Rediscovering Zygorhizidium affluens Canter: MolecularTaxonomy, Infectious Cycle, and Cryopreservation of a ChytridInfecting the Bloom-Forming Diatom Asterionella formosa

Cecilia Rad-Menéndez,a Mélanie Gerphagnon,b Andrea Garvetto,b Paola Arce,b Yacine Badis,b Télesphore Sime-Ngando,c

Claire M. M. Gachonb

aCulture Collection of Algae and Protozoa, Scottish Association for Marine Science, Scottish Marine Institute,Oban, United Kingdom

bScottish Association for Marine Science, Scottish Marine Institute, Oban, United KingdomcLaboratoire Microorganismes: Génome et Environnement, Université Clermont Auvergne, UMR CNRS 6023,Aubière, France

ABSTRACT Parasitic Chytridiomycota (chytrids) are ecologically significant in var-ious aquatic ecosystems, notably through their roles in controlling bloom-forming phytoplankton populations and in facilitating the transfer of nutrientsfrom inedible algae to higher trophic levels. The diversity and study of these ob-ligate parasites, while critical to understand the interactions between pathogensand their hosts in the environment, have been hindered by challenges inherentto their isolation and stable long-term maintenance under laboratory conditions.Here, we isolated an obligate chytrid parasite (CCAP 4086/1) on the freshwaterbloom-forming diatom Asterionella formosa and characterized its infectious cycleunder controlled conditions. Phylogenetic analyses based on 18S, 5.8S, and 28Sribosomal DNAs (rDNAs) revealed that this strain belongs to the recently de-scribed clade SW-I within the Lobulomycetales. All morphological features ob-served agree with the description of the known Asterionella parasite Zygorhiz-idium affluens Canter. We thus provide a phylogenetic placement for this chytridand present a robust and simple assay that assesses both the infection successand the viability of the host. We also validate a cryopreservation method for sta-ble and cost-effective long-term storage and demonstrate its recovery afterthawing. All the above-mentioned tools establish a new gold standard for theisolation and long-term preservation of parasitic aquatic chytrids, thus openingnew perspectives to investigate the diversity of these organisms and their physi-ology in a controlled laboratory environment.

IMPORTANCE Despite their ecological relevance, parasitic aquatic chytrids are un-derstudied, especially due to the challenges associated with their isolation andmaintenance in culture. Here we isolated and established a culture of a chytrid para-site infecting the bloom-forming freshwater diatom Asterionella formosa. The chytridmorphology suggests that it corresponds to the Asterionella parasite known as Zy-gorhizidium affluens. The phylogenetic reconstruction in the present study supportsthe hypothesis that our Z. affluens isolate belongs to the order Lobulomycetales andclusters within the novel clade SW-I. We also validate a cryopreservation method forstable and cost-effective long-term storage of parasitic chytrids of phytoplankton.The establishment of a monoclonal pathosystem in culture and its successful cryo-preservation opens the way to further investigate this ecologically relevant parasiticinteraction.

KEYWORDS biobanking, bloom dynamics, Chytridiomycota, cryopreservation,molecular methods, pathosystem, phytopathogens, phytoplankton

Received 25 July 2018 Accepted 20September 2018

Accepted manuscript posted online 28September 2018

Citation Rad-Menéndez C, Gerphagnon M,Garvetto A, Arce P, Badis Y, Sime-Ngando T,Gachon CMM. 2018. RediscoveringZygorhizidium affluens Canter: moleculartaxonomy, infectious cycle, andcryopreservation of a chytrid infecting thebloom-forming diatom Asterionella formosa.Appl Environ Microbiol 84:e01826-18. https://doi.org/10.1128/AEM.01826-18.

Editor Karyn N. Johnson, University ofQueensland

Copyright © 2018 American Society forMicrobiology. All Rights Reserved.

Address correspondence to Claire M. M.Gachon, claire.gachon@sams.ac.uk.

C.R.-M. and M.G. contributed equally to thisarticle.

ENVIRONMENTAL MICROBIOLOGY

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Fungi belonging to the phylum Chytridiomycota (i.e., chytrids) are important para-sites of freshwater phytoplankton (1). Their multifaceted contribution to food web

dynamics is increasingly recognized (2, 3): chytrid infections have been shown to beone of the main factors controlling the density and genetic structure of their hostpopulation (4–6), with a huge impact on the succession of phytoplankton species andthe entire trophic food web (7). In particular, chytrid infections of phytoplankton drivethe mycoloop, a trophic shortcut that facilitates the transfer of organic carbon and keynutrients from inedible phytoplankton to higher trophic levels (2, 8–10). Large andheavily silicified diatom cells are a good example of inedible phytoplankton exploitedby chytrids, the outbreaks of which can inhibit the development of blooms, as observedin Synedra (65.5% prevalence) and Asterionella (51.3% prevalence) from Lake Pavin andLake Aydat (France) (11, 12). Studies on the interactions between chytrids and theirhosts are rapidly moving from field-based observations to integrated “omics” and“metaomics” investigations, with the former requiring chytrid cultures and resourcepooling. For this purpose however, establishing and maintaining pure laboratorycultures of obligate parasites of phytoplankton remain a bottleneck. Renewed effortsare currently being made, leading to the successful cultivation of chytrids parasitizingdiatoms (13, 14), cyanobacteria (15), and green algae (16–18). Short life cycles, usuallycomplete within a few days, require frequent medium transfer and the supply of freshhost to ensure the viability of cultures. This skill- and labor-intensive subculturingrestricts the availability of isolates and makes them potentially subject to discontinuedmaintenance. To enable in-depth, long-term studies of this group of organisms, thereis a need to ease the maintenance burden of cultures, while also guaranteeing theirphenotypic and genotypic stability. Cryopreservation is thus an attractive option forlong-term storage, reducing both the time employed in maintaining the culture andthe risks associated with serial transfer and minimizing genetic drift, as well as thepossibility of contamination and accidental loss (19, 20). Whereas several cryopreser-vation methods have been proposed for Chytridiomycota, further protocol optimiza-tion is needed to achieve quantitative recovery of infectivity postcryopreservation (21).It is widely known that cryopreservation protocols need to be adapted to individualspecies or even strains of the same species due to variable susceptibility to cryoinjury(22–24).

