbiol.63291–310. 10.1146/annurev.micro.091208.073435 [DOI] [PubMed] [Google Scholar]
74. Ford S. E. (1996). Range extension by the oyster parasitePerkinsus marinus into the northeastern United States: response to climate change?Oceanogr. Lit. Rev.12:1265. [Google Scholar]
75. Ford S. E., Smolowitz R. (2007). Infection dynamics of an oyster parasite in its newly expanded range.Mar. Biol.151119–133. 10.1007/s00227-006-0454-6 [DOI] [Google Scholar]
76. Freeman M. (2009). X-cell parasites in the European dabLimanda limanda are related to other X-cell organisms: a discussion on the potential identity of this new group of parasites.Parasitology136967–980. 10.1017/S0031182009006507 [DOI] [PubMed] [Google Scholar]
77. Freeman M. A., Eydal M., Yoshimizu M., Watanabe K., Miura K., Ogawa K. (2011). Molecular identification and transmission studies of X-cell parasites from Atlantic codGadus morhua (Gadiformes: Gadidae) and the northern black flounderPseudopleuronectes obscurus (Pleuronectiformes: Pleuronectidae).Parasit. Vectors4:15. [DOI] [PMC free article] [PubMed] [Google Scholar]
78. Freeman M. A., Fuss J., Kristmundsson Á, Bjorbækmo M. F. M., Mangot J.-F., del Campo J., et al. (2017). X-cells are globally distributed, genetically divergent fish parasites related to Perkinsids and Dinoflagellates.Curr. Biol.271645–1651.e3. 10.1016/j.cub.2017.04.045 [DOI] [PubMed] [Google Scholar]
79. Galil B. S. (2000). A sea under siege – alien species in the Mediterranean.Biol. Inv.2177–186. 10.1023/A:1010057010476 [DOI] [Google Scholar]
80. Gallardi D. (2014). Effects of bivalve aquaculture on the environment and their possible mitigation: a review.Fish. Aquac. J.5:105. 10.4172/2150-3508.1000105 [DOI] [Google Scholar]
81. Garcés E., Alacid E., Reñé A., Petrou K., Simó R. (2013b). Host-released dimethylsulphide activates the dinoflagellate parasitoidParvilucifera sinerae.ISME J.71065–1068. 10.1038/ismej.2012.173 [DOI] [PMC free article] [PubMed] [Google Scholar]
82. Garcés E., Alacid E., Bravo I., Fraga S., Figueroa R. I. (2013a).Parvilucifera sinerae (Alveolata, Myzozoa) is a generalist parasitoid of dinoflagellates.Protist164245–260. 10.1016/j.protis.2012.11.004 [DOI] [PubMed] [Google Scholar]
83. Gewin V. (2008). Riders of a modern-day Ark.PLoS Biol.6:e24. 10.1371/journal.pbio.0060024 [DOI] [PMC free article] [PubMed] [Google Scholar]
84. Goedken M., Morsey B., Sunila I., Dungan C., Guise S. D. (2005). The effects of temperature and salinity on apoptosis ofCrassostrea virginica hemocytes andPerkinsus marinus.J. Shell. Res.24177–183. [Google Scholar]
85. Goggin L. (1992). Occurence of parasites of the genusPerkinsus in France.Bull. Eur. Ass. Fish. Pathol.12:174. [Google Scholar]
86. Goulletquer P., Héral M., Rotschild B. (1994). Causes of decline of oyster production (Crassostrea virginica) in the Maryland portion of the Chesapeake Bay: a literature study.Haliotis2387–112. [Google Scholar]
87. Gray M., Miller D., Hoverman J. (2009). Ecology and pathology of amphibian ranaviruses.Dis. Aquat. Org.87243–266. 10.3354/dao02138 [DOI] [PubMed] [Google Scholar]
