Fungi, bacteria and oomycota opportunistically isolated from the seagrass, Zostera marina.

Fungi in the marine environment are often neglected as a research topic, despite that fungi having critical roles on land as decomposers, pathogens or endophytes. Here we used culture-dependent methods to survey the fungi associated with the seagrass, Zostera marina, also obtaining bacteria and oomycete isolates in the process. A total of 108 fungi, 40 bacteria and 2 oomycetes were isolated. These isolates were then taxonomically identified using a combination of molecular and phylogenetic methods. The majority of the fungal isolates were classified as belonging to the classes Eurotiomycetes, Dothideomycetes, and Sordariomycetes. Most fungal isolates were habitat generalists like Penicillium sp. and Cladosporium sp., but we also cultured a diverse set of rare taxa including possible habitat specialists like Colletotrichum sp. which may preferentially associate with Z. marina leaf tissue. Although the bulk of bacterial isolates were identified as being from known ubiquitous marine lineages, we also obtained several Actinomycetes isolates and a Phyllobacterium sp. We identified two oomycetes, another understudied group of marine microbial eukaryotes, as Halophytophthora sp. which may be opportunistic pathogens or saprophytes of Z. marina. Overall, this study generates a culture collection of fungi which adds to knowledge of Z. marina associated fungi and highlights a need for more investigation into the functional and evolutionary roles of microbial eukaryotes associated with seagrasses.

Introduction

Despite their global importance in terrestrial systems, the diversity, function, evolution, and global importance of fungi in the marine environment remains understudied. There are only ∼1100 currently accepted species of marine fungi despite estimates that true diversity is much higher, at 10,000 or more species [1, 2]. It is well known that fungi play vital roles in land plant health and fitness (e.g. as pathogens or endophytes), and although much less is known about fungi in aquatic ecosystems, it is thought they have important roles in organic matter degradation and food web dynamics [3]. Thus, it is likely that fungi engage in similarly vital functional roles when associated with marine plants, like seagrasses.

Seagrasses are fully submerged marine angiosperms and are foundation species in coastal ecosystems. Seagrass beds perform important ecosystem services and can store carbon over very long timescales in their above and below ground tissues and in surrounding sediments (i.e. "blue carbon") [4]. Unfortunately, seagrass beds are threatened by human-related activities such as pollution, climate change and coastal development, and restoration efforts thus far have been mostly ineffective [5]. In addition to their global ecological importance, seagrasses also have a unique evolutionary history. Sometimes referred to as the “whales of the plant world”, seagrasses are a paraphyletic group of multiple lineages that convergently adapted to the marine environment between 70 and 100 million years ago [6, 7]. There are only ~60 species of seagrass compared to the ~250,000 species of terrestrial flowering plants, a testament to the strict selective pressure posed by re-entry to the marine environment. This work focuses on one widespread seagrass species, Zostera marina, which occurs throughout much of the Northern Hemisphere.

Previous work has characterized the composition and structure of the bacterial community associated with Z. marina and found the community to be distinct for different seagrass tissues (e.g., roots, leaves, rhizomes) [8-10]. Many of the abundant bacteria found associated with Z. marina are thought to have important functions related to nitrogen and sulfur cycling [10-15] and several culture-dependent studies have obtained bacterial isolates associated with Z. marina, ranging from ubiquitous marine lineages to putative sulfate-reducers [16-25].

In comparison, less is known about the fungal community associated with Z. marina and seagrasses generally. Culture-based studies have found fungi associated with leaves, roots and rhizomes of seagrasses, but there is little agreement between studies about the taxonomic composition of these communities within and between seagrass species [26-42]. Recently culture-independent studies of seagrass-associated fungi have more thoroughly investigated the diversity of these microorganisms and highlighted a need to further understand factors affecting their biogeography and community dynamics [43-46]. However, these studies were severely hampered by a lack of representation of fungal sequences from the marine environment in public databases and found that taxonomic assignments could not be made for many fungal sequences associated with seagrasses. This suggests both a need to expand molecular knowledge of marine and seagrass-associated fungi in public databases and that seagrasses may harbor diverse and understudied fungal lineages.

Fungi and bacteria are not the only microbes associated with Z. marina and there are many other understudied microorganisms that likely have important roles in the seagrass ecosystem. For example, one culture-independent effort sought to investigate the composition of the entire eukaryote community associated with Z. marina, and found that the bacterial and eukaryotic epibiont communities were highly correlated [9]. Additionally, oomycetes [47, 48], protists [49], and viruses [50] have all been cultured in association with Z. marina and seagrass wasting disease is thought to be caused by the heterokont, Labyrinthula zosterae [51].

Here we used a culture-dependent survey followed by molecular and phylogenetic identification to (i) obtain and identify a diverse collection of fungi associated with Z. marina, (ii) place this fungal collection in the phylogenetic context of isolates obtained from other seagrass surveys, and (iii) compare and contrast the composition of this fungal collection to high throughput sequencing results of the composition of the fungal community associated with Z. marina from the same location.

Methods

Sample collection and isolation

Zostera marina tissues were collected under California Department of Fish and Wildlife Scientific Collecting Permit # SC 4874 granted to Dr. John J. Stachowicz. Individual Z. marina plants and associated sediment were collected opportunistically from Westside Point (GPS: 38°19'10.67"N, 123° 3'13.71"W) in Bodega Bay, CA during several sampling trips (October 2017, May 2018, July 2018, August 2018 and January 2019) at low tide using a 2.375 inch diameter modified PVC pipe as described in Ettinger & Eisen [43]. Generally, 2–3 cores were obtained per sampling trip. Bulk plant tissue from multiple Zostera marina plants of varying ages was also collected during these trips using gloves and placed in sterile plastic bags for use as both an inoculation source and for inclusion in media recipes. Seawater was also collected in autoclaved 1 L nalgene bottles for use in media recipes. All samples were kept cold on ice in a dark cooler for transport back to the lab. Plant tissues, sediment and seawater were stored at 4°C until plating could occur which happened within 4–24 hours of collection.

Plant tissues, sediment and seawater were plated on a variety of different media types. Only “green” leaf tissue was used as an isolation source. Generally for seagrass tissues (leaf, root or rhizome) this involved, (1) rinsing the tissue with autoclaved nanopure water to remove loosely associated sediment for ~30 sec, (2) using flame sterilized scissors to cut ~1 cm pieces of tissue, (3) placing a subset of these tissue segments directly on plates using flame sterilized tweezers (1–3 segments / plate), (4) taking another subset of tissue segments and placing these segments into 1.5 mL centrifuge tubes with 1 mL of autoclaved nanopure water, (5) vortexing the 1.5 mL centrifuge tubes for ~30 sec, (6) either smashing tissue segments using a sterile pestel or leaving the segments intact, and (7) directly plating intact tissue segments on media using flame sterilized tweezers (1–3 segments / plate) and pipetting 350 μL of wash liquid or of smashed tissue directly on plates. A further subset of tissue segments were subjected to a bleach treatment or were surface cleaned following step (2) above. For the bleach treatment, this involved taking tissue segments and, (1) immersing segments for 5 min in 1 mL 0.5% NaOCl (~10% bleach), (2) then in 1 mL of 95% EtOH for 1 min, (3) then in 1 mL autoclaved nanopure water for 3 min, and (4) directly plating intact bleached tissue segments on media using flame sterilized tweezers (1–3 segments / plate). For the surface cleaned tissues, this involved taking tissue segments and, (1) immersing segments in 500 μL 95% ethanol for ~5 sec, (2) then in 500 μL 0.5% NaOCl (∼10% bleach) for 2 min, (3) then in 500 μL 70% ethanol for 2 min, (4) then rinsing segments with autoclaved nanopure water for 1 min, and (5) directly plating intact surface cleaned tissue segments on media using flame sterilized tweezers (1–3 segments / plate). For sediment this process involved, (1) placing sediment into 1.5 mL centrifuge tubes with 1 mL of autoclaved nanopure water, (2) vortexing the tubes for ~30 sec, and (3) then pipetting 350 μL of sediment suspension directly onto plates. For seawater this process involved pipetting 350 μL of seawater directly onto plates.

