Labyrinthula can be identified in culture through observation of distinctive fusiform cells embedded within hyaline ectoplasmic networks, or ‘slimeways’ (Porter, 1969). Individual Labyrinthula cells are colorless, though dense aggregations may have some yellowish/orangish coloring (Figure 2), measuring between 5 µm – 30 µm long by 3-8 µm wide (Pokorny, 1967; Muehlstein et al., 1988; Leano and Damare, 2012). Each Labyrinthula cell contains a single large central nucleus, flanked by 2 large vacuoles where stores of lipid bodies are easily visible (Jepps, 1931; Young, 1943; Porter, 1969). Labyrinthula cells also contain specialized posteriorly oriented organelles, including dictyosomes and an endoplasmic reticulum (Porter, 1969).

Early in isolation, Labyrinthula cells appear enigmatic, as actin-myosin reactions in the ectoplasm generate distinctive gliding motion of cells within the colony. Active cells may travel up to 50 µm per minute (Dietz and Schnetter, 1999; Preston and King, 2005). The plasmodium generating organelle allows the cells to develop sometimes extensive networks, which are used by the cells for both motility and osmotic feeding (Porter, 1969) by secreting exoenzymes for digestion during nutrient uptake (Tsui et al., 2009). The unique ectoplasmodic networks generated by Labyrinthula grow in root-like, or ‘lacey’ patterns (Vergeer and den Hartog, 1994). They are most often observed following a period of intensive axenic culturing and growth in a laboratory setting. Cell colonies are readily identifiable through microscopy where they can be observed forming distinctive and robust fractal-like vegetative cell communities in culture (Figure 3) (Jepps, 1931; Young, 1943; Trevathan-Tackett et al., 2018a). In the literature, Labyrinthula colonies have been variably described by morphological qualities, however there is agreement in their tendency to branch out from single-point infections. Growth characteristics of the colony may be characterized by cellular alignments, density, agar penetration and cellular sizes (Sullivan et al., 2013; Sullivan et al., 2017; Trevathan-Tackett et al., 2018a). Finally, care should be taken to avoid confusing Labyrinthula with a closely-related and commonly co-occurring taxa, such as aplanochytrids, especially for smaller-bodied Labyrinthula spp. (Figure 2, Leander et al., 2004). With experience and under similar culture conditions, aplanochytrids can appear to have a more distinct nucleus, be more rounded, and have a more ‘dotted line’ like presentation when observing continuity among cells in the leading edge/arm of a colony on agar (Leander et al., 2004).

Labyrinthula cells reproduce both sexually and asexually (Figure 4). Vegetative cells multiply through binary fission or mitotic division of vegetative cells. This is considered to be the most commonly used strategy for replication and propagation of Labyrinthula cells in naturally occurring colonies (Pokorny, 1967). A cyst-like resting phase, possibly asexual, is also reported (Pokorny, 1967; Amon, 1978). Sexual modes of reproduction have been described through accounts of aggregating plasmodia (Pokorny, 1967), sporulating cells, and zoospore recovery (Amon and Perkins, 1968; Perkins and Amon, 1969). Several accounts of cells restructuring in culture to form dense aggregations of cells with nucleated walls are described as ‘sori’ (Pokorny, 1967). The ‘spores’ aggregating inside the ‘sori’ are extremely difficult to visualize and often usual staining methods are not effective (Jepps, 1931). However, bi-flagellated zoospores were isolated from a sorus and photographed using a scanning electron microscope (Amon and Perkins, 1968). Imaging and assessment of chemical signaling and quorum-sensing behaviors has been described for other organisms, such as bacteria (De Kievit and Iglewski, 2000), but is generally lacking for stramenopiles (Hassani et al., 2018). The role of chemotaxis in Labyrinthula life cycle and behavior are unknown.