88. Green D. E., Converse K. A., Schrader A. K. (2002). Epizootiology of sixty-four amphibian morbidity and mortality events in the USA, 1996–2001.Ann. N. Y. Acad. Sci.969323–339. 10.1111/j.1749-6632.2002.tb04400.x [DOI] [PubMed] [Google Scholar]
89. Guidi L., Chaffron S., Bittner L., Eveillard D., Larhlimi A., Roux S., et al. (2016). Plankton networks driving carbon export in the oligotrophic ocean.Nature532465–470. 10.1038/nature16942 [DOI] [PMC free article] [PubMed] [Google Scholar]
90. Guo X., Ford S. E. (2016). Infectious diseases of marine molluscs and host responses as revealed by genomic tools.Phil. Trans. R. Soc. B371:20150206. 10.1098/rstb.2015.0206 [DOI] [PMC free article] [PubMed] [Google Scholar]
91. Hamaguchi M., Suzuki N., Usuki H., Ishioka H. (1998). Perkinsus protozoan infection in short-necked clamTapes (= Ruditapes) philippinarum in Japan.Fish Pathol.33473–480. 10.3147/jsfp.33.473 [DOI] [Google Scholar]
92. Hartikainen H., Ashford O. S., Berney C., Okamura B., Feist S. W., Baker-Austin C., et al. (2014). Lineage-specific molecular probing reveals novel diversity and ecological partitioning of haplosporidians.ISME J.8177–186. 10.1038/ismej.2013.136 [DOI] [PMC free article] [PubMed] [Google Scholar]
93. Harvell C. D. (1999). Emerging marine diseases–climate links and anthropogenic factors.Science2851505–1510. 10.1126/science.285.5433.1505 [DOI] [PubMed] [Google Scholar]
94. Héral M., Rothschild B. J., Goulletquer P. (1990).Decline of Oyster Production in the Maryland Portion of the Chesapeake Bay: Causes and Perspectives.Copenhagen: International Council for the Exploration the Sea. [Google Scholar]
95. Ibelings B. W., De Bruin A., Kagami M., Rijkeboer M., Brehm G. M., Van Donk E. (2004). Review of host parasite interactions between freshwater phytoplankton and chytrid fungi (Chytridiomycota).J. Phycol.40437–453. [Google Scholar]
96. Isidoro-Ayza M., Grear D. A., Chambouvet A. (2019). Pathology and case definition of severe Perkinsea infection of frogs.Vet. Path.56133–142. 10.1177/0300985818798132 [DOI] [PubMed] [Google Scholar]
97. Isidoro-Ayza M., Lorch J. M., Grear D. A., Winzeler M., Calhoun D. L., Barichivich W. J. (2017). Pathogenic lineage of Perkinsea associated with mass mortality of frogs across the United States.Sci. Rep.7:10288. 10.1038/s41598-017-10456-1 [DOI] [PMC free article] [PubMed] [Google Scholar]
98. Jeon B. S., Nam S. W., Kim S., Park M. G. (2018). Revisiting the Parvilucifera infectans /P. sinerae (Alveolata, Perkinsozoa) species complex, two parasitoids of dinoflagellates.Algae331–19. 10.4490/algae.2018.33.3.6 [DOI] [Google Scholar]
99. Jobard M., Wawrzyniak I., Bronner G., Marie D., Vellet A., Sime-Ngando T., et al. (2020). Freshwater Perkinsea: diversity, ecology and genomic information.J. Plank. Res.423–17. 10.1093/plankt/fbz068 [DOI] [Google Scholar]
100. Jones C. G., Lawton J. H., Shachak M. (1996). “Organisms as ecosystem engineers,” inEcosystem Management: Selected Readings, edsSamson F. B., Knopf F. L. (New York, NY: Springer; ), 130–147. 10.1007/978-1-4612-4018-1_14 [DOI] [Google Scholar]