A variety of media recipes were used to try to obtain a diverse collection of fungal isolates. These media included 1% tryptone agar (10 g tryptone, 10 g agar, 1 L distilled water), potato dextrose agar (PDA), potato carrot agar (PCA), palm oil media (12 g agar, 10 g dextrose, 10 g yeast extract, 3 g peptone, 2 g L-arginine, 10 mL Tween80,10 mL palm oil, 1 L distilled water, final pH: 8.0), lecithin media (12 g agar, 10 g dextrose, 10 g yeast extract, 3 g peptone, 2 g L-arginine, 10 mL Tween80, 0.7 g lecithin, 1 L distilled water, final pH: 8.0), malt extract agar (MEA; 30 g malt extract, 15 g agar, 1 L distilled water, final pH: 5.5), glucose yeast peptone agar (GYPA; 15 g agar, 5 g yeast extract, 5 g peptone, 40 g glucose,1 L distilled water), and a Zostera marina agar (20 g of leaves in 100 mL of 0.45 μM Millipore filtered natural aged seawater heated up to 60°C for 30 min, 18 g agar, 0.45 μM Millipore filtered natural aged seawater make up volume to 1L) inspired by Agar Posidonia from Panno et al. [33]. A variety of salt amendments were used including: adding no salt, adding varying amounts of instant ocean (8 g, 16 g, or 32 g) or substituting distilled water for 0.45 μM Millipore filtered natural aged seawater. All media was amended with 50 mg/mL ampicillin, with some media batches also amended with 50 mg/mL trimethoprim or 50 mg/mL streptomycin. Additionally, some media batches also included the addition of 5 g/L dehydrated crushed Z. marina leaf tissue. For the exact media conditions each isolate was grown on see S1 Table.

Plates were wrapped in parafilm to prevent contamination and incubated at room temperature (e.g. as in [27, 42]) in the dark (e.g. as in [26, 32]) in a cabinet drawer for a minimum of 4 weeks (e.g. as in [26, 27, 32, 33]), up to a maximum of 12 weeks. Plates were observed every 2–3 days for fungal growth. Fungal isolates were then sterilely subcultured onto new plates and the process repeated until we were confident we had a single isolate. We were confident when we had subcultured the organism three times each with consistent morphology and no signs of contamination. During the isolation process, all parent plates and subcultures for an organism were stored at 4°C for comparative purposes. Plates with contamination were tossed (e.g. with a morphology inconsistent with what had been previously observed or colonies not near tissues or areas that were streaked).

DNA extraction, Polymerase chain reaction (PCR) and Sanger sequencing

DNA was extracted from isolates using the MoBio PowerSoil DNA Isolation kit (MO BIO Laboratories, Inc., Carlsbad, CA, United States) with minor changes to the manufacturer’s protocol as follows. To improve fungal lysis, samples were heated at 70°C for 10 minutes between steps 4 and 5. For step 5, samples were bead beaten on the homogenize setting for 2 minutes using a mini-bead beater (BioSpec Products). For a subset of isolates DNA was instead extracted with either the Qiagen Plant DNeasy (QIAGEN, Hilden, Germany), the Qiagen DNeasy PowerSoil Pro Kit (QIAGEN, Hildren, Germany) or the Zymo Xpedition Fungal/Bacterial DNA Mini Prep (Zymo Research Inc, Irvine, CA, United States) according to the manufacturer's instructions. The reason for the discrepancy between which DNA extraction kit was used is that we initially tried several different DNA extraction kits, before finding that the MoBio PowerSoil DNA Isolation kit provided the best DNA yield and subsequently, extracting isolates only with that kit moving forward. For the DNA extraction kit used for each isolate see S1 Table.

Polymerase chain reaction (PCR) was performed using Taq DNA Polymerase (QIAGEN, Hilden, Germany). Initially, PCR was performed on DNA from all isolates to amplify the fungal ITS-28S rRNA gene region. For isolates where PCR was not successful after three attempts, we then attempted to amplify the bacterial 16S rRNA gene. A few samples that had successful amplification for the bacterial 16S rRNA gene had close matches in NCBI GenBank to oomycete mitochondria, so in these cases we then attempted to amplify the oomycete 28S rRNA gene.

The fungal ITS-28S rRNA gene region was obtained using the ITS5 [52] and LR3 [53] primer set, the bacterial 16S rRNA gene was obtained using the 27F [54] and 1391R [55] primer set, and the oomycete 28S rRNA gene was obtained using the LR0R [56] and Un-Lo28S1220 [57] primer set. When amplifying the fungal ITS-28S rRNA gene region, PCR was performed with the following conditions: 95°C for 5 minutes, 35 cycles at 94°C for 30 seconds, 52°C for 15 seconds, 72°C for 1 minute, and a final extension at 72°C for 8 minutes [58]. When amplifying the bacterial 16S rRNA gene, PCR was performed with the following protocol: 95°C for 3 minutes, 40 cycles at 95°C for 15 seconds, 54°C for 30 seconds, 72°C for 1 minute and 30 seconds, and a final extension at 72°C for 5 minutes (modified from [59]). When amplifying the oomycete 28S rRNA gene, PCR was performed with the following protocol: 94°C for 4 minutes, 35 cycles at 94°C for 30 seconds, 57°C for 30 seconds, 72°C for 30 sec, and a final extension at 72°C for 10 minutes (adapted from Bourret et al. [60]).

PCR products were visualized on 2% agarose E-gels (Invitrogen, Carlsbad, CA, United States). PCR products were then purified using the Nucleospin Gel and PCR kit (QIAGEN, Hilden, Germany) and quantified using the Qubit dsDNA HS Assay Kit (Invitrogen, Carlsbad, CA, United States). The PCR products were sequenced using the Sanger method by the UC Davis College of Biological Sciences UCDNA Sequencing Facility (http://dnaseq.ucdavis.edu/). The resulting ABI files were visualized and consensus sequences were generated using seqtrace v. 0.9.0 [60] following the Swabs to Genomes workflow [59]. Consensus sequences for the PCR products were deposited at NCBI Genbank under the following accession no. MN543905-MN544012 for the fungal ITS-28S rRNA gene region, MN931878-MN931917 for the bacterial 16S rRNA gene, and MN944508-MN944509 for the oomycete 28S rRNA gene.

Taxonomic analyses

Preliminary taxonomic assignment of sequences from the PCR products generated above were obtained by comparing the best results (or “top match”) across three methods to obtain a consensus assignment. The three methods included (1) using NCBI’s Standard Nucleotide BLAST’s megablast option against the nr/nt database with default settings for all isolates and against the 16S ribosomal RNA sequence database for bacterial isolates, (2) using the Ribosomal Database Project (RDP) classifier with the appropriate respective database (e.g. the 16S rRNA training set for bacteria, the Fungal LSU, WARCUP and UNITE datasets for fungi, the Fungal LSU for oomycetes) and default settings, (3) using the SILVA Alignment, Classification and Tree (ACT) service with the appropriate database (SSU for bacteria, LSU for fungi and oomycetes) and default settings [61-65]. Taxonomic assignments for isolates and associated isolation conditions were then imported into R (v. 3.6.0) for visualization and analysis using the following packages: ggplot2 (v. 3.2.1), dplyr (v. 0.8.4), reshape (v. 0.8.8), patchwork (v. 1.0.0), and tidyverse (v. 1.3.0) [66-70] (S1 File).

Phylogenetic analyses of fungal isolates

Sequences closely related to the fungal ITS-28S rRNA gene PCR products generated above were identified using NCBI’s Standard Nucleotide BLAST’s megablast option with default settings to further confirm fungal taxonomy through phylogenetic placement (S2 Table). Additionally, we wanted to place the Z. marina associated fungal isolates in the context of the phylogenetic diversity of available other seagrass-associated fungal isolates. To this end, we performed a literature search to obtain, to our knowledge at the time of the search, all available 28S rRNA sequences obtained from seagrass associated fungal isolates for inclusion in phylogenetic analyses (S3 Table) [26, 27, 42, 71, 72]. Finally, to provide a further framework for these phylogenies, as well as appropriate outgroup taxa, we downloaded the available 28S rRNA sequences previously used in James et al. [73, 74] (S4 Table).

Using the sequences listed in Tables 1 and S2–S4, we generated four different sequence alignments, (1) an alignment to investigate seagrass isolates in the Basidiomycota and Mucoromycota, (2) an alignment to investigate seagrass isolates in the Eurotiomycetes class in the Ascomycota phylum, (3) an alignment to investigate seagrass isolates in the Sordariomycetes class in the Ascomycota phylum, and (4) an alignment to investigate seagrass isolates in the Dothideomycetes class in the Ascomycota phylum.

Table 1

Fungi isolated from the seagrass, Zostera marina.