The first attempt to morphologically and taxonomically describe separate Labyrinthula (Cienkowski) species evaluated and summarized cell structure, colony morphology, physiology, ecology and pathogenic considerations, for 10 species (L. macrocystis, L. minuta, L. vitellina var. vitellina, L. vitellina var. pacifica, L. cienkowski, L. zopfii, L. valkonovii, L. chattonii, L. algeriensis, L. roscoffensis, L. coenosystis) of known isolates (Pokorny, 1967). A review of taxonomy reveals early nomenclature was abandoned when two epithets emerged in the literature as pathogenic phenomenon were discovered in Labyrinthula, including L. zosterae (Porter, 1990) affecting seagrasses and L. terrestris (Bigelow et al., 2005) affecting turf grasses. A third species, Labyrinthula diatomea, is more recently described and is associated with marine sediments where it primarily, if not entirely, consumes diatoms (Popova et al., 2020). There may be redundancy in the descriptions of cultured specimens based on morphological features, however, molecular sequence-based approaches help clarify, replicate and underpin resolution efforts in this arena (e.g. Martin et al., 2016).

Metabarcoding studies commonly target the V9 section of the 18S rDNA gene to describe protist communities (Amaral-Zettler et al., 2009). Huge data collection surveys like the Tara Ocean Expedition or the Earth Microbiome Project accumulate an immense database of 18S rDNA sequences and associated environmental data. These efforts drive advances in protistan phylogeny and accelerate our understanding of ecological needs for otherwise cryptic taxa, including Labyrinthulomycetes (Pan et al., 2017). Global Labyrinthula surveys, including two nuclear DNA sequencing methods (18S and ITS amplicons) report there are at least 16-21 Labyrinthula species covering both terrestrial and marine habitats (Bigelow et al., 2005; Martin et al., 2016). ITS sequences are more commonly used as a barcode for fungi phylogeny (Schoch et al., 2012). Therefore, both currently used molecular markers have high quality reference data and are a good tool for discriminating taxonomic units and to describe diversity within the genus Labyrinthula. Newer whole genome approaches will eventually resolve phylogenetic uncertainties.

Given the plastic and somewhat featureless nature of Labyrinthula cells at the level of gross morphology, genetic signatures are a critical pathway for resolution. In the last decade, molecular techniques have been more commonly used to identify Labyrinthula spp. cultured from field samples (see Section 3 for details on molecular markers). Progress has been made in classifying Labyrinthula in the context of other genera and families in this order (Pan et al., 2016; Pan et al., 2017). The most recent key developed to describe distinguishable morphological characteristics of species in this genus was prepared over 20 years ago (Dick, 2001). Genetic evidence for Labyrinthulomycetes species diversity has recently leaned more on barcoding-type approaches, but also ecological context, to establish speciation and develop a stable taxonomic key (Martin et al., 2016). Given the robust record of publishing species descriptions, recent advances in genome sequencing (Tan et al., 2021) and success in developing prototypic genetic-level species delineations (Martin et al., 2016), there are new opportunities to resolve historic and ongoing discoveries into speciation and evolution of Labyrinthula.

Prior to sampling collected plant material for transfer onto agar media, it is advisable to pre-treat tissues to reduce contaminants (See Supplemental Manual). Labyrinthulids are able to utilize a range of microbial or plant substrates (See Supplemental Manual) for survival (Sykes and Porter, 1973). This is consistent with findings about their diverse biogeographic range, hosts, and trophism. Variabilities in nutrients and other additives in media may have a substantial impact on the cell and colony characteristics of isolates. Accordingly, care should be taken with the intent of using any culture medium, as the consequences of different media on growth results remains an open question, especially with regards to affecting strain virulence. A summary of standard agar and liquid culture methods successfully used to culture Labyrinthula can be found in the Supplementary Manual.