101. Jones M., Armién A., Rothermel B., Pessier A. (2012). Granulomatous myositis associated with a novel alveolate pathogen in an adult southern leopard frog (Lithobates sphenocephalus).Dis. Aquat. Org.102163–167. 10.3354/dao02539 [DOI] [PubMed] [Google Scholar]
102. Karlsbakk E., Nystøyl C. F., Plarre H., Nylund A. (2021). A novel protist parasite,Salmoxcellia vastator n. gen., n. sp. (Xcelliidae, Perkinsozoa), infecting farmed salmonids in Norway.Parasites Vectors14:431. 10.1186/s13071-021-04886-0 [DOI] [PMC free article] [PubMed] [Google Scholar]
103. Katsura K., Yamazaki F., Hamada K., Oishi K., Harada T., Shinkawa T. (1984). Geographic distribution and frequency of tumorous fishes collected from the coastal waters of Hokkaido, Japan.Nippon Suisan Gakkaishi50979–984. 10.2331/suisan.50.979 [DOI] [Google Scholar]
104. Lafferty K. D., Dobson A. P., Kuris A. M. (2006). Parasites dominate food web links.Proc. Nat. Acad. Sci. U.S.A.10311211–11216. 10.1073/pnas.0604755103 [DOI] [PMC free article] [PubMed] [Google Scholar]
105. Lassalle G., de Montaudouin X., Soudant P., Paillard C. (2007). Parasite co-infection of two sympatric bivalves, theManila clam (Ruditapes philippinarum) and the cockle (Cerastoderma edule) along a latitudinal gradient.Aquat. Living Resour.2033–42. 10.1051/alr:2007013 [DOI] [Google Scholar]
106. Le Roux F., Lorenzo G., Peyret P., Audemard C., Figueras A., Vivares C., et al. (2001). Molecular evidence for the existence of two species ofMarteilia in Europe.J. Eukaryot. Microbiol.48449–454. 10.1111/j.1550-7408.2001.tb00178.x [DOI] [PubMed] [Google Scholar]
107. Leander B. S., Hoppenrath M. (2008). Ultrastructure of a novel tube-forming, intracellular parasite of dinoflagellates:Parvilucifera prorocentri sp. nov. (Alveolata, Myzozoa).Eur. J. Protistol.4455–70. 10.1016/j.ejop.2007.08.004 [DOI] [PubMed] [Google Scholar]
108. Lee M.-K., Cho B.-Y., Lee S.-J., Kang J.-Y., Jeong H. D., Huh S. H., et al. (2001). Histopathological lesions ofManila clam,Tapes philippinarum, from Hadong and Namhae coastal areas of Korea.Aquaculture201199–209. 10.1016/S0044-8486(01)00648-2 [DOI] [Google Scholar]
109. Lefranc M., Thénot A., Lepere C., Debroas D. (2005). Genetic diversity of small eukaryotes in lakes differing by their trophic status.Appl. Environ. Microbiol.715935–5942. [DOI] [PMC free article] [PubMed] [Google Scholar]
110. Leite R. B., Afonso R., Cancela M. L. (2004).Perkinsus sp. infestation in carpet-shell clams,Ruditapes decussatus (L), along the Portuguese coast. Results from a 2-year survey.Aquaculture24039–53. 10.1016/j.aquaculture.2004.05.014 [DOI] [Google Scholar]
111. Lepelletier F., Karpov S. A., Alacid E., Le Panse S., Bigeard E., Garcés E., et al. (2014a).Dinomyces arenysensis gen. et sp. nov. (Rhizophydiales,Dinomycetaceae fam. nov.), a chytrid infecting marine dinoflagellates.Protist165230–244. 10.1016/j.protis.2014.02.004 [DOI] [PubMed] [Google Scholar]
112. Lepelletier F., Karpov S. A., Le Panse S., Bigeard E., Skovgaard A., Jeanthon C., et al. (2014b).Parvilucifera rostrata sp. nov. (Perkinsozoa), a novel parasitoid that infects planktonic dinoflagellates.Protist16531–49. 10.1016/j.protis.2013.09.005 [DOI] [PubMed] [Google Scholar]