Strain	Isolation Source	Class	Order	Putative Taxonomy	GenBank Accession (ITS-LSU)	Genus includes known marine fungi	Genus detected in ITS amplicon data
CLE116	Leaf	Dothideomycetes	Capnodiales	Cladosporium sp.	MN543969	yes	yes
CLE118	Leaf	Dothideomycetes	Capnodiales	Cladosporium sp.	MN543970	yes	yes
CLE127	Leaf	Dothideomycetes	Capnodiales	Cladosporium sp.	MN543975	yes	yes
CLE152	Leaf	Dothideomycetes	Capnodiales	Cladosporium sp.	MN543985	yes	yes
CLE37	Leaf	Dothideomycetes	Capnodiales	Cladosporium sp.	MN543925	yes	yes
CLE39	Leaf	Dothideomycetes	Capnodiales	Cladosporium sp.	MN543926	yes	yes
CLE109	Root	Dothideomycetes	Capnodiales	Cladosporium sp.	MN543962	yes	yes
CLE14	Root	Dothideomycetes	Capnodiales	Cladosporium sp.	MN543914	yes	yes
CLE90	Root	Dothideomycetes	Capnodiales	Cladosporium sp.	MN543951	yes	yes
CLE157	Seawater	Dothideomycetes	Capnodiales	Cladosporium sp.	MN543992	yes	yes
CLE121	Sediment	Dothideomycetes	Capnodiales	Cladosporium sp.	MN543973	yes	yes
CLE103	Leaf	Dothideomycetes	Capnodiales	Ramularia sp.	MN543956	no	yes
CLE164	Leaf	Dothideomycetes	Capnodiales	Ramularia sp.	MN544001	no	yes
CLE32	Leaf	Dothideomycetes	Capnodiales	Ramularia sp.	MN543922	no	yes
CLE81	Leaf	Dothideomycetes	Capnodiales	Ramularia sp.	MN543944	no	yes
CLE89	Leaf	Dothideomycetes	Capnodiales	Ramularia sp.	MN543950	no	yes
CLE158	Rhizome	Dothideomycetes	Capnodiales	Ramularia sp.	MN543993	no	yes
CLE160	Rhizome	Dothideomycetes	Capnodiales	Ramularia sp.	MN543996	no	yes
CLE1	Root	Dothideomycetes	Capnodiales	Ramularia sp.	MN543907	no	yes
CLE111	Root	Dothideomycetes	Capnodiales	Ramularia sp.	MN543964	no	yes
CLE112	Root	Dothideomycetes	Capnodiales	Ramularia sp.	MN543965	no	yes
CLE122	Sediment	Dothideomycetes	Capnodiales	Ramularia sp.	MN543974	no	yes
CLE104	Leaf	Dothideomycetes	Dothideales	Aureobasidium sp.	MN543957	yes	yes
CLE102	Leaf	Dothideomycetes	Pleosporales	Pleosporales sp.	MN543955	NA	NA
CLE3	Leaf	Dothideomycetes	Pleosporales	Pleosporales sp.	MN543909	NA	NA
CLE55	Leaf	Dothideomycetes	Pleosporales	Pleosporales sp.	MN543927	NA	NA
CLE56	Leaf	Dothideomycetes	Pleosporales	Pleosporales sp.	MN543942	NA	NA
CLE57	Leaf	Dothideomycetes	Pleosporales	Pleosporales sp.	MN543928	NA	NA
CLE159	Root	Dothideomycetes	Pleosporales	Pleosporales sp.	MN543995	NA	NA
CLE2	Rhizome	Dothideomycetes	Pleosporales	Pleosporales sp.	MN543908	NA	NA
CLE101	Leaf	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543954	yes	yes
CLE12	Leaf	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543912	yes	yes
CLE128	Leaf	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543976	yes	yes
CLE129	Leaf	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543977	yes	yes
CLE13	Leaf	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543913	yes	yes
CLE130	Leaf	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543978	yes	yes
CLE131	Leaf	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543979	yes	yes
CLE133	Leaf	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543981	yes	yes
CLE139	Leaf	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543983	yes	yes
CLE151	Leaf	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543984	yes	yes
CLE163	Leaf	Eurotiomycetes	Eurotiales	Penicillium sp.	MN544000	yes	yes
CLE17	Leaf	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543916	yes	yes
CLE171	Leaf	Eurotiomycetes	Eurotiales	Penicillium sp.	MN544004	yes	yes
CLE172	Leaf	Eurotiomycetes	Eurotiales	Penicillium sp.	MN544005	yes	yes
CLE174	Leaf	Eurotiomycetes	Eurotiales	Penicillium sp.	MN544007	yes	yes
CLE175	Leaf	Eurotiomycetes	Eurotiales	Penicillium sp.	MN544008	yes	yes
CLE20	Leaf	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543918	yes	yes
CLE34	Leaf	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543923	yes	yes
CLE35	Leaf	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543924	yes	yes
CLE42	Leaf	Eurotiomycetes	Eurotiales	Penicillium sp.	MN544012	yes	yes
CLE62	Leaf	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543933	yes	yes
CLE66	Leaf	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543936	yes	yes
CLE73	Leaf	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543940	yes	yes
CLE83	Leaf	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543946	yes	yes
CLE84	Leaf	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543947	yes	yes
CLE95	Leaf	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543953	yes	yes
CLE132	Leaf	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543980	yes	yes
CLE106	Rhizome	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543959	yes	yes
CLE107	Rhizome	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543960	yes	yes
CLE108	Rhizome	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543961	yes	yes
CLE113	Rhizome	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543966	yes	yes
CLE114	Rhizome	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543967	yes	yes
CLE115	Rhizome	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543968	yes	yes
CLE145	Rhizome	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543994	yes	yes
CLE155	Rhizome	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543988	yes	yes
CLE25	Rhizome	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543919	yes	yes
CLE26	Rhizome	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543920	yes	yes
CLE41	Rhizome	Eurotiomycetes	Eurotiales	Penicillium sp.	MN544011	yes	yes
CLE85	Rhizome	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543948	yes	yes
CLE110	Root	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543963	yes	yes
CLE15	Root	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543915	yes	yes
CLE161	Root	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543997	yes	yes
CLE162	Root	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543998	yes	yes
CLE58	Root	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543929	yes	yes
CLE59	Root	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543930	yes	yes
CLE60	Root	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543931	yes	yes
CLE68	Root	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543937	yes	yes
CLE119	Sediment	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543971	yes	yes
CLE120	Sediment	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543972	yes	yes
CLE138	Sediment	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543982	yes	yes
CLE167	Sediment	Eurotiomycetes	Eurotiales	Penicillium sp.	MN544003	yes	yes
CLE173	Sediment	Eurotiomycetes	Eurotiales	Penicillium sp.	MN544006	yes	yes
CLE18	Sediment	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543917	yes	yes
CLE64	Sediment	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543935	yes	yes
CLE69	Sediment	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543938	yes	yes
CLE70	Sediment	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543939	yes	yes
CLE77	Sediment	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543941	yes	yes
CLE80	Sediment	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543943	yes	yes
CLE31	Seawater	Eurotiomycetes	Eurotiales	Penicillium sp.	MN543921	yes	yes
CLE144	Seawater	Eurotiomycetes	Eurotiales	Talaromyces sp.	MN543991	yes	yes
CLE82	Seawater	Eurotiomycetes	Eurotiales	Talaromyces sp.	MN543945	yes	yes
CLE92	Seawater	Eurotiomycetes	Eurotiales	Talaromyces sp.	MN543952	yes	yes
CLE154	Rhizome	Microbotryomycetes	Sporidiobolales	Rhodotorula sp.	MN543987	yes	yes
CLE88	Leaf	Sordariomycetes	Glomerellales	Colletotrichum sp.	MN543949	no	yes
CLE143	Rhizome	Sordariomycetes	Glomerellales	Colletotrichum sp.	MN543989	no	yes
CLE4	Rhizome	Sordariomycetes	Glomerellales	Colletotrichum sp.	MN543905	no	yes
CLE5	Rhizome	Sordariomycetes	Glomerellales	Colletotrichum sp.	MN543906	no	yes
CLE7	Leaf	Sordariomycetes	Hypocreales	Acrostalagmus sp.	MN543911	yes	yes
CLE63	Rhizome	Sordariomycetes	Hypocreales	Emericellopsis sp.	MN543934	yes	no
CLE105	Leaf	Sordariomycetes	Hypocreales	Hypocreales sp.	MN543958	NA	NA
CLE153	Leaf	Sordariomycetes	Hypocreales	Hypocreales sp.	MN543986	NA	NA
CLE61	Root	Sordariomycetes	Hypocreales	Hypocreales sp.	MN543932	NA	NA
CLE6	Rhizome	Sordariomycetes	Hypocreales	Sarocladium sp.	MN543910	yes	yes
CLE146	Leaf	Sordariomycetes	Hypocreales	Trichoderma sp.	MN543999	yes	yes
CLE165	Leaf	Tremellomycetes	Filobasidiales	Naganishia sp.	MN544002	yes	yes
CLE156	Rhizome	Ustilaginomycetes	Ustilaginales	Pseudozyma sp.	MN543990	yes	no
CLE40	Rhizome	Ustilaginomycetes	Ustilaginales	Pseudozyma sp.	MN544010	yes	no
CLE24	Leaf	Mucoromycetes	Mucorales	Absidia cylindrospora	MN544009	no	no