In lakes, the freshwater diatom Asterionella formosa Hassall is one of the principalbloom-forming diatom species that are inedible to zooplankton (13, 25) and is knownto be susceptible to chytrid parasitism (9, 26). A. formosa is infected by three well-described chytrid species, Rhizophydium planktonicum Canter emend., Zygorhizidiumplanktonicum Canter, and Zygorhizidium affluens Canter. The morphological similaritiesamong these three species led to initial misidentification, which was later resolved byextensive morphological observation on sporangium operculation (26–28). Furtherstudies on zoospore ultrastructure confirmed the existence of the three species, whilealso suggesting that the two Zygorhizidium species should be separated at a highertaxonomical level (29, 30).

Subsequently, molecular investigation resulted in placement of R. planktonicum andZ. planktonicum into the order Chytridiales and “novel clade II” (sensu Jobard et al., 2012[31]), respectively (32). To the best of our knowledge, the phylogenetic position of Z.affluens remains to be ascertained by molecular methods, despite it being one of themajor players in the epidemic outbreaks that control its host population (28).

Molecular ecology techniques have been applied to investigate chytridiomycosisoutbreaks and, more generally, to explore fungal ecology in freshwater lakes (31, 33, 34)and in a range of other aquatic ecosystems spanning deep-sea hydrothermal vents (35),the Arctic Ocean and sea ice (36), and coastal marine habitats (37). Chytridiomycota arean important component of the fungal diversity in all aquatic ecosystems surveyed andare often the dominant fungal taxon, especially in the context of phytoplankton blooms(38). Despite their prevalence, the bulk of this environmental diversity remains unan-notated both taxonomically and functionally (e.g., referred to as “dark matter fungi”[39]). Phylogenetic reconstructions show that novel chytrid lineages are composed

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almost entirely of uncultured organisms (40). Therefore, bringing chytrids into cultureis needed not only to investigate their biology but also to establish a reference enablingthe annotation of metagenomic data. Here, we isolated and molecularly characterizedan obligate chytrid parasite on the freshwater bloom-forming diatom A. formosa anddescribed its infectious cycle under controlled conditions. Furthermore, we developeda method for the cryopreservation of the chytrid that will allow us to continue thestudies on this organism. Quantitative data on the chytrid life cycle pre- and postcryo-preservation were obtained, including infection parameters (prevalence and intensityof infection [41]) and host viability over time. To investigate the putative conservationof the relationships within the host-parasite pairing as well as the infectivity of thechytrid after cryopreservation, we also propose a double-staining method based on acombination of two fluorochromes (calcofluor white [CFW] and carboxyfluoresceinsuccinimidyl ester [CFSE]) coupled with epifluorescence microscopy.

RESULTSMorphological characterization of chytrid strain CCAP 4086/1. The progression

of the chytrid through its life cycle is described in Fig. 1. Dissemination is ensured bya spherical zoospore (2 to 3.7 �m in diameter) bearing a posterior flagellum, a largelipid globule, and a crescent-shaped nuclear cap (Fig. 1A). The spore swims toward thehost cell and encysts at its surface (Fig. 1B). The initial phase (stage I) (Fig. 1C)corresponds to the direct development of the zoospore into a young sporangium(endogenous development), characterized by the appearance of a germination tubewhich penetrates the diatom wall on the girdle region. It is followed by a maturationphase that comprises the development of the young sporangium and the growth of thegermination tube into the host cell (Fig. 1D). This is followed by the differentiation ofvisible zoospores inside the sporangium, as well as the growth of a mainly unbranched(rarely laterally unibranched) rhizoid into the host cell (stage II) (Fig. 1E). Finally, a fullymature sporangium with numerous zoospores inside is produced (stage III) (Fig. 1F).During the dehiscence phase, the mature sporangium releases zoospores via a lateral

FIG 1 Life cycle of Zygorhizidium affluens CCAP 4086/1. (A) Spherical zoospore with posterior flagellum (F), lipidglobule (LG), and nuclear cap (NC). (B) Encystment of zoospores to diatom cells. (C) Development of a zoospore intoa young sporangium (stage I) and appearance of a germination tube (GT). (D and E) Development of a youngsporangium followed by further maturation with visible zoospores inside (stage II) and rhizoid (R) growth into thehost cell. (F) Fully mature sporangium with numerous zoospores inside (stage III). (G) Empty sporangium afterrelease of zoospores (stage IV). (H) Sporangia at stages I, II, and III stained with CFW. Note the ring-shaped shade(RS) on the mature (stage III) sporangium. (I) Operculate (OP) empty sporangium (stage IV) stained with CFW. Scalebars, 5 �m.

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(rarely basal or apical) operculum. The first sign of operculation can be observed withcalcofluor white staining as a ring-shaped shade on the sporangium (Fig. 1H). Theoperculum is rarely seen attached to the empty sporangium and most often detachescompletely from it. The sporangium keeps its shape after spore release (stage IV) (Fig.1G). No sign of sexual reproduction or resting-spore formation was observed in thecultured strain CCAP 4086/1.