113. Lepère C., Demura M., Kawachi M., Romac S., Probert I., Vaulot D. (2011). Whole-genome amplification (WGA) of marine photosynthetic eukaryote populations: Whole-genome amplification of photosynthetic eukaryotes.FEMS Microbiol. Ecol.76513–523. 10.1111/j.1574-6941.2011.01072.x [DOI] [PubMed] [Google Scholar]
114. Lepère C., Domaizon I., Debroas D. (2008). Unexpected importance of potential parasites in the composition of the freshwater small-eukaryote community.Appl. Environ. Microbiol.742940–2949. 10.1128/AEM.01156-07 [DOI] [PMC free article] [PubMed] [Google Scholar]
115. Lesbarrères D., Balseiro A., Brunner J., Chinchar V. G., Duffus A., Kerby J., et al. (2012).Ranavirus: past, present and future.Biol. Lett.8481–483. 10.1098/rsbl.2011.0951 [DOI] [PMC free article] [PubMed] [Google Scholar]
116. Lester R. J. G., Davis G. H. G. (1981). A newPerkinsus species (Apicomplexa, Perkinsea) from the abaloneHaliotis ruber.J. Invertebr. Pathol.37181–187. 10.1016/0022-2011(81)90073-2 [DOI] [Google Scholar]
117. Letunic I., Bork P. (2019). Interactive Tree Of Life (iTOL) v4: recent updates and new developments.Nucleic Acids Res.47W256–W259. 10.1093/nar/gkz239 [DOI] [PMC free article] [PubMed] [Google Scholar]
118. Lima-Mendez G., Faust K., Henry N., Decelle J., Colin S., Carcillo F., et al. (2015). Ocean plankton. Determinants of community structure in the global plankton interactome.Science348:1262073. 10.1126/science.1262073 [DOI] [PubMed] [Google Scholar]
119. Litchman E. (2010). Invisible invaders: non-pathogenic invasive microbes in aquatic and terrestrial ecosystems: Invasive microbes.Ecol. Lett.131560–1572. 10.1111/j.1461-0248.2010.01544.x [DOI] [PubMed] [Google Scholar]
120. Logares R., Bråte J., Bertilsson S., Clasen J. L., Shalchian-Tabrizi K., Rengefors K. (2009). Infrequent marine–freshwater transitions in the microbial world.Trends Microbiol.17414–422. 10.1016/j.tim.2009.05.010 [DOI] [PubMed] [Google Scholar]
121. López-García P., Philippe H., Gail F., Moreira D. (2003). Autochthonous eukaryotic diversity in hydrothermal sediment and experimental microcolonizers at the Mid-Atlantic Ridge.Proc. Nat. Acad. Sci. U.S.A.100697–702. 10.1073/pnas.0235779100 [DOI] [PMC free article] [PubMed] [Google Scholar]
122. López-García P., Rodriguez-Valera F., Pedrós-Alió C., Moreira D. (2001). Unexpected diversity of small eukaryotes in deep-sea Antarctic plankton.Nature409603–607. [DOI] [PubMed] [Google Scholar]
123. Lough R., Valentine P., Potter D., Auditore P., Bolz G., Neilson J., et al. (1989). Ecology and distribution of juvenile cod and haddock in relation to sediment type and bottom currents on eastern Georges Bank.Mar. Ecol. Prog. Ser.561–12. 10.3354/meps056001 [DOI] [Google Scholar]
124. Lymbery A. J. (1989). Host specificity, host range and host preference.Parasitol. Today5:298. 10.1016/0169-4758(89)90021-5 [DOI] [PubMed] [Google Scholar]
125. Mackin J. G., Owen H. M., Collier A. (1950). Preliminary note on the occurrence of a new protistan parasite,Dermocystidium marinum n. sp. inCrassostrea virginica (Gmelin).Science111328–329. 10.1126/science.111.2883.328 [DOI] [PubMed] [Google Scholar]