Here we report the taxonomic information for each fungal isolate (Class, Order) and the putative taxonomy, provide the GenBank accession number for the ITS-28S rRNA gene sequence for each isolate, and report on the isolation source the isolate was obtained from (e.g. leaf, rhizome, root, seawater or sediment). We also report on whether the genus of each isolate includes marine fungal representatives based on the consensus compiled in Jones et al. [2] and whether the genus of each isolate was detected in the ITS amplicon data in Ettinger & Eisen [43]. Organisms for which a taxonomic identification below the order level was not possible, have a “NA” value for these columns. It is important to note that there is likely significant biological variation within the genera reported here (e.g. among Penicillium), such that finding members of these genera should not be interpreted as meaning that the specific variants isolated here have the same biology as variants found to be member of the same genera in other datasets.

Each of the four sequence alignments was generated using MAFFT (v. 7.402) [75] with default parameters on the CIPRES Science Gateway web server [76]. The alignments were trimmed using trimAl (v.1.2) with the -gappyout method [77] and then manually inspected with JalView [78]. Sequence alignments were then further trimmed to the D1/D2 regions of the 28S rRNA gene with trimAl using the select option (e.g. Basidiomycota / Mucoromycota alignment {614–2899 }, Eurotiomycetes alignment {0–569 }, Sordariomycetes alignment {501–1224 }, and Dothideomycetes alignment {0–429, 993–1755 }). Spurious sequences (e.g. sequences which contained few or no nucleotides after trimming) were then removed with trimAl using -resoverlap .75 -seqoverlap 50. The resulting alignments contained: 80 sequences with 614 positions (Basidiomycota / Mucoromycota), 91 sequences with 509 positions (Eurotiomycetes), 96 sequences with 501 positions (Sordariomycetes), and 107 sequences with 563 positions (Dothideomycetes).

JModelTest2 (v. 2.1.10) was run with the number of substitution schemes (-s) set to 3 (JC/F81, K80/HKY, SYM/GTR) and then otherwise default parameters on the CIPRES Science Gateway web server to select a best-fit model of nucleotide substitution for use in phylogenetic analyses for each alignment [79, 80]. The best-fit model based on the Akaike Information Criterion values for all alignments was the GTR + I + G evolutionary model.

Using the CIPRES Science Gateway web server, Bayesian phylogenetic inference for each alignment was performed using MrBayes (v. 3.2.2) with four incrementally heated simultaneous Monte Carlo Markov Chains (MCMC) run over 10,000,000 generations. The analysis stopped early if the optimal number of generations to reach a stop value of 0.01 or less for the convergence diagnostic was achieved [81]. This occurred for the Eurotiomycetes, Sordariomycetes and Dothideomycetes alignments at 2,150,000 generations, 1,375,000 generations and 2,140,000 generations, respectively. The Basidiomycota / Mucoromycota alignment ran for the full 10,000,000 generations, only achieving an average standard deviation of split frequencies of 0.049. The first 25% of trees generated for each alignment were discarded as burn-in and for the remaining trees, a majority rule consensus tree was generated and used to calculate the Bayesian Posterior Probabilities. The resulting phylogenies were then visualized with the ggtree (v. 2.0.1), treeio(v. 1.11.2), ggplot2 (v. 3.2.1), and tidyverse (v. 1.3.0) packages in R (v. 3.6.0) and clade labels were added in Adobe Photoshop CS6 [66, 67, 82–85] (S1 File). Alignments and phylogenies generated here were deposited to Dryad [86].

Comparisons to ITS amplicon data from Ettinger & Eisen [43]

To compare to high throughput sequencing data associated with Z. marina from the same location (Westside Point, Bodega Bay, CA), we utilized an amplicon sequence variant (ASV) dataset previously analysed in Ettinger & Eisen [43]. Specifically, we are using the subset ASV table that was used to investigate differences between bulk sample types. Briefly, this ASV table was previously subset to a depth of 10,000 sequences and included 49 samples from four sample types: leaf epiphytes (n = 13), root epiphytes (n = 14), rhizome epiphytes (n = 7), and sediment (n = 15). We then used this ASV to make comparisons to the fungal taxa isolated in this study. To investigate whether fungal genera isolated in this study were also detected in the high throughput sequencing data, we generated a list of the unique genera found in the ASV table and compared it to the list of fungal genera isolated here. To investigate whether the fungal genera isolated in this study were detected from the same Z. marina tissues, we collapsed the ASV table to the genus level using the tax_glom function in phyloseq. We then subsampled the ASV table to only include the genera of fungi isolated in this study, transformed this ASV table represent presence / absence and visualized a comparison of the distribution of these genera across sample types (leaf, root, rhizome, sediment) to the distribution of these genera across isolation sources (leaf, root, rhizome, sediment). To investigate the mean relative abundance of the fungal orders isolated in this study in the high throughput sequencing data, we collapsed the ASV table to the order level using the tax_glom function in phyloseq. We then subsampled the ASV table to only include the orders of fungi isolated in this study and visualized the distribution of these orders across sample types (leaf, root, rhizome, sediment). These analyses were performed in R (v. 3.6.0) using the ggplot2 (v. 3.2.1), dplyr (v. 0.8.4), reshape (v. 0.8.8), patchwork (v. 1.0.0), phyloseq (v. 1.30.0) and tidyverse (v. 1.3.0) packages [66–69, 82, 87, 88] (S1 File).

Results

Isolation efficacy

A total of 160 plates were initially inoculated, 81 with leaves (nwhole = 44, ncrushed = 22, nwashes = 11, nbleached = 2, nsurface cleaned = 2), 38 with rhizomes (nwhole = 13, ncrushed = 8, nwashes = 2, nbleached = 2, nsurface cleaned = 2), 27 with roots (nwhole = 13, ncrushed = 8, nwashes = 2, nbleached = 2, nsurface cleaned = 2), 4 with seawater, and 10 with sediment (S1 and S2 Figs). Microbial growth was observed on 135 plates (84.4% of all inoculated plates). We subcultured 1–5 organisms from all plates with observed microbial growth. However, we only obtained isolates that met our criteria for putatively being single organisms (e.g. which had been subcultured three times each with consistent morphology and no signs of contamination) from 86 of these plates (63.7% of plates with observed growth, 53.8% of all inoculated plates). No isolates were ultimately obtained from bleached or surface cleaned tissues and only one isolate was obtained from Zostera marina agar.

In total 185 putatively anexic microbial isolates were obtained. Of these 185 isolates, we were able to generate PCR products for 176 isolates to send for Sanger sequencing for taxonomic identification. Despite multiple attempts we were unable to generate PCR products for 9 isolates across all primer sets tried here (possibly due to primer mismatch and/or too low concentrations of DNA). Of the 170 isolates where PCR products were sent for sequencing, we received good quality sequencing results for and were able to taxonomically identify 150 isolates. For the 26 isolates where sequencing either failed, was low quality or appeared mixed, 17 appeared to be bacterial in origin, 4 appeared to be fungal and 5 were too poor quality to identify (e.g. comprised of only N’s) based on searches using NCBI’s Standard Nucleotide BLAST’s megablast option with default settings.

Taxonomic diversity of fungi isolated from Z. marina

In an attempt to cultivate a wide diversity of fungal isolates, we used a variety of media types including several which had been used previously to isolate fungi from seagrasses (e.g. PDA [26, 27, 42], GPYA [33], MEA [32]). A total of 108 fungal isolates were obtained, with the majority cultured from Z. marina leaf tissue (n = 51), resulting in a range of morphological diversity (Fig 1). The rest of isolates were cultured from rhizome tissue (n = 23), root tissue (n = 16), associated sediment (n = 13), and seawater (n = 5) (Figs 2 and S3 and S4).