Molecular characterization of chytrid strain CCAP 4086/1. In the concatenatedmaximum-likelihood (ML) tree of 18S, 5.8S, and 28S ribosomal DNA (rDNA) sequences,CCAP 4086/1 is firmly placed in the order Lobulomycetales (Fig. 2); it does not groupwith any other sequenced Zygorhizidium or Rhizophydium species, which all fall in the“novel clade II” (sensu Jobard et al., 2012 [31]) and in the order Chytridiales, respectively.

Together with “uncultured Chytridiomycota Ay2007E7” from Lake Aydat in France(31), CCAP 4086/1 defines a novel clade (100% support). This clade is a sister to asecond well-supported (100%) clade that contains uncultured chytrids from the BalticSea (3c-D9, VM3-110, and 5-C10) (42). In turn, these two clades are sisters to the recentlydescribed chytrid parasite Algomyces stechlinensis within the robust group (100%bootstrap support) named SW-I (Fig. 2) (18).

Analysis of the environmental molecular diversity surrounding CCAP 4086/1 con-firms the existence of a well-defined SW-I clade (100% support) within the Lobulomy-cetales and reveals substantial diversity within it (Fig. 3, shaded clade).

Quantitative data on the life cycle of the chytrid strain CCAP 4086/1 in culture.In order to obtain a clear understanding of the chytrid development and the diatomviability upon infection, we developed a double-staining method combining the vitalcytosolic stain CFSE with the chytrid stain CFW. CFSE is a lipophilic molecule that easilypermeates the cell membrane, and it is intracellularly sequestered after hydrolysisby esterases and covalent conjugation with cytoplasmic amino groups (43). There-fore, no interaction can occur with calcofluor white, which stains extracellularN-acetylglucosamine in the cell wall (44, 45). This allowed us to follow live and deaddiatom cells in A. formosa colonies, the number of chytrids per host cell, and thedevelopmental stage of the chytrid (Fig. 4) (see details in Materials and Methods).Under optimal temperature conditions for the chytrid (15°C), the maturation of youngsporangia occurred in 24 h, as judged from the maturation of stage I and II at day 1 intomature sporangia (stage III) at day 2. At day 3, 75.6% of the chytrid population was backto stage I, demonstrating that the entire development cycle (transformation of stage Iinto stage IV and release of new infectious spores) was completed in less than 3 days(Fig. 5A).

The prevalence of infection rose steadily from 12.5% � 1% to 26.4% � 0.8% for thefirst 3 days, increasing significantly (Kruskal-Wallis test, P � 0.05) from day 4 to day 6to reach a maximum of 72.7% � 3.6% at the end of the experiment (Fig. 5B).Accordingly, the number of stage I sporangia increased significantly between day 2 andday 3 (from 37.66 � 1.5 to 107.33 � 3.05 sporangia · ml�1, respectively) and then againbetween day 4 and day 5 (from 133.3 � 6.1 to 378 � 45.4 sporangia · ml�1) (not shown).The intensity of infection stayed stable for the first 4 days (1.33 � 0.07 to 1.34 � 0.06 at day1 and day 4, respectively) before increasing significantly (P � 0.05) to reach 2.79 � 0.26at day 6 (Fig. 5B).

Quantitative assessment of chytrid infectivity postcryopreservation. Severalmethods for the cryopreservation of CCAP 4086/1 were tested (Table 1). The optimalprotocol involved dimethyl sulfoxide (DMSO) (10% [vol/vol]) as a cryoprotectant and atwo-step cooling approach. However, it was apparent that chytrid zoospores lost theirability to swim very rapidly and died within minutes during incubation in 10% DMSO.Cultures with a majority of stage I sporangia, as well as 1-week-old cultures, with a highintensity of infection and a range of life cycle stages all failed to survive the process.After several attempts to find the optimal developmental stage of the chytrid, it wasconcluded that a 3-day-old culture with high prevalence but low intensity of infectionand with a majority of mature sporangia (stage III) was most suitable to ensure

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FIG 2 Maximum-likelihood reconstruction (1,000 bootstraps) of chytrid fungal phylogeny based on three concatenated rRNA-encodinggene sequences (18S, 5.8S, and 28S). Symbols near the species name indicate the presence (*) or absence (�) of genes encoding 18S, 5.8S,and 28S rRNAs, respectively, in the alignment. Species names in bold indicate known parasites of the diatom Asterionella formosa.

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successful cryopreservation. The diatom host did not survive cryopreservation, regard-less of the conditions used.

The capability of the chytrid to infect its host after cryopreservation was comparedwith quantitative data gathered from the culture and a control containing 10% DMSOin order to check for the stability of infectivity postcryopreservation and optimize

FIG 3 Maximum-likelihood reconstruction (1,000 bootstraps) of the phylogeny of the order Lobulomycetales based on the 18S rRNA gene sequences. Alignedreference sequences of OTUs from environmental barcode surveys in aquatic ecosystems highlight the hidden diversity of close relatives of Zygorhizidiumaffluens CCAP 4086/1 (bold) within the novel clade SW-I (gray background).

FIG 4 Infected A. formosa stained by CFW and CFSE in bright-field (A) and UV (B) illumination as observed byoptical microscopy. D, dead cell; L, living cell; C, chytrid. Scale bars, 10 �m.

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propagation/recovery (Fig. 5). The use of the vital staining CFSE together with CFWallowed witnessing of the propagation of the chytrid to freshly added diatom cells thathad not undergone cryopreservation.