Since Labyrinthula does not grow fully homogeneous in liquid culture or on agar plates, it is difficult to control Labyrinthula cell densities at the start of any assay. For this reason, it is important to advance protocol to standardize precise conditions and controls to account for the overall variance in Labyrinthula growth between replicates. The length of the assay will depend on how fast isolates are growing. All assays should be stopped before the colony reaches the edge of the tank, petri dish or well-plate. Those types exhibiting more agar-surface growth tend to grow faster and often reach the edge within days, while predominantly in-agar types are usually slower and may never reach the edge. Given the phenotypic plasticity observed in Labyrinthula, care should be taken to make sure the Labyrinthula isolate used for all replicates of the assay exhibit similar cell size and shape, and that they are growing at roughly the same rate. Thus, colonies used in assays should be the same age, and maintained by transferring to new agar plates at the same time (ideally, not more than 1 week or so old). Finally, as noted by Martin et al. (2009), abiotic conditions (and likely biotic, as well) can alter both colony extent and cell densities within colonies/ectoplasmic networks, with experimental designs and questions dictating whether both measures are necessary. Three key parameters (dish size, borer size and incubation time) can be altered to fit the need of the assay. For example, if the assay is used to investigate Labyrinthula growth on seagrass extracts it can be beneficial to use smaller petri dishes or well-plates to minimize the amount of extracts used for each replicate (Trevathan-Tackett et al., 2015; Jakobsson-Thor et al., 2018). If long term data are required, a larger dish and longer incubation time may be needed.

Staining of plant tissues has been effective in revealing important cellular characteristics of Labyrinthula, including descriptions of key structures and functions (Jepps, 1931; Young, 1938; Watson and Raper, 1957; Armstrong et al., 2000; Preston and King, 2005). Early visualizations were drawn by Jepps (1931) observed from stained and fixed Labyrinthula cells, as viewed through a microscope. More recently, color images of histological investigations of seagrass demonstrate the presence of Labyrinthula cells within the leaf tissue of its seagrass host (Groner et al., 2014; Groner et al., 2016). For example, in our experience and that of Harrington and Hageage (2003) when utilizing a fluorescing stain, it is strongly advised to carefully evaluate stain performance under particular conditions, as it may vary by organism, pH, production batch, and many other factors. Notably for seagrasses, background fluorescence can vary significantly by species (e.g., Figure 5A, B), leaf age (possibly related to cuticle thickness and/or composition), degree of fouling and damage to the epidermis (Figures 5A–C), and microscope filters used.

An inverted microscope with phase contrast allows for the best visualization of cell and colony formations and measurements needed to verify the presence of Labyrinthula organisms. Scanning electron microscopy (SEM) has been successfully used to view zoospores (Amon and Perkins, 1968; Perkins and Amon, 1969) and ultrastructure (Perkins, 1972) in Labyrinthulomycota. To improve microscopic images of cultured colonies, staining and light manipulations may be required. Exploration and discovery of these and other visualization tools may aid in further discoveries of temporal and spatial qualities of host-pathogen relationships and trophic transfers.

Research on Labyrinthula disease and ecology have primarily focused on laboratory pathogenicity assays. Investigations into seagrass pathogenicity testing of Labyrinthula spp. was pioneered by Charles Renn (Renn, 1936, Watson and Ordal 1957, Muehlstein, 1992; Martin et al., 2009; Sullivan et al., 2013) using Labyrinthula to test Koch’s postulate. Direct observation of infection and pathogenicity are most common (Figure 6, See Supplemental Manual). Laboratory and field assays are both used to investigate a) pathogenicity and virulence of newly isolated cultures (Vergeer and den Hartog, 1994; Martin et al., 2016; Trevathan-Tackett et al., 2018a), b) how infection develops naturally on infected plants under various environmental conditions (Buchsbaum et al., 1990; Vergeer et al., 1995; Beets et al., 2014; Dawkins et al., 2018), and c) variability in infectiousness of healthy or asymptomatic plants by inoculating under different environmental conditions (McKone and Tanner, 2009; Trevathan et al., 2011). Pathogenicity in seagrass has also been researched indirectly through correlation of environmental parameters, and spatial distribution and coverage (Cottam and Munro, 1954; Vergeer and den Hartog, 1994; Lathrop et al., 2001).