Fig 1

Microbes isolated from the seagrass, Zostera marina.

An example of the morphological diversity of microbial isolates (bacteria, fungi and oomycota) associated with the seagrass, Z. marina. Depicted plates were arbitrarily chosen to depict the morphological diversity of the isolates cultured in this study. Putative taxonomy of isolates shown: (a) Penicillium sp. CLE73, (b) Cladosporium sp. CLE116, (c) Colletotrichum sp. CLE5, (d) Hypocreales sp. CLE105, (e) unidentified microorganism, (f) Penicillium sp. CLE130, (g) Penicillium sp. CLE68, (h) Halophytophthora sp. CLE94, (i) Pleosporales sp CLE57, (j) unidentified microorganism, (k) Pleosporales sp. CLE102, (l) Penicillium sp. CLE26, (m) Cladosporium sp. CLE118, (n) Ramularia sp. CLE122, (o) Pseudoalteromonas sp. CLE126, (p) Talaromyces sp. CLE92, (q) Colletotrichum sp. CLE4, (r) Talaromyces sp. CLE82, (s) unidentified microorganism, (t) Acrostalagmus sp. CLE7, (u) Ramularia sp. CLE1, (v) Pleosporales sp. CLE56, (w) Penicillium sp. CLE77, (x) Ramularia sp. CLE112, (y) Penicillium sp. CLE106, (z) unidentified microorganism, (aa) Streptomyces sp. CLE117, (ab) Penicillium sp. CLE114, (ac) Cladosporium sp. CLE127, and (ad) Penicillium sp. CLE110. Unidentified microorganisms were unable to be identified using molecular methods (i.e. a PCR product was not successfully generated).

Fig 2

Distribution of counts of fungal isolates across isolation sources.

A histogram representing the number of fungal isolates grouped by order and colored by isolation source (leaf, rhizome, root, seawater or sediment). The numbers included on each bar represent the count of isolates obtained from that particular isolation source.

Almost all of the fungal isolates were taxonomically classified as belonging to the Ascomycota (n = 103), with the remaining five isolates classified as Basidiomycota (n = 4) and Mucoromycota (n = 1), respectively (Table 1). Within the Ascomycota, isolates were further identified as being in three classes: the Eurotiomycetes (n = 62), Dothideomycetes (n = 30), and Sordariomycetes (n = 11).

Eurotiomycetes isolates were further taxonomically classified as Penicillium sp. (n = 59) and Talaromyces sp. (n = 3). Sordariomycetes isolates were putatively classified as Colletotrichum sp. (n = 4), Acrostalagmus sp. (n = 1), Emericellopsis sp. (n = 1), Sarocladium sp. (n = 1), Trichoderma sp. (n = 1), and unidentified Hypocreales sp. (n = 3). Dothideomycetes isolates were classified as Cladosporium sp. (n = 11), Ramularia sp. (n = 11), Aureobasidium sp. (n = 1), and unidentified Pleosporales sp. (n = 7). Basidiomycota isolates were putatively classified as Pseudozyma sp. (n = 2), Rhodotorula sp. (n = 1), and Naganishia sp. (n = 1). The single Mucoromycota isolate was putatively classified as Absidia cylindrospora.

We observed a positive relationship between the number of tissue types and number of media types a fungal genus was isolated from (R = 0.86; S5 Fig) which we hypothesize may indicate that some fungal genera are habitat generalists. A similar positive relationship is also observed between the number of tissue types and the number of salt sources (R = 0.92) as well as between the number of media types and number of salt sources (R = 0.87). However, we did not perform any experiments to confirm this pattern. We also did not always attempt to control for effort (e.g. plating the same number of tissue segments on all media types [S1 and S2 Figs]). Therefore, we suggest that these positive relationships be interpreted with caution.

Taxonomic diversity of bacteria and oomycota isolated from Z. marina

Our intent here was to isolate fungi which was why we included broad spectrum antibiotics in our culturing media to help eliminate bacteria which might be associated with Z. marina. However, we still cultivated and identified 40 bacteria and 2 oomycetes. The bacteria are likely naturally resistant to the antibiotics used and the oomycetes, as eukaryotes, are unlikely to be affected by their presence in the media. As with the fungal cultivation results, the majority of bacterial isolates were obtained from Z. marina leaf tissue (n = 17). The rest of the bacterial isolates were cultured from rhizome tissue (n = 9), root tissue (n = 7), associated sediment (n = 5), and seawater (n = 2) (S6 Fig).

Bacterial isolates were taxonomically identified as belonging to the Actinobacteria (n = 4), Firmicutes (n = 2), Bacteroidetes (n = 2), and Proteobacteria (n = 33) (Table 2). The two Firmicute isolates were further classified as Bacillus sp., the two Bacteroidetes isolates as Joostella sp., and the Actinobacteria isolates as Streptomyces sp. (n = 2), Rhodococcus sp. (n = 1), and Isoptericola sp. (n = 1). The Proteobacteria isolates were classified as Vibrio sp. (n = 18), Pseudoalteromonas sp. (n = 8), Hafnia sp. (n = 2), Pseudomonas sp. (n = 1), Shewanella sp. (n = 1), Marinomonas sp. (n = 1), and Phyllobacterium sp. (n = 1).

Table 2

Bacteria isolated from the seagrass, Zostera marina.