126. Mangot J.-F., Debroas D., Domaizon I. (2011). Perkinsozoa, a well-known marine protozoan flagellate parasite group, newly identified in lacustrine systems: a review.Hydrobiologia65937–48. 10.1007/s10750-010-0268-x [DOI] [Google Scholar]
127. Mangot J.-F., Lepère C., Bouvier C., Debroas D., Domaizon I. (2009). Community structure and dynamics of small eukaryotes targeted by new oligonucleotide probes: new insight into the lacustrine microbial food web.Appl. Environ. Microbiol.756373–6381. 10.1128/AEM.00607-09 [DOI] [PMC free article] [PubMed] [Google Scholar]
128. McCauley D. J., Pinsky M. L., Palumbi S. R., Estes J. A., Joyce F. H., Warner R. R. (2015). Marine defaunation: animal loss in the global ocean.Science347:1255641. 10.1126/science.1255641 [DOI] [PubMed] [Google Scholar]
129. Miwa S., Nakayasu C., Kamaishi T., Yoshiura Y. (2004). X-cells in fish pseudotumors are parasitic protozoans.Dis. Aquat. Org.58165–170. 10.3354/dao058165 [DOI] [PubMed] [Google Scholar]
130. Moon-van der Staay S. Y., De Wachter R., Vaulot D. (2001). Oceanic 18S rDNA sequences from picoplankton reveal unsuspected eukaryotic diversity.Nature409607–610. 10.1038/35054541 [DOI] [PubMed] [Google Scholar]
131. Morley N. J. (2010). Interactive effects of infectious diseases and pollution in aquatic molluscs.Aquat. Toxicol.9627–36. 10.1016/j.aquatox.2009.09.017 [DOI] [PubMed] [Google Scholar]
132. Morrison C. M., Shum G., Appy R. G., Odense P., Annand C. (1982). Histology and prevalence of X-cell lesions in atlantic cod (Gadus morhua).Can. J. Fish. Aquat. Sci.391519–1530. 10.1139/f82-204 [DOI] [Google Scholar]
133. Norén F., Moestrup Ø, Rehnstam-Holm A.-S. (1999).Parvilucifera infectans Norén etMoestrup gen. et sp. nov. (Perkinsozoa phylum nov.): a parasitic flagellate capable of killing toxic microalgae.Eur. J. Protistol.35233–254. 10.1016/S0932-4739(99)80001-7 [DOI] [Google Scholar]
134. Oie-Listed Diseases (2021).OIE - World Organisation for Animal Health. Available online at:https://www.oie.int/en/animal-health-in-the-world/oie-listed-diseases-2021/(accessed February 10, 20210). [Google Scholar]
135. Olson D. H., Aanensen D. M., Ronnenberg K. L., Powell C. I., Walker S. F., Bielby J., et al. (2013). Mapping the global emergence ofBatrachochytrium dendrobatidis, the amphibian chytrid fungus.PLoS One8:e56802. 10.1371/journal.pone.0056802 [DOI] [PMC free article] [PubMed] [Google Scholar]
136. Pagenkopp Lohan K. M., Hill-Spanik K. M., Torchin M. E., Fleischer R. C., Carnegie R. B., Reece K. S., et al. (2018). Phylogeography and connectivity of molluscan parasites:Perkinsus spp. in Panama and beyond.Int. J. Parasitol.48135–144. 10.1016/j.ijpara.2017.08.014 [DOI] [PubMed] [Google Scholar]
137. Park K.-I., Choi K.-S. (2001). Spatial distribution of the protozoan parasitePerkinsus sp. found in theManila clams,Ruditapes philippinarum, in Korea.Aquaculture2039–22. 10.1016/S0044-8486(01)00619-6 [DOI] [Google Scholar]
138. Park K.-I., Choi K.-S., Choi J.-W. (1999). Epizootiology ofPerkinsus sp. found in the Manila clam,Ruditapes philippinarum in Komsoe Bay, Korea.Kor. J. Fish Aquat. Sci.32303–309. [Google Scholar]