Different kinds of L. zosterae vectors, including inoculated leaf tissues, agar plugs (Jakobsson-Thor et al., 2019), gauze and water-baths (Groner et al., 2016) have been used to perform infection experiments on healthy seagrass shoots. It is important to standardize experimental vector infection protocol for pathogenicity assays. While several techniques have been used to infect plants with Labyrinthula (Martin et al., 2016), the majority of pathogenicity assay techniques use surface-to-surface transfer. A limitation with the method above is that it is difficult to control for L. zosterae quantities added during inoculation. It is important to be able to control pathogen loads when studying disease severity and other density-dependent effects. Only the in vitro water-borne method by Groner et al. (2016) makes it possible to evaluate and control the concentration of the Labyrinthula cells when performing infection experiments prior to treatment. In this method, a hemacytometer is used to determine the Labyrinthula cell dosage in either a concentrated inoculum applied directly to the plant or as an inoculum bath in which the ramet is immersed. Similar assays have been performed on fresh leaf sections thereby providing a relative measure of potential virulence or severity (Groner et al., 2014; Dawkins et al., 2018). This method can be used for transferring cells as mentioned above, but concentrations can also be sampled and assessed with this method at any time to understand pathogen loading and variability due to concentration.

Infection experiments have so far mainly been used to investigate direct infection effects on a given seagrass host (Short et al., 1993; Steele et al., 2005; McKone and Tanner, 2009; Garcias-Bonet et al., 2011; Trevathan et al., 2011). Recent studies on the effect of physical and chemical environmental stressors, e.g. temperature, salinity, light, pH and interactions of these factors, have shed some light on how a changing environment will affect L. zosterae infection (Bishop et al., 2017; Dawkins et al., 2018; Brakel et al., 2019; Jakobsson-Thor et al., 2020).

Compounds produced by Labyrinthula spp. and their hosts may provide valuable information on their interactions. Measurement of particular compounds or classes thereof (e.g. phenolics) may inform targeted questions, while overall metabolic signatures (usually of a certain class of compounds) of a given sample may provide broader or more exploratory information. Metabolomics has been useful in the examination of phylogeny and halophyte defense compounds. For instance, Huang et al. (2003) used fatty acid (FA) signatures to cluster unknown isolates of thraustochytrids against a phylogeny constructed with the 18S rRNA gene. For such methods to be applicable for Labyrinthula phylogeny, a precise understanding of how FA are synthesized and how production may change through disease progression is required. Also, though the presence of defense compounds can be detected if there is a change in concentration between different treatments, such as between an infected and uninfected plant (Brakel et al., 2014), it is difficult to detect these defense compounds if the compounds are at the same levels and always active (constitutive defense).

It has been hypothesized that seagrasses defend against pathogenic Labyrinthula species by the production of chemical defense compounds. Overall, secondary metabolites, including chemical defenses against Labyrinthula are poorly understood (Zidorn, 2016). Despite research, no direct links between defense compounds and resilience have been identified thus far.

Several studies have investigated the effect of phenolic acids on Labyrinthula, a well-known group of defense compounds in terrestrial plants. While some have found a positive correlation between Labyrinthula infection and phenolic acid production in seagrasses (Vergeer and Develi, 1997; McKone and Tanner, 2009), others suggest phenolics accumulation in diseased leaves is ‘pseudo-induced’ due to lesions disrupting the allocation and storage of leaf resources (Steele et al., 2005). These methods have made it possible to compare phenolic acids in seagrass shoots collected from the field, or from shoots used in different kinds of infection experiments (Groner et al., 2016; Groner et al., 2018). Still, the role of phenolics in moderating Labyrinthula is not certain.