Strain	Isolation Source	Class	Order	Putative Taxonomy	GenBank Accession (SSU)	Top BLAST match	BLAST % Identity	Top BLAST accession no.
CLE44	Leaf	Actinobacteria	Actinomycetales	Isoptericola sp.	MN931913	Isoptericola halotolerans	99.2	NR_043198.1
CLE150	Sediment	Actinobacteria	Actinomycetales	Rhodococcus sp.	MN931907	Rhodococcus erythropolis	99.92	NR_037024.1
CLE117	Leaf	Actinobacteria	Streptomycetales	Streptomyces sp.	MN931916	Streptomyces argenteolus	99.36	NR_112300.1
CLE43	Leaf	Actinobacteria	Streptomycetales	Streptomyces sp.	MN931912	Streptomyces beijiangensis	98.35	NR_112607.1
CLE16	Root	Alphaproteobacteria	Rhizobiales	Phyllobacterium sp.	MN931909	Phyllobacterium loti	95.73	NR_133818.1
CLE136	Sediment	Bacilli	Lactobacillales	Bacillus sp.	MN931897	Bacillus mycoides	99.61	NR_036880.1
CLE53	Sediment	Bacilli	Lactobacillales	Bacillus sp.	MN931915	Bacillus thuringiensis	99.92	NR_043403.1
CLE8	Rhizome	Flavobacteria	Flavobacteriales	Joostella sp.	MN931878	Joostella marina	99.3	NR_044346.1
CLE10	Root	Flavobacteria	Flavobacteriales	Joostella sp.	MN931908	Joostella marina	99.22	NR_044346.1
CLE126	Leaf	Gammaproteobacteria	Alteromonadales	Pseudoalteromonas sp.	MN931894	Pseudoalteromonas spiralis	99.46	NR_114801.1
CLE71	Leaf	Gammaproteobacteria	Alteromonadales	Pseudoalteromonas sp.	MN931884	Pseudoalteromonas spiralis	99.52	NR_114801.1
CLE74	Leaf	Gammaproteobacteria	Alteromonadales	Pseudoalteromonas sp.	MN931886	Pseudoalteromonas hodoensis	98.42	NR_126232.1
CLE140	Rhizome	Gammaproteobacteria	Alteromonadales	Pseudoalteromonas sp.	MN931899	Pseudoalteromonas hodoensis	98.98	NR_126232.1
CLE141	Rhizome	Gammaproteobacteria	Alteromonadales	Pseudoalteromonas sp.	MN931900	Pseudoalteromonas spiralis	99.14	NR_114801.1
CLE142	Rhizome	Gammaproteobacteria	Alteromonadales	Pseudoalteromonas sp.	MN931901	Pseudoalteromonas spiralis	99.92	NR_114801.1
CLE98	Rhizome	Gammaproteobacteria	Alteromonadales	Pseudoalteromonas sp.	MN931892	Pseudoalteromonas spiralis	99.3	NR_114801.1
CLE147	Sediment	Gammaproteobacteria	Alteromonadales	Pseudoalteromonas sp.	MN931903	Pseudoalteromonas hodoensis	98.6	NR_126232.1
CLE47	Rhizome	Gammaproteobacteria	Alteromonadales	Shewanella sp.	MN931882	Shewanella surugensis	97.78	NR_040950.1
CLE149	Seawater	Gammaproteobacteria	Enterobacteriales	Hafnia sp.	MN931906	Hafnia alvei	99.54	NR_112985.1
CLE87	Seawater	Gammaproteobacteria	Enterobacteriales	Hafnia sp.	MN931890	Hafnia alvei	99	NR_112985.1
CLE19	Leaf	Gammaproteobacteria	Oceanospirillales	Marinomonas sp.	MN931910	Marinomonas rhizomae	97.5	NR_116233.1
CLE28	Rhizome	Gammaproteobacteria	Pseudomonadales	Pseudomonas sp.	MN931911	Pseudomonas sabulinigri	97.69	NR_044415.1
CLE123	Leaf	Gammaproteobacteria	Vibrionales	Vibrio sp.	MN931893	Vibrio ostreicida	99.15	NR_133887.1
CLE125	Leaf	Gammaproteobacteria	Vibrionales	Vibrio sp.	MN931917	Vibrio lentus	99.68	NR_114982.1
CLE148	Leaf	Gammaproteobacteria	Vibrionales	Vibrio sp.	MN931904	Vibrio kanaloae	98.53	NR_114804.1
CLE170	Leaf	Gammaproteobacteria	Vibrionales	Vibrio sp.	MN931902	Vibrio alginolyticus	99.61	NR_122050.1
CLE176	Leaf	Gammaproteobacteria	Vibrionales	Vibrio sp.	MN931905	Vibrio penaeicida	96	NR_042121.1
CLE29	Leaf	Gammaproteobacteria	Vibrionales	Vibrio sp.	MN931879	Vibrio kanaloae	99.12	NR_114804.1
CLE30	Leaf	Gammaproteobacteria	Vibrionales	Vibrio sp.	MN931880	Vibrio kanaloae	98.65	NR_114804.1
CLE72	Leaf	Gammaproteobacteria	Vibrionales	Vibrio sp.	MN931885	Vibrio alginolyticus	98.69	NR_122050.1
CLE78	Leaf	Gammaproteobacteria	Vibrionales	Vibrio sp.	MN931889	Vibrio tasmaniensis	99.43	NR_036929.1
CLE65	Leaf	Gammaproteobacteria	Vibrionales	Vibrio sp.	MN931883	Vibrio penaeicida	96.66	NR_042121.1
CLE75	Rhizome	Gammaproteobacteria	Vibrionales	Vibrio sp.	MN931887	Vibrio kanaloae	99.45	NR_114804.1
CLE76	Rhizome	Gammaproteobacteria	Vibrionales	Vibrio sp.	MN931888	Vibrio kanaloae	99.14	NR_114804.1
CLE134	Root	Gammaproteobacteria	Vibrionales	Vibrio sp.	MN931895	Vibrio penaeicida	95.89	NR_042121.1
CLE135	Root	Gammaproteobacteria	Vibrionales	Vibrio sp.	MN931896	Vibrio penaeicida	95.89	NR_042121.1
CLE36	Root	Gammaproteobacteria	Vibrionales	Vibrio sp.	MN931881	Vibrio tasmaniensis	99.68	NR_036929.1
CLE48	Root	Gammaproteobacteria	Vibrionales	Vibrio sp.	MN931914	Vibrio kanaloae	98.96	NR_114804.1
CLE52	Root	Gammaproteobacteria	Vibrionales	Vibrio sp.	MN931891	Vibrio penaeicida	96.92	NR_042121.1
CLE137	Sediment	Gammaproteobacteria	Vibrionales	Vibrio sp.	MN931898	Vibrio anguillarum	98.92	NR_042509.1

Here we report the taxonomic information for each bacterial isolate (Class, Order) and the putative taxonomy, provide the GenBank accession number for the 16S rRNA gene sequence for each isolate, and report on the isolation source the isolate was obtained from (e.g. leaf, rhizome, root, seawater or sediment). We also report the taxonomic identity of the top BLAST match against NCBI’s targeted loci 16S ribosomal RNA sequence database, the BLAST % identity to the bacterial isolate and the GenBank accession number for the 16S rRNA gene sequence for the top BLAST match.

The two oomycete isolates were obtained from Z. marina leaf tissue and were both taxonomically identified as Halophytophthora sp. (Table 3).

Table 3

Oomycota isolated from the seagrass, Zostera marina.

Strain	Isolation Source	Class	Order	Putative Taxonomy	GenBank Accession (SSU)	Top BLAST match	BLAST % Identity	Top BLAST accession no.
CLE33	Leaf	Oomycota	Pythiales	Halophytophthora sp.	MN944508	Halophytophthora polymorphica	98.9	AY598669.1
CLE94	Leaf	Oomycota	Pythiales	Halophytophthora sp.	MN944509	Halophytophthora polymorphica	98.69	AY598669.1

Here we report the taxonomic information for each oomycete isolate (Class, Order) and the putative taxonomy, provide the GenBank accession number for the 28S rRNA gene sequence for each isolate, and report on the isolation source the isolate was obtained from (e.g. leaf, rhizome, root, seawater or sediment). We also report the taxonomic identity of the top BLAST match against NCBI’s nr/nt database, the BLAST % identity to the oomycete isolate and the GenBank accession number for the 28S rRNA gene sequence for the top BLAST match.

Phylogenetic comparison of fungal isolates across seagrass species

To confirm fungal isolate identity and investigate if Z. marina fungal isolates were closely related to fungal isolates obtained from other seagrass species, we built four phylogenetic trees, 1) a phylogeny of seagrass isolates in the Basidiomycota and Mucoromycota (Figs 3 and S7), (2) a phylogeny of seagrass isolates in the Eurotiomycetes class in the Ascomycota phylum (Figs 4 and S8), (3) a phylogeny of seagrass isolates in the Sordariomycetes class in the Ascomycota phylum (Figs 5 and S9), and (4) a phylogeny of seagrass isolates in the Dothideomycetes class in the Ascomycota phylum (Figs 6 and S10). The placements of isolates in these phylogenies were consistent with the taxonomic identities previously determined. Additionally, isolates that were only identified at the order level taxonomically (unidentified Pleosporales sp. and Hypocreales sp.) were not able to be confidently further identified via the phylogenetic methods used here. The closest phylogenetic relatives of unidentified Hypocreales sp. are other unidentified fungi in the order Hypocreales (Fig 5). While the unidentified Pleosporales sp. form an unresolved clade with members in the family Didymellaceae in the order Pleosporales (Fig 6).

Fig 3

Phylogenetic placement of seagrass fungal isolates in the Basidiomycota and Mucoromycota.

A molecular phylogeny of 28S rRNA genes for isolates in the Basidiomycota and Mucoromycota was constructed using Bayesian inference. This alignment was generated using MAFFT (v. 7.402) on the CIPRES Science Gateway web server, trimmed using trimAl (v.1.2) and a phylogenetic tree was inferred on the trimmed alignment with a GTR + I + G model using MrBayes (v. 3.2.2) [75–77, 81]. Displayed at each node as a circle in the tree are the Bayesian posterior probabilities (e.g. a black circle represents probabilities greater or equal to 90%, a grey circle represents probabilities greater or equal to 70%, a white circle represents probabilities less than 70%). The names of fungi isolated from Z. marina are shown in green, fungi isolated from other seagrass species are shown in black, and all other fungi are shown in grey. For visualization purposes, selected clades have been collapsed and the number of sequences within that clade is indicated. Collapsed clades are shown in green if the majority of sequences in the clade are from isolates associated with Z. marina, black if the majority of isolates are from other seagrass species, and grey otherwise. Clade names that are followed by an asterisk contain sequences from multiple sources. An expanded version of this phylogeny can be found in S7 Fig. The GenBank accession numbers of the sequences used to build this phylogeny can be found in Tables 1 and S2–S4.

Fig 4

Phylogenetic placement of seagrass fungal isolates in the Eurotiomycetes.