139. Patoka J., Magalhães A. L. B., Kouba A., Faulkes Z., Jerikho R., Vitule J. R. S. (2018). Invasive aquatic pets: failed policies increase risks of harmful invasions.Biodivers. Conserv.273037–3046. 10.1007/s10531-018-1581-3 [DOI] [Google Scholar]
140. Perkins F. O. (1976). Zoospores of the oyster pathogen,Dermocystidium marinum. I. Fine structure of the conoid and other sporozoan-like organelles.J. Parasitol.62959–974. 10.2307/3279192 [DOI] [Google Scholar]
141. Perkins F. O. (1996). The structure ofPerkinsus marinus (Mackin, Owen And Collier, 1950) Levine, 1978 with comments on taxonomy and phylogeny ofPerkinsus spp.J. Shellfish Res.15:67. [Google Scholar]
142. Perkins F. O., Menzel R. W. (1967). Ultrastructure of sporulation in the oyster pathogenDermocystidium marinum.J. Invertebr. Pathol.9205–229. 10.1016/0022-2011(67)90009-2 [DOI] [PubMed] [Google Scholar]
143. Poulin R., Mouillot D. (2003). Parasite specialization from a phylogenetic perspective: a new index of host specificity.Parasitology126473–480. 10.1017/S0031182003002993 [DOI] [PubMed] [Google Scholar]
144. Pretto T., Zambon M., Civettini M., Caburlotto G., Boffo L., Rossetti E., et al. (2014). Massive mortality inManila clams (Ruditapes philippinarum) farmed in the lagoon of Venice, caused byPerkinsus olseni.Bull. Eur. Ass. Fish Pathol.34:43. [Google Scholar]
145. Råberg L., Alacid E., Garces E., Figueroa R. (2014). The potential for arms race and Red Queen coevolution in a protist host-parasite system.Ecol. Evol.44775–4785. 10.1002/ece3.1314 [DOI] [PMC free article] [PubMed] [Google Scholar]
146. Ramilo A., Carrasco N., Reece K. S., Valencia J. M., Grau A., Aceituno P., et al. (2015). Update of information on perkinsosis in NW Mediterranean coast: identification ofPerkinsus spp. (Protista) in new locations and hosts.J. Invertebr. Pathol.12537–41. 10.1016/j.jip.2014.12.008 [DOI] [PubMed] [Google Scholar]
147. Reece K., Dungan C., Burreson E. (2008). Molecular epizootiology ofPerkinsus marinus andP. chesapeaki infections among wild oysters and clams in Chesapeake Bay, USA.Dis. Aquat. Org.82237–248. 10.3354/dao01997 [DOI] [PubMed] [Google Scholar]
148. Reece K. S., Siddall M. E., Burreson E. M., Graves J. E. (1997). Phylogenetic analysis ofPerkinsus based on actin gene sequences.J. Parasitol.83417–423. [PubMed] [Google Scholar]
149. Reñé A., Alacid E., Ferrera I., Garcés E. (2017a). Evolutionary trends ofPerkinsozoa (Alveolata) characters based on observations of two new genera of parasitoids of dinoflagellates,Dinovorax gen. nov. andSnorkelia gen. nov.Front. Microbiol.8:1594. 10.3389/fmicb.2017.01594 [DOI] [PMC free article] [PubMed] [Google Scholar]
150. Reñé A., Alacid E., Figueroa R. I., Rodríguez F., Garcés E. (2017b). Life-cycle, ultrastructure, and phylogeny ofParvilucifera corolla sp. nov. (Alveolata, Perkinsozoa), a parasitoid of dinoflagellates.Eur. J. Protistol.589–25. 10.1016/j.ejop.2016.11.006 [DOI] [PubMed] [Google Scholar]