Initially during the first 2 days in the DMSO control, the prevalence of infection onlive cells appeared to be stable (Fig. 5D); however, as the prevalence on live cellsincreased at day 3 to reach 32.9% � 6.7%, the number of live cells started to declinefrom 1.78 · 104 to 1.05 · 104 cells/ml at day 3 and day 6, respectively (not shown). Thisoccurred as the development of the chytrid life cycle progressed through a majority ofyoung sporangia (stage I) at day 4 exactly when the intensity started to dramatically rise

FIG 5 Chytrid infection development in culture (A and B), under control DMSO conditions (C and D), and aftercryopreservation (E and F). Chytrid sporangium development (A, C, and E) and prevalence (Pr) and intensity (I) of infection(B, D, and F) were assessed over 6 days (A, B, C, and D) or 28 days (E and F). dx/y indicates x days after the yth event withintroduction of fresh A. formosa cells (see Fig. 6 for details). Panels B, D, and F show means of replicates � standarddeviation.

TABLE 1 Regeneration of viable cultures of CCAP 4086/1 using various cryopreservationprocedures and a range of developmental stages of chytrid culture

Stage (culture age,days)

Regeneration obtained with the following conditions and thawingtemp (°C):

Nalgene Controlled cooler

10%Glycerol 10% DMSO

10%Glycerol 10% DMSO

30 40 30 40 30 40 30 40

Zoospores only No No No No No No No NoI (1) No No No No No No No NoI, II, III, and IV (6) No No No No No No No NoIII (3) No No No No No No Yes Yes

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from 1.47 � 0.05 at day 4 to 2.28 � 0.91 at day 6 (Fig. 5D). After this, the youngsporangia developed into stage II (Fig. 5C) at day 5, followed by further maturation overthe course of a day into mature and then dehiscent sporangia releasing the zoosporesand generating a majority of young sporangia again at day 6 (Fig. 5C).

Initially, 6 days after the samples were thawed (Fig. 5F, d6/1 [i.e., at day 6 and afterone addition of fresh A. formosa host]), the prevalence of infection was significantlylower (14.8% � 3.13%; P � 0.05) than in the DMSO control at day 6 (54.6% � 28.1%)(Fig. 5D), showing that the infectivity of the chytrid was reduced by the cryopreserva-tion process. After addition of a second batch of fresh host (Fig. 5F), the prevalence ofinfection remained low (9.8% � 3.2%); however, it rose steadily after the addition of athird batch of diatoms. After two further additions of bait (Fig. 5F, d6/5), the prevalenceof infection in the cryopreserved culture finally reached 67.7% � 10.9%, a levelcomparable to that in both the noncryopreserved DMSO-containing control (Fig. 5D)and the reference culture (Fig. 5B) at day 6 (P � 0.05; 54.6% � 28.1% and 72.7% � 3.6%,respectively). At that point, a high proportion of the chytrid was at young or maturingdevelopmental stages (stages I and II), demonstrating the dynamism of infection. Wealso verified that the life cycle was completed in less than 3 days.

DISCUSSIONIdentification of CCAP 4086/1 as Zygorhizidium affluens and its taxonomic

implications. The establishment of a coculture from a single sporangium of the chytridCCAP 4086/1 on a monoclonal culture of A. formosa revealed that the morphologicalfeatures and the development stages of this parasite are extremely similar to thosedescribed for Rhizophydium planktonicum (26, 27): spherical zoospores with a posteriorflagellum and a single large lipid globule attach (encyst) to the host frustule, where theydevelop directly (endogenous growth) into a eucarpic monocentric sporangium, andthe rhizoidal system is long and thread-like (rarely with a single lateral branch),occupying nearly the entire infected cell. However, careful observation revealed thepresence of an operculum detaching after zoospore release. Coupled with the robustand firm nature of the sporangium, which does not collapse after spore release, thisobservation ruled out R. planktonicum as a plausible affiliation for CCAP 4086/1 (26–28).We also ruled out rarely reported chytrid parasites of A. formosa such as species 4 andspecies 5 (observed only once in Tarns, Cumbria, United Kingdom [26]), because theyare characterized by an obovoid or irregularly shaped sporangium derived from theasymmetrical swelling of the encysted zoospore, coupled with a long, laterallybranched rhizoid, in sharp disagreement with the morphology of CCAP 4086/1. Thepeculiar sporangial morphology of Rhizophydium tetragenum, i.e., originally a tetroidevolving into a Sarcina-like zoosporangium (71), made it easy to dismiss this species asa possible candidate for our organism. Of the remaining described chytrid parasitesinfecting A. formosa, only Zygorhizidium planktonicum and Zygorhizidium affluens areoperculated (26, 28). Z. planktonicum is distinct from both R. planktonicum and Z.affluens on the basis of (i) an obpyriform sporangium that is taller than it is broad, (ii)a short and heavily branched rhizoidal system, and (iii) the presence of an apical papillawhere the operculum is formed (26, 46). This operculum tends to remain hinged to theempty sporangium (26), in contrast to our observations, where the operculum had atendency to be cast off after dehiscence. Similar to the case for CCAP 4086/1, theoperculum of Z. affluens tends to detach, leaving behind only a long-lasting sphericalsporangium with a broad lateral opening. The development of the operculum startswith a circular thinning usually on one side of the sporangium, observed in transmis-sion electron microscopy (TEM) and identified as an “opercular rim” by Beakes et al. (46).A similar feature was observable in mature CFW-stained sporangia of CCAP 4086/1 (Fig.1H, ring-shaped shade), consistent with a degradation of the chitinous cell wall aroundthe opening of the forming operculum. We were also able to detect the presence of a“nuclear cap” in zoospores (Fig. 1A), which was already reported as a character specificto Z. affluens as seen by bright-field optical microscopy (28) and TEM observations (29,30). In summary, our observations match in all respects the description of Zygorhizidium