A molecular phylogeny of 28S rRNA genes for isolates in the Eurotiomycetes was constructed using Bayesian inference. This alignment was generated using MAFFT (v. 7.402) on the CIPRES Science Gateway web server, trimmed using trimAl (v.1.2) and a phylogenetic tree was inferred on the trimmed alignment with a GTR + I + G model using MrBayes (v. 3.2.2) [75–77, 81]. Displayed at each node as a circle in the tree are the Bayesian posterior probabilities (e.g. a black circle represents probabilities greater or equal to 90%, a grey circle represents probabilities greater or equal to 70%, a white circle represents probabilities less than 70%). The names of fungi isolated from Z. marina are shown in green, fungi isolated from other seagrass species are shown in black, and all other fungi are shown in grey. For visualization purposes, selected clades have been collapsed and the number of sequences within that clade is indicated. Collapsed clades are shown in green if the majority of sequences in the clade are from isolates associated with Z. marina, black if the majority of isolates are from other seagrass species, and grey otherwise. Clade names that are followed by an asterisk contain sequences from multiple sources. An expanded version of this phylogeny can be found in S8 Fig. The GenBank accession numbers of the sequences used to build this phylogeny can be found in Tables 1 and S2–S4.

Fig 5

Phylogenetic placement of seagrass fungal isolates in the Sordariomycetes.

A molecular phylogeny of 28S rRNA genes for isolates in the Sordariomycetes was constructed using Bayesian inference. This alignment was generated using MAFFT (v. 7.402) on the CIPRES Science Gateway web server, trimmed using trimAl (v.1.2) and a phylogenetic tree was inferred on the trimmed alignment with a GTR + I + G model using MrBayes (v. 3.2.2) [75–77, 81]. Displayed at each node as a circle in the tree are the Bayesian posterior probabilities (e.g. a black circle represents probabilities greater or equal to 90%, a grey circle represents probabilities greater or equal to 70%, a white circle represents probabilities less than 70%). The names of fungi isolated from Z. marina are shown in green, fungi isolated from other seagrass species are shown in black, and all other fungi are shown in grey. For visualization purposes, selected clades have been collapsed and the number of sequences within that clade is indicated. Collapsed clades are shown in green if the majority of sequences in the clade are from isolates associated with Z. marina, black if the majority of isolates are from other seagrass species, and grey otherwise. Clade names that are followed by an asterisk contain sequences from multiple sources. An expanded version of this phylogeny can be found in S9 Fig. The GenBank accession numbers of the sequences used to build this phylogeny can be found in Tables 1 and S2–S4.

Fig 6

Phylogenetic placement of seagrass fungal isolates in the Dothideomycetes.

A molecular phylogeny of 28S rRNA genes for isolates in the Dothideomycetes was constructed using Bayesian inference. This alignment was generated using MAFFT (v. 7.402) on the CIPRES Science Gateway web server, trimmed using trimAl (v.1.2) and a phylogenetic tree was inferred on the trimmed alignment with a GTR + I + G model using MrBayes (v. 3.2.2) [75–77, 81]. Displayed at each node as a circle in the tree are the Bayesian posterior probabilities (e.g. a black circle represents probabilities greater or equal to 90%, a grey circle represents probabilities greater or equal to 70%, a white circle represents probabilities less than 70%). The names of fungi isolated from Z. marina are shown in green, fungi isolated from other seagrass species are shown in black, and all other fungi are shown in grey. For visualization purposes, selected clades have been collapsed and the number of sequences within that clade is indicated. Collapsed clades are shown in green if the majority of sequences in the clade are from isolates associated with Z. marina, black if the majority of isolates are from other seagrass species, and grey otherwise. Clade names that are followed by an asterisk contain sequences from multiple sources. An expanded version of this phylogeny can be found in S10 Fig. The GenBank accession numbers of the sequences used to build this phylogeny can be found in Tables 1 and S2–S4.

We expected to see more shared taxonomic groups and phylogenetic clustering between the fungal isolates of Z. marina and those of other seagrass species than was observed in Figs 3–6. Many of the fungal isolates from Z. marina did not cluster with fungal isolates that had been previously cultivated in association with other seagrass species. One interpretation might be that each seagrass species harbors a unique fungal community. However, this could also be the result of slight differences in collection protocols, media recipes or other methodology involved in isolating fungi in the compared studies.

The fungal taxa that did have close relatives that were associated with other species included Penicillium sp. (Fig 4), Trichoderma sp. (Fig 5), Cladosporium sp. and Ramularia sp. (Fig 6). We note that Penicillium sp., Cladosporium sp. and Ramularia sp. are drivers of the positive relationship observed previously between the number of tissue types and number of media types a fungal genus was isolated from (S5 Fig).

Comparisons to ITS amplicon sequencing data from Ettinger & Eisen

We compared the diversity of the fungi isolated here to high throughput sequencing data associated with Z. marina from the same location (as previously analyzed in Ettinger & Eisen [43]. We found that the fungal genera isolated in this study were generally also detected in the sequencing data (Table 1). Only three genera were not detected in the sequencing data, Pseudozyma sp., Emericellopsis sp. and Absidia cylindrospora. The absence of these genera in the sequencing data may be methodological (e.g. they do not amplify with the primer set used to generate the sequencing data) or biological (e.g. due to seasonal variation).

We then investigated whether the fungal genera isolated in this study were also detected in association with the same types of samples (e.g. leaf, root, rhizome, sediment) in the sequencing data. We observed that many of rare (e.g. not as frequently isolated) genera were not consistently detected on the same sample type with both methods, whereas many of the abundant (e.g. more frequently isolated) genera were detected with both methods (Fig 7). When we looked at the mean relative abundance of the fungal orders isolated in this study in the high throughput sequencing data, we observed that the genera detected using both methods generally were in orders that had higher mean relative abundances in the seagrass ecosystem (S11 Fig). We also observed that some orders such as the Eurotiales (e.g. Penicillium sp.) and Capnodiales (e.g. Cladosporium sp. and Ramularia sp.) had similar mean relative abundances across all sample types. While other orders such as Glomerellales (e.g. Colletotrichum sp.) had a higher mean relative abundance on one sample type (e.g. leaves).

Fig 7

Comparison of the detection of a fungal genus across methods.

A heatmap representing a comparison of the detection of the presence / absence of fungal genera isolated in this study (using a culture-dependent method) and fungal genera identified in high throughput sequencing data from Ettinger & Eisen [43] (using a culture-independent method). For each fungal genera, we visualize if it was not detected (light grey), detected using only the culture-dependent method (medium grey), detected using only the culture-independent method (dark grey) or detected by both methods (black) for each sample type / isolation source (leaf, root, rhizome, sediment).

Discussion

Here, we generated a fungal collection of 108 isolates expanding understanding of the diversity of Z. marina associated fungi, while also underscoring how little we know about these understudied microorganisms. Generally, the taxonomic diversity observed in our cultivation efforts is consistent with that of other culture-dependent studies which found Eurotiomycetes, Dothideomycetes, and Sordariomycetes to be the main classes of fungi associated with seagrasses [26, 27]. This is also consistent with what is known of the diversity of fungal associations with terrestrial plants. Members of the Sordariomycetes and Dothideomycetes have been found to be the predominant members of land plant fungal endophyte communities [89], while Eurotiomycetes have been found to be the dominant members of freshwater plant communities [90].

Dark septate endophytes (DSE), particularly members of the Pleosporales within the Dothideomycetes (Fig 6), have been observed to form associations with several seagrass species [33, 42, 44, 71, 72, 91–93]. DSE are a morphological, not phylogenetic (e.g. not each other's closest relatives) group of plant associated fungi, and are largely uncharacterized. The most well described of these DSE associations is between the Mediteranean seagrass, Posidonia oceanica, and its dominant root-associated fungus, Posidoniomyces atricolor. This Pleosporales member has been found associated with changes in root hair development and can form ecto-mycorrhizal-like structures [42, 72, 92, 93]. Here, although we isolated seven members of the Pleosporales, none appeared to be close relatives to Posidoniomyces atricolor.

Chytridiomycota were found to be prevalent members of the Z. marina leaf microbiome in Ettinger & Eisen [43], however, no chytrids were cultured in this study. This is likely because the isolation methods used here favor cultivation of Dikarya. Alternative methods and media recipes should be utilized (e.g. baiting) to isolate representatives of these important members of the fungal community. Similarly we note that the methods used here would fail to culture fungi involved in obligate associations with seagrasses. In these cases, a combination of microscopy and/or cell sorting for directed cultivation or sequencing might prove valuable for assessing the functional roles of these organisms to the seagrass ecosystem. Finally, in this study, we only attempted to cultivate aerobic fungi, but there could be anaerobic fungi living in these ecosystems as well.

Previous work on the fungal community associated with the seagrass, Enhalus acoroides, identified a pattern of distance decay, where the fungal community was more similar between seagrass that were closer together geographically than between seagrass that were distant from each other [46]. This suggests that dispersal limitation and/or habitat specialization are playing important roles in structuring the fungal community associated with seagrasses. In this study, we opportunistically sampled fungi associated with a single seagrass species, Z. marina, from a single seagrass patch in Bodega Bay, CA. We did not investigate the fungal community of this seagrass species at other locations and thus, we cannot test for a pattern of distance decay here. However, we do see some examples of habitat specificity/generalism (and/or dispersal efficiency) at a local level in the fungal genera isolated here from Z. marina.