Additional compounds with inhibitory effects on Labyrinthula growth have been isolated from T. testudinum, but the structures have not been confirmed (Trevathan-Tackett et al., 2015). The isolation was done using bioassay-guided fractionation, a common method for isolation of bioactive compounds. This method allows detection and isolation of inhibitory compounds from complex samples without targeting a specific group of compounds. However, the extraction technique used by Trevathan-Tackett et al. (2015) did not target phenolic acids. It would be beneficial for future studies to apply a broader extraction technique that also includes phenolic acids. This would better resolve whether phenolic acids are actively inhibiting Labyrinthula growth in seagrasses or if other compounds are responsible for this interaction. Immune-related enzymes represent another group of compounds that may act on and in response to Labyrinthula, at least for turtlegrass, which is thought to impact virulence. Duffin et al. (2020) developed assays for four enzymes (peroxidase, exochitinase, polyphenol oxidase, and lysozyme activity), two of which correlate with Labyrinthula load. Further, all are expressed constitutively in turtlegrass, and appear useful in terms of elucidating Labyrinthula defense strategies such as tolerance (Duffin et al., 2020), or even defense ‘syndromes’ (Agrawal and Fishbein, 2006; Defossez et al., 2018), in wild seagrass populations. Duffin et al. (2021) subsequently combined all four markers when evaluating host immune activity relative to Labyrinthula load in the field.

Generally, the production of secondary metabolites is considered a cost to a plant, with multiple functions as a potential means to offset its toll (Erb and Kliebenstein, 2020). Still, the production of defensive compounds in plants afflicted with SWD, along with reduced host photosynthetic ability (Ralph and Short, 2002), may in turn reduce host energetic resources such as non-structural carbohydrates. As valuable measures of seagrass fitness broadly, assessments of non-structural carbohydrates (i.e. starch and/or sugar content) have also been applied to SWD systems. Brakel et al. (2019) found limited effects of Labyrinthula inoculation on fitness measures, including rhizome starch, leaf starch, and leaf sucrose concentrations, in eelgrass relative to other stressors (temperature, light, and salinity) in a factorial, multi-stressor experiment. In contrast, Graham et al. (2021) found that rhizome sugar content decreased with the severity of SWD in field-collected eelgrass plants.

Long-chain polyunsaturated FA (LCPUFA), especially the omega-3 FA docosahexaenoic acid (DHA) and eicosapentaenoic acid (EPA), are valuable dietary constituents for human and aquatic life (Parrish, 2009). As effective synthesizers of LCPUFA, thraustochytrids have received considerable research and industry attention (Marchan et al., 2018). Labyrinthulids are also known to produce LCPUFA, notably DHA, with much research focused on potential industrial applications of high-LCPUFA producing strains (Sakata et al., 2000; Kumon et al., 2002; Kumon et al., 2003; Kumon et al., 2005; Kumon et al., 2006; Wang et al., 2019). However, most FA studies on Labyrinthulids use single-taxon microbial (bacteria or yeasts) or derived (serum, yeast, and/or peptone-based) substrates for culture (Sakata et al., 2000; Kumon et al., 2002; Kumon et al., 2003; Kumon et al., 2005; Kumon et al., 2006; Wang et al., 2019; Yoshioka et al., 2019). Consequently, little is known about Labyrinthula LCPUFA production in situ.

Pathogenic Labyrinthula can produce substantial amounts of DHA (Yoshioka et al., 2019). Grown on a modified serum seawater agar and an eelgrass-based agar (see Growth Media), Labyrinthula isolates from diseased Z. marina produced DHA as their dominant FA. In both laboratory-inoculated and field-collected eelgrass tissues, DHA content was greater in diseased tissue than healthy tissue, suggesting that pathogenic Labyrinthula produces DHA in situ (Yoshioka et al., 2019). For Yoshioka et al. (2019), plant DHA content associated with disease was detectable but modest. Fatty acid changes in diseased plants are likely subject to other variables that may affect the plant-pathogen relationships. Lastly, surface metabolites, including FA of Z. marina, can inhibit or modulate microbes that in turn may affect disease (Papazian et al., 2019), suggesting more work is needed to understand production and the role of microbes in modulating FA levels in plants.