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affluens Canter (28) and distinguish the isolate CCAP 4086/1 from Z. planktonicum, R.planktonicum, and all other known species of chytrid parasites infecting A. formosa. Onthese bases, CCAP 4086/1 is here named Zygorhizidium affluens CCAP 4086/1. Recentmultigene phylogenetic analysis of the rDNAs of chytrids morphologically identified asZ. planktonicum and Zygorhizidium melosirae Canter emend. confirmed their closerelationship (14) and placed these two species in the so-called “novel clade II” (sensuJobard et al., 2012 [31]), which is otherwise composed only of environmental se-quences. In the same study, a multigene rDNA sequence of R. planktonicum wasproduced, confirming its affiliation to the Chytridiales (14, 47). Our data show that Z.affluens CCAP 4086/1 belongs to the order Lobulomycetales (48) and is thus distantlyrelated to both R. planktonicum and the Zygorhizidium species already sequenced.

The order Lobulomycetales has been described on the basis of genetic markers andzoospore ultrastructure (48) and so far contains eight characterized species (17, 31,45–48), to which metabarcoding surveys added a high richness of environmentalsequences from various habitats, including corn rhizosphere (C. Hussels, unpublisheddata), salt marshes (49), abyssal hydrothermal vents (35), alpine snow (50), and theArctic Ocean/sea ice (36). The brown and red seaweed obligate parasite Algochytropspolysiphoniae (51, 52) and the recently described parasite of volvocacean algae Algo-myces stechlinensis (18) are so far the only known parasitic members of the order. In thephylogenetic reconstruction presented here, both Z. affluens CCAP 4086/1 and Algo-myces stechlinensis belong to the well-supported novel clade SW-I (18). Within SW-I, Z.affluens CCAP 4086/1 groups together with “uncultured Chytridiomycota Ay2007E7,”which was retrieved from the eutrophic freshwater Lake Aydat (France), close to LakePavin where our strain was isolated (31) (Fig. 2). These two organisms are sisters to arobust clade composed of three environmental sequences from the Baltic Sea (42; L.Montonen, V. Kinnunen, and K. T. Steffen, unpublished data), while Algomyces stechli-nensis (the basal taxon within SW-I in our phylogenetic reconstruction) infects Eudorinaelegans in the oligotrophic Lake Stechlin (Germany). Within SW-I, Zygorhizidium affluensCCAP 4086/1 and Algomyces stechlinensis are the only two species for which theecological function has been ascertained, both being parasites of microalgae in fresh-water habitats. It is worth mentioning that environmental 28S sequences from theArctic Ocean which have been coupled with observations of chytrid parasitism ondiatoms (36) clustered as a sister taxon to the above-described group within clade SW-Iin the phylogenetic tree presented in reference 18. This suggests that algal parasitismcould be a conserved or widespread ecological strategy within SW-I. Our phylogenyagrees with the work of Beakes et al. (29), who hypothesized that Z. planktonicum andZ. affluens belonged to different genera on the basis of zoospore ultrastructure. Inparticular, Beakes et al. observed that Z. affluens zoospores lack microtubule roots, aGolgi apparatus, and a rumposome (fenestrated cisterna) (29, 30, 46). The lack of thesefeatures together with the presence of an opaque flagellar plug bearing extensions arethe principal ultrastructural characteristics used to define the order Lobulomycetales(48). On this basis, Simmons et al. (48) hypothesized the possibility of an inclusion of Z.affluens into the Lobulomycetales, in agreement with our phylogenetic conclusions.TEM analysis of Z. affluens CCAP 4086/1 zoospore ultrastructure is under way. Opera-tional taxonomic units (OTUs) retrieved using the Z. affluens CCAP 4086/1 18S rDNA asa query to screen metabarcoding data sets from different aquatic ecosystems supportthe evidence of a high diversity within SW-I, which is confirmed as a well-definedsubclade in the order Lobulomycetales. Our results do not allow us to speculate on thehabitat preferences within SW-I, since we did not investigate ecosystem others thanaquatic ones. However, our results suggest that SW-I can potentially be a high-ranktaxon within the Lobulomycetales, although this hypothesis will need further molecularand ultrastructural data to be confirmed. Overall, our data add weight to the hypothesisthat the genus Zygorhizidium is polyphyletic and therefore will need revision. For thispurpose, ultrastructural data for CCAP 4086/1 would be required, together with a betterresolution of the position of Z. affluens CCAP 4086/1 within the SW-I subclade and,ideally, the ultrastructural characterization of the closely related species Algomyces

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stechlinensis in order to identify morphological synapomorphies defining the clade.Finally, molecular investigation of the type species for Zygorhizidium (i.e., Zygorhizidiumwillei Löwenthal, which is parasitic on the green alga Cylindrocystis brebissonii) isrequired to determine whether clade SW-I or novel clade II (sensu Jobard et al., 2012[31]), if either, should retain the name Zygorhizidium.

Cryopreservation. The cryopreservation method proposed here uses the standardcryoprotectant (DMSO) and cooling rate already proposed for other fungi (53). How-ever, previous studies have highlighted the need of incorporating the physiologicalstate of the organism and the analysis of infectivity posttreatment (21). In the presentstudy, the biological condition of the organism precryopreservation was assessed andthe comparison on infectivity pre- and posttreatment investigated to ensure thestability of the culture, allowing the long-term study of the organism.