151. Reñé A., Alacid E., Gallisai R., Chambouvet A., Fernández-Valero A. D., Garces E. (2021a). New Perkinsea parasitoids of dinoflagellates distantly related to theParviluciferaceae members.Front. Microbiol.12:2199. 10.3389/fmicb.2021.701196 [DOI] [PMC free article] [PubMed] [Google Scholar]
152. Reñé A., Timoneda N., Sampedro N., Alacid E., Gallisai R., Gordi J., et al. (2021b). Host preferences of coexisting Perkinsea parasitoids during coastal dinoflagellate blooms.Mol. Ecol.302417–2433. 10.1111/mec.15895 [DOI] [PubMed] [Google Scholar]
153. Rodríguez F., Figueroa R. I. (2020). Confirmation of the wide host range ofParvilucifera corolla (Alveolata, Perkinsozoa).Eur. J. Protistol.74:125690. 10.1016/j.ejop.2020.125690 [DOI] [PubMed] [Google Scholar]
154. Ruano F., Batista F. M., Arcangeli G. (2015). Perkinsosis in the clamsRuditapes decussatus andR. philippinarum in the northeastern Atlantic and Mediterranean Sea: a review.J. Invertebr. Pathol.13158–67. 10.1016/j.jip.2015.07.015 [DOI] [PubMed] [Google Scholar]
155. Ruano F., Cachola R. (1986). “Outbreak of a severe epizootic ofPerkinsus marinus (LEVIN-87) at Ria de Faro clam’s culture beds,” inProccedings of the 2nd International Colloquium on Pathology in Marine Aquaculture (Porto), (Portugal: ), 41–43. [Google Scholar]
156. Ruiz G. M., Rawlings T. K., Dobbs F. C., Drake L. A., Mullady T., Huq A., et al. (2000). Global spread of microorganisms by ships.Nature40849–50. 10.1038/35040695 [DOI] [PubMed] [Google Scholar]
157. Sakai A. K., Allendorf F. W., Holt J. S., Lodge D. M., Molofsky J., With K. A., et al. (2001). The population biology of invasive species.Annu. Rev. Ecol. Evol. Syst.32305–332. 10.1146/annurev.ecolsys.32.081501.114037 [DOI] [Google Scholar]
158. Saldarriaga J. F., McEwan M. L., Fast N. M., Taylor F. J. R., Keeling P. J. (2003). Multiple protein phylogenies show thatOxyrrhis marina andPerkinsus marinus are early branches of the dinoflagellate lineage.Int. J. Syst. Evol. Microbiol.53355–365. 10.1099/ijs.0.02328-0 [DOI] [PubMed] [Google Scholar]
159. Sanil N. K., Vijayan K. K., Kripa V., Mohamed K. S. (2010). Occurrence of the protozoan parasite,Perkinsus olseni in the wild and farmed Pearl Oyster,Pinctada fucata (Gould) from the Southeast coast of India.Aquaculture2998–14. 10.1016/j.aquaculture.2009.12.007 [DOI] [Google Scholar]
160. Scheele B. C., Pasmans F., Skerratt L. F., Berger L., Martel A., Beukema W., et al. (2019). Amphibian fungal panzootic causes catastrophic and ongoing loss of biodiversity.Science3631459–1463. 10.1126/science.aav0379 [DOI] [PubMed] [Google Scholar]
161. Seebens H., Blackburn T. M., Dyer E. E., Genovesi P., Hulme P. E., Jeschke J. M., et al. (2017). No saturation in the accumulation of alien species worldwide.Nat. Commun.8:14435. 10.1038/ncomms14435 [DOI] [PMC free article] [PubMed] [Google Scholar]
162. Sheppard B., Phillips A. (2008).Perkinsus olseni detected in Vietnamese aquacultured reef clamsTridacna crocea imported to the USA, following a mortality event.Dis. Aquat. Org.79229–235. 10.3354/dao01888 [DOI] [PubMed] [Google Scholar]
163. Siddall M. E., Reece K. S., Graves J. E., Burreson E. M. (1997). ‘Total evidence’ refutes the inclusion ofPerkinsus species in the phylumApicomplexa.Parasitology115165–176. 10.1017/S0031182097001157 [DOI] [PubMed] [Google Scholar]