A pattern observed across culture-based studies of seagrass-associated fungi is that ubiquitous fungi are the dominant members of the communities, but that seagrasses also consistently host a diverse set of rare taxa. For example, ubiquitous fungi like Penicillium sp. and Cladosporium sp. have been previously reported as the dominant fungi of leaves in other culture-based studies of Z. marina [36, 38, 94], other seagrass species [26, 34, 35, 41, 95, 96] and freshwater aquatic plants [90]. Additionally, Penicillium sp. and Cladosporium sp. were some of the only fungi in this study which were found to have close relatives associated with different seagrass species. We hypothesize that these fungal genera may be habitat generalists (taxa that occur evenly distributed across a wide range of habitats [97, 98]) in the seagrass (and potentially larger marine) ecosystem as they were isolated from multiple media types, detected from most sample types (Fig 7) and found to have similar mean relative abundances across sample types (S11 Fig). However, just because these fungi are ubiquitous, does not reflect negatively on their potential importance. These habitat generalists have been shown to be highly adaptable with the innate ability to survive in wide range of extreme conditions (e.g. high salinity), are known to perform ecologically important functions (e.g. degradation of organic matter) and represent sources of biologically interesting and active secondary metabolites [33, 94, 99].

We hypothesize that some fungi associated with Z. marina may be habitat specialists (taxa that are more restricted to a specific habitat range [97, 98]). For example, some Colletotrichum sp. may be habitat specialists that preferentially associate with Z. marina leaf tissue. A Colletotrichum sp. ASV (SV10) was found to be dominant on leaves in Ettinger & Eisen [43] and a Colletotrichum spp. isolate was previously reported from another seagrass species as a leaf endophyte [95]. However, in this study, we are unable to decouple the contribution of environmental factors (e.g. habitat or niche specialization) from life history strategies (e.g. dispersal, growth rate). For example, although we hypothesize that some genera may be habitat generalists, it is possible that these patterns may also reflect that some genera have more efficient dispersal mechanisms or faster-growth rates and are able to outcompete slower-growing taxa. We realize these ideas may not be unconnected and that habitat generalists, by their nature, may be assembled by dispersal related mechanisms, and specialists by species sorting [97]. Regardless, future studies could use alternative approaches such as adding different combinations and concentrations of fungicides in order to fully survey rare and slow-growing fungi in this system.

Many of the fungal taxa isolated here are known to have complex life history strategies when associated with land plants. For example, the genus Ramularia includes species that are pathogens of a variety of important agricultural plants including barley and sugar beets [100, 101] and the genus Colletotrichum includes members that can form endophytic or pathogenic associations with land plants [102]. Additionally there is mounting support for a multi-niche view of fungi, with many phyto-pathogens now being found able to form benign or even beneficial endophytic associations with plants [103]. Thus, future research should endeavour to investigate the true functional roles these fungal genera may have when associated with Z. marina and whether these functional roles shift when Z. marina is stressed or challenged.

Although our goal here was to isolate seagrass-associated fungi, we also identified 40 bacterial isolates associated with Z. marina. Since we were using antibiotics, we do not expect these isolates to be representative of the true diversity of the culturable bacterial community associated with Z. marina, just the subset of the community naturally resistant to the antibiotics used here. Most of the bacterial isolates we obtained are from known ubiquitous marine lineages (Vibrio, Pseudoalteromonas, Pseudomonas, Shewanella, and Bacillus) which are likely habitat generalists and have all been previously cultured from Z. marina from Bodega Bay, CA [18, 20, 22–25]. We also isolated several bacterial isolates that may represent rare or slow-growing taxa with interesting ecological implications for the seagrass ecosystem. This includes several Actinomycetes (Streptomyces sp., Rhodococcus sp. and Isoptericola), representatives of which are known to produce a variety of antibiotics and interesting secondary metabolites [104] and members of the genus, Isoptericola, have been previously isolated as endophytes of mangrove plants [105]. We also isolated a Phyllobacterium sp. and representatives of this genus are slow-growing N2-fixing plant-growth promoting bacteria which have been previously isolated from mangrove rhizosphere and the roots of land plants [106-108]. Land plants often overcome nitrogen limitation through beneficial relationships with N2-fixing bacteria and similar associations have been observed between N2-fixing bacteria and Zostera [14, 109–112]. Based on their role as established N2-fixers in other plant systems, it is possible Phyllobacterium sp. may be involved in fixing nitrogen for seagrasses and this possibility should be further investigated.

Just like marine fungi, oomycetes are neglected in marine systems even though they are often implicated as important pathogens of land plants (e.g. Phytophthora ramorum [113]), Phytophthora infestans [114], Pythium spp. [115]). Historically, oomycetes were thought to be fungi based on their similar morphology, but phylogenetic methods have now shown that these organisms are more closely related to diatoms and brown algae [116-119]. In the course of this study, we isolated two members of the Halophytophthora. Representatives of Halophytophthora have been previously isolated associated with Z. marina [47] and this genus includes known saprophytes (organisms living on organic matter) and are thought to be important decomposers in mangrove ecosystems [120]. Recently, Govers et al. [121] suggested that Halophytophthora sp. Zostera may be common in Z. marina beds, and that this oomycete may serve as an opportunistic pathogen by decreasing seed germination in Z. marina populations under certain environmental conditions. More work is needed to understand the possible implications of these oomycetes in the seagrass ecosystem.

Conclusion

Overall, this study generated a fungal culture collection which broadens understanding of the diversity of Z. marina associated fungi and highlights a need for further investigation into the functional and evolutionary roles of fungi and microbial eukaryotes (e.g. oomycetes) associating with seagrasses more generally. We placed this fungal collection in the phylogenetic context of isolates obtained from other seagrass surveys and found that only habitat generalists were isolated in association with multiple species. We then compared the composition of this fungal collection to high throughput sequencing results of the fungal community associated with Z. marina from Ettinger & Eisen [43] and found that taxa isolated here were generally present in the sequencing data, but that they were not prevalent, with the exception of the Glomerellales (e.g. Colletotrichum sp.) on the leaves. Although this study adds to general knowledge of the diversity of Z. marina associated fungi, there are still many unanswered questions to be addressed related to the life history strategies, functional roles, and dispersal mechanisms of marine and seagrass-associated fungi. One of the biggest challenges in marine mycology is assessing whether the fungal taxa observed are actively growing in the marine ecosystem [122]. For our study here we could ask—are many of the proposed habitat generalists actively growing in the seagrass ecosystem or merely passing through as spores? Additionally, many of the fungi cultured here in association with Z. marina have close relatives that are also known to be opportunistic pathogens of land plants. Are these fungi Z. marina pathogens or do they serve some other function in the marine environment? Ultimately this work serves as a necessary first step towards experimental and comparative genomic studies investigating the functional roles of these understudied microorganisms and which may lead to important discoveries related to molecular biology, natural product discovery, fungal diversity and evolution, and global importance of marine fungi.

Distribution of counts of inoculated plates across collection trips used for isolation.

A histogram representing the number of inoculated plates faceted by collection trip (October 2017, May 2018, July 2018, August 2018, January 2019), grouped by inoculum source (leaf, root, rhizome, seawater, sediment) and colored by media recipe used for isolation. The media recipes used included 1% tryptone agar, potato dextrose agar (PDA), potato carrot agar (PCA), palm oil media, lecithin media, malt extract agar (MEA), glucose yeast peptone agar (GYPA), and Zostera marina agar (Zostera). The numbers included on each bar represent the count of plates inoculated for that media recipe with each inoculum source after each collection trip. For example, the first column shows the count of plates from sampling on October 2017 from leaf samples, with three plates on GPYA and two on ME.

(TIF)

Click here for additional data file.

Distribution of counts of inoculated plates across tissue treatments and media types used for isolation.

A histogram representing the number of inoculated plates faceted by media recipe, grouped by inoculum source (leaf, root, rhizome) and colored by tissue treatment (tissue wash, crushed tissue, rinsed whole tissue, bleached whole tissue, surface cleaned whole tissue) used for isolation. The media recipes used included 1% tryptone agar, potato dextrose agar (PDA), potato carrot agar (PCA), palm oil media, lecithin media, malt extract