Samples with a majority of just-encysted sporangia (stage I) did not survive thecryopreservation process, which is probably related to the fact that they are trophicallydependent on their host cell (54), which also did not survive cryopreservation. Likewise,we assume that 1-week-old cultures with a high intensity of sporangia in all develop-ment stages did not survive because the host was unable to support the furtherdevelopment of the chytrid after thawing.

As previously described (23), motile zoospores did not survive cryopreservation;therefore, we hypothesized and experimentally verified that the best results would beobtained with stage III chytrids that no longer trophically rely on the host, i.e., maturesporangia full of zoospores. Once thawed, the zoospores are released and continue theinfection upon addition of fresh host cells. This addition of fresh host is imperativebecause A. formosa did not survive the cryopreservation process. Repeated inoculationof new host after each life cycle (�3 days) allowed the propagation of the chytrid.

The decrease in the prevalence and intensity of infection observed immediatelyafter cryopreservation was likely due to increased stress levels resulting from tissuedamage due to ice formation and other cryopreservation-related drawbacks (55–57).However, after 6 life cycles (�18 days) and 5 inoculums of the host, both the prevalenceand intensity of infection steadily rose to become comparable to those pretreatment(Fig. 5B to D).

Although cryopreservation protocols must be tailored to the species level withspecific cryoprotectants and cooling rates (21, 22, 24, 53), the tools described here arethe basis for the appropriate study of infectivity pre- and postcryopreservation. Thus,they illustrate the reliability of the method. We hope that the novel protocols estab-lished here will ease the maintenance burden for obligate chytrid parasites andtherefore stimulate efforts on the isolation of novel strains, the investigation of theirphysiology and phenotypic plasticity, and the generalization of our results. We alsohope that these protocols may inspire future research on other parasites, for example,obligate biotrophic plant pathogens.

MATERIALS AND METHODSSample collection. Samples were collected in Lake Pavin (45°29=41�N, 2°53=12�E), an oligotrophic

deep volcanic mountain lake (maximum depth [Zmax] � 92 m) characterized by small surface (44 ha) andsmall drainage basin (50 ha) areas. A weekly sampling mission was undertaken from March to April 2013during the annual diatom bloom, near the center of the lake at the point of maximum depth. Twentyliters was sampled using an 8-liter Van Dorn bottle at the middle of the euphotic layer (estimated fromSecchi depth). To eliminate the metazoan zooplankton, collected samples were immediately prefilteredthrough a 150-�m-pore-size nylon filter. The filtrate was then concentrated on a 25-�m-pore-size nylonfilter, collected by washing the filter with 0.2-�m-pore-size-filtered lake water, poured into sterilizedtransparent recipient flasks, and then transferred immediately to the laboratory for processing.

Strain isolation, purification, and culture conditions. A. formosa was isolated by micropipettingusing a 20-�l glass capillary (Brauband; intraMARK, Germany). Single colonies of A. formosa were pickedand transferred into 6-well plates containing fresh sterile diatom medium (DM) (29). The diatom wasmaintained at 20°C under a 12/12-h light/dark regime (irradiance, �64 �mol · m�2 · s�1). Likewise,colonies of A. formosa infected with one sporangium of the chytrid were isolated following the samemethod and incubated at 15°C using the same light conditions as described above.

A number of strategies were combined to purify these clonal cultures. Initially, serial dilution wasused, by micropipetting single colonies and inoculating them into fresh sterile medium; A. formosainfected with chytrids was then filtered on 50-�m and 20-�m filter units (Celltrics; Partec, Germany); the

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diatom colonies were retained by the filter and then washed into sterile medium. This was repeated untilbacteria were the only other organisms present in the culture. Once established in culture, the chytrid-A.formosa pairing and the uninfected A. formosa strain were maintained by serial transfer every 6 days.Specifically, a 180-ml 1-week-old A. formosa culture was infected with 20 ml of a 1-week-old chytridculture.

All strains used in this study are freely available from the Culture Collection of Algae and Protozoa(CCAP) under the following accession numbers: A. formosa, CCAP 1005/23; and chytrid, CCAP 4086/1.

Strain characterization and culture synchronization. In order to characterize the chytrid strain andquantify the infection process under optimal conditions (as described above), a synchronized chytridculture was studied over 6 days. To obtain a synchronized chytrid culture, a 1-week-old chytrid culturewas filtered successively through 25-�m, 10-�m, and 5-�m nylon meshes to obtain a suspension offungal zoospores, free of host cells. As previously observed, the filtration process does not affect thezoospore swimming activity (13). After 1 week, the chytrid culture was subcultured as described above(10% [vol/vol]) into 1-week-old A. formosa previously grown at 20°C under a 12/12-h light/dark regime(irradiance, �64 �mol · m�2 · s�1). Daily, the host density, the chytrid life cycle, and the prevalence(percentage of infected cells in a host population) and intensity (number of sporangia per infected cell)of infection, two classical parameters used to study this group of organisms (41), were studied. Diatomconcentrations were determined with a Sedgwick-Rafter counting chamber (Hausser Scientific, Horsham,PA, USA). To determine the prevalence and intensity, 1 ml of the chytrid culture was stained with thefluorochrome calcofluor white (CFW) (final concentration, 2.5% [vol/vol]) and examined using UVexcitation (405 nm) under an inverted Zeiss Axioskop 2 epifluorescence microscope (Carl Zeiss, Ger-many). Systematically, 100 colonies, representing at least 400 cells, were examined, and both theprevalence and intensity of infection, as well as the chytrid life stage and morphological characteristics,were recorded.