164. Šimková A., Verneau O., Gelnar M., Morand S. (2006). Specificity and specialization of congeneric monogeneans parasitizing cyprinid fish.Evolution601023–1037. [PubMed] [Google Scholar]
165. Stich H. F., Acton A. B., Forrester C. R. (1976). Fish tumors and sublethal effects of pollutants.J. Fish Res. Board Can.331993–2001. [Google Scholar]
166. Stuart S. N., Chanson J. S., Cox N. A., Young B. E., Rodrigues A. S. L., Fischman D. L., et al. (2004). Status and trends of amphibian declines and extinctions worldwide.Science3061783–1786. 10.1126/science.1103538 [DOI] [PubMed] [Google Scholar]
167. Thomas F., Poulin R. (1998). Manipulation of a mollusc by a trophically transmitted parasite: convergent evolution or phylogenetic inheritance?Parasitology116431–436. 10.1017/S003118209800239X [DOI] [PubMed] [Google Scholar]
168. Valiulis G. A., Mackin J. G. (1969). Formation of sporangia and zoospores byLabyrinthomyxa sp. parasitic in the clamMacoma balthica.J. Invertebr. Pathol.14268–270. 10.1016/0022-2011(69)90115-3 [DOI] [Google Scholar]
169. Van den Wyngaert S., Gsell A. S., Spaak P., Ibelings B. W. (2013). Herbicides in the environment alter infection dynamics in a microbial host-parasite system: Infection dynamics and environmental pollution.Environ. Microbiol.15837–847. 10.1111/j.1462-2920.2012.02874.x [DOI] [PubMed] [Google Scholar]
170. Vilas R., Cao A., Pardo B. G., Fernández S., Villalba A., Martínez P. (2011). Very low microsatellite polymorphism and large heterozygote deficits suggest founder effects and cryptic structure in the parasitePerkinsus olseni.Infect. Genet. Evol.11904–911. 10.1016/j.meegid.2011.02.015 [DOI] [PubMed] [Google Scholar]
171. Vilela H. (1951).Sporozoaires Parasites de la Palourde, Tapes decussatus (L.). Lisboa: Revisita da Faculdade de Ciências, 379–386. [Google Scholar]
172. Wang Y., Yoshinaga T., Itoh N. (2018). New insights into the entrance ofPerkinsus olseni in theManila clam,Ruditapes philippinarum.J. Invertebr. Pathol.153117–121. 10.1016/j.jip.2018.03.005 [DOI] [PubMed] [Google Scholar]
173. Williamson M., Fitter A. (1996). The varying success of invaders.Ecology771661–1666. 10.2307/2265769 [DOI] [Google Scholar]
174. Winters A. D., Faisal M. (2014). Molecular and ultrastructural characterization ofHaplosporidium diporeiae n. sp., a parasite ofDiporeia sp. (Amphipoda, Gammaridea) in the Laurentian Great Lakes (USA).Parasit Vectors7:343. 10.1186/1756-3305-7-343 [DOI] [PMC free article] [PubMed] [Google Scholar]
175. YuBo L., XiChang Z., LiJun W., Bo Y., Ying Z., ChunLei C. (2001). Prevalence ofPerkinsus sp. in theManila clam Ruditapes philippinarum along northern coast of Yellow sea in China.Oceanol. Limnol. Sin.32502–511. [Google Scholar]
176. Zuendorf A., Bunge J., Behnke A., Barger K. J.-A., Stoeck T. (2006). Diversity estimates of microeukaryotes below the chemocline of the anoxic Mariager Fjord, Denmark: diversity estimates of microeukaryotes.FEMS Microbiol. Ecol.58476–491. 10.1111/j.1574-6941.2006.00171.x [DOI] [PubMed] [Google Scholar]

Supplementary Materials