Retrieving CCAP 4086/1 rDNAs. A transcriptomic database generated from the pathosysteminvolving the chytrid CCAP 4086/1 and A. formosa CCAP 1005/23 was queried for the presence of theparasite and its host. Briefly, reads for each sample were quality checked with FastQC (58), trimmed usingTrimmomatic (59), and quality checked a second time (FastQC), and then each sample was lane-wiseassembled via Trinity (60). Using BLAST and NCBI E-utilities, de novo-assembled contigs from a heavilyinfected sample of A. formosa and CCAP 4086/1 were queried to obtain rDNA belonging to the hostdiatom and the chytrid parasite using GenBank 18S rDNA sequences of A. formosa (HQ912633) and“uncultured Chytridiomycota Ay2007E7” (JQ689413). Contigs whose similarity to the query sequenceswas above 95% were subjected to further BLAST analysis against GenBank, and via this procedure contigTrinity_DN14199_c0_g7_i3 (4,962 bp containing 18S rDNA, internal transcribed spacer 1 [ITS1], 5.8SrDNA, ITS2, and 28S rDNA) was identified as belonging to the chytrid parasite (with 99.48% identity toJQ689413 over 1,162 bp) and subsequently chosen as a genetic marker for further phylogenetic analysis.

Phylogenetic analysis. We assembled a data set of the 18S, 5.8S, and 28S rDNA sequences ofchytrids based on the work of Seto et al. (14) in order to encompass all the known molecular diversityof chytrid parasites of A. formosa. Since preliminary findings pointed toward inclusion of CCAP 4086/1 inthe recently established order Lobulomycetales (48), long and informative rDNA sequences within thisorder were included in the tree (18, 32, 61). Finally, “uncultured Chytridiomycota Ay2007E7” was includedas the best GenBank match against CCAP 4086/1, and three 18S rDNA sequences from environmentalsurveys (uncultured fungi 3c-D9, 5-C10, and VM3-110) matching the query sequence with identities of�95% were also added to our data set (42). Sequences were aligned in Geneious 6.1.8 (62) using MAFFT(63), manually checked, and trimmed for the presence of introns. With IQ-TREE 1.5.5 (64), substitutionmodels best fitting the data were assessed for each gene separately via ModelFinder (65), resulting inTIM2R5 (18S and 28S) and TPM2IG4 (5.8S). A concatenated alignment was analyzed with the samesoftware using a partitioned model (66) under the �spp option, i.e., allowing each gene to evolve at itsown speed, and a maximum-likelihood tree was inferred using a bootstrap test of phylogeny with 1,000replicates.

Diversity assessment in environmental barcode data sets. An in-house script combining EDirectand SRA Toolkit utilities, referred to as MOULINETTE (67), was used to screen �19,000 metabarcode datasets from projects deposited in the NCBI Sequence Read Archive (SRA). Those were selected using thekeywords “freshwater,” “lake,” “wastewater,” and “aquatic” (see reference 67 for details). Briefly reads areretained when they are at least 97% identical over 80% of the length of the query sequence (in this caseCCAP 4086/1), extracted, paired, and filtered (expected error, over 1.0). Paired reads that survived thefiltering process were then clustered into OTUs using usearch (v9.1.13) (68). All OTUs were then alignedto reference Lobulomycetales sequences, including the parasite CCAP 4086/1, using MAFFT (63). An MLtree (1,000 bootstraps) was inferred using IQ-TREE 1.5.5 (64) and ModelFinder (65) to assess thebest-fitting model of molecular evolution (i.e., HKYR3).

Cryopreservation. A range of cryoprotectants and cooling and thawing procedures were tested oncultures at different life stages (Table 1). After this initial screen, the following optimal protocol was usedfor all subsequent experiments. Cultures were cryopreserved in triplicate, using dimethyl sulfoxide(DMSO) (10% [vol/vol]) as a cryoprotectant and following a two-step cooling approach involving initialcontrolled-rate cooling followed by plunging into liquid nitrogen. Three-day-old infected cultures wereemployed since they had both a high prevalence and a low intensity of infection. Aliquots of thechytrid-A. formosa pairing (0.5 ml) were dispensed into cryovials (Greiner Bio-One GmbH, Germany).DMSO (Sigma-Aldrich Ltd., UK) was filter sterilized in sterile DM to a final concentration of 20% (vol/vol)using a 0.20-�m sterile syringe filter (Anachem, UK). An aliquot (0.5 ml) of the 20% (vol/vol) DMSOsolution was added to the harvested cells to give a final DMSO concentration of 10% (vol/vol). This was

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then incubated at room temperature (�20°C) for 20 min prior to cryopreservation. The cryovials werethen transferred to a controlled-rate cooler (Kryo 360 3.3; Planer plc, UK) and cooled at 1°C · min�1

between 20°C and �40°C. After being held for a further 15 min at �40°C, the cryovials were rapidlyremoved, plunged into liquid N2, and then transferred to a cryostorage container filled with liquid N2.

FIG 6 Schematic representation of the cryopreservation procedure (left, gray flasks) and control conditions (right, black flasks). An inoculum of a three-day-oldinfected culture (A) was mixed with the cryoprotectant DMSO (B) in a cryovial. The chytrid culture was then cryopreserved using a two-step cooling approachinvolving initial controlled-rate cooling followed by plunging into liquid nitrogen. Immediately after thawing, the samples were inoculated into cell culture flaskscontaining 3-fold more fresh A. formosa CCAP 1005/23 (C) to allow the development of chytrid CCAP 4086/1. Finally, DM (D) was added to dilute the DMSO10-fold, avoiding possible toxicity. Viability was assessed by cell counts and subsequent determination of prevalence and intensity of infection.

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To investigate the potential effect of the DMSO on the infectivity pattern of the chytrid, the sameprocedure (except for the cryopreservation/cooling of the samples) was followed to establish 3 controlreplicates. After incubation with DMSO, the control samples were inoculated into 9 ml sterile DM todilute the DMSO 10-fold, and an inoculum of the diatom host (3-fold more than the density in the initialsample) was added. The control replicates were then incubated under the same light and temperatureconditions used for the cryopreserved samples (see “Thawing and recovery” below) and were sampleddaily for 6 days by removing 1 ml and inspecting at least 100 A. formosa colonies (Fig. 6). For each colonyencountered, the number of A. formosa cells, their viability, the number of chytrids in each cell, and thechytrid life cycle stage were recorded.

Thawing and recovery. After 1 week, the samples were transferred in liquid N2 from the cryostoragefacility to the lab. They were then thawed by direct immersion in a preheated water bath at 30°C andwere removed as soon as all visible ice had melted. Immediately after thawing, the samples wereaseptically inoculated into 50-ml cell culture flasks containing fresh A. formosa CCAP 1005/23 to allow thedevelopment of chytrids, in the proportion of 1 thawed cell to 3 fresh A. formosa cells. The flask wastopped up with 9 ml sterile DM to dilute the DMSO 10-fold. The flasks were then transferred at 15°Cunder reduced light intensity (irradiance of ca. 12 �mol photons · m�2 · s�1) and a 12/12-h light/darkregime for the first 24 h to reduce potential light-induced stress (19). The samples were then incubatedat an irradiance of ca. 64 �mol photons · m�2 · s�1 for another 6 days to generate sufficient material toundertake postpreservation functional stability assessment.

Postcryopreservation viability assay. To assess postcryopreservation viability, we developed adouble-staining method with carboxyfluorescein diacetate succinimidyl ester (CFSE) used as a vital stain(69) together with the fluorochrome calcofluor white (CFW), allowing us to simultaneously assess theviability of the diatom host and stain the chytrid cell wall (Fig. 4). CFSE in the form of a 10 mM stocksolution in DMSO was freshly prepared and added to the samples to give a final concentration of 2 �M,and then the cells were incubated at room temperature for 15 min (70). Following this, CFW was addedto the samples at a 2.5% (vol/vol) final concentration following a protocol described previously (4) andincubated for a further 10 min before examination under a Zeiss Axioskop 2 epifluorescence microscope(Carl Zeiss, Germany) fitted with 100-W UV illumination and two filter sets, i.e., a type 09 filter (excitation,450 to 490 nm; dichroic mirror, 510 nm; emission, long path, 515 nm) and a type 02 filter (excitation, 365nm; dichroic mirror, 395 nm; emission, long path, 420 nm). Chitin walls stained with CFW were examinedusing UV excitation (405 nm), and the viability of diatoms stained with CFSE was explored under bluelight illumination (488 nm) using UV light excitation. Micrographs were taken with an AxioCam HRccamera (Carl Zeiss, Germany) using the AxioVision software, version 4.7.1 (Carl Zeiss, Germany).

The viability of thawed samples was estimated by systematically inspecting at least 100 A.formosa colonies. For each colony encountered, the number of A. formosa cells, their viability, thenumber of chytrids on each cell, and the chytrid life cycle stage were recorded to elucidate theprevalence and intensity of infection (41). These parameters were recorded 6 days (�2 life cycles)after thawing to allow the culture to recover from the stress induced by the cryopreservationmethod. After this, a new inoculum of the diatom host (3-fold cell/cell) was added to allow thedevelopment of the infection, and the same counts were repeated after periods of 3 days (�1 lifecycle) for 3 times, adding a new inoculum of the host (3-fold) each time. Following this, the sampleswere left for 3 days under normal conditions and a new inoculum of the host (3-fold) was added; thesamples were then left to develop a normal infection, and the counts were repeated to be able tocompare the infection levels after 2 life cycles (�6 days) postcryopreservation with the control. Theentire procedure is summarized in Fig. 6.

Statistical analysis. Due to the nonnormal data, differences in the prevalence and intensity ofinfection among time points and experimental conditions were tested with the nonparametric Kruskal-Wallis test, followed by a Mann-Whitney pairwise comparison with Bonferroni correction. All statisticalanalyses were conducted using PAST 3.08 (http://palaeo-electronica.org/2001_1/past/issue1_01.htm).

Accession number(s). The rDNA sequence of CCAP 4086/1 is available in GenBank under accessionnumber MH626496.

ACKNOWLEDGMENTSThis project has received funding from the French ANR under grant agreement

ANR-12-BSV7-0019, from the European Union’s Horizon 2020 research and innova-tion program under Marie Skłodowska-Curie grant agreement no. 642575, and fromthe UK NERC under grants MultiMARCAPP (NE/L013029/1) and GlobalSeaweed(NE/L013223/1).

We thank Matilda Haraldsson and Alain Franc for insightful discussions.C.R.-M., M.G., T.S.-N., and C.M.M.G. designed the study; C.R.-M., M.G., A.G., P.A., and

Y.B. performed experimental work; and C.R.-M., M.G., A.G., P.A., and C.M.M.G. wrote thepaper.

We have no conflict of interest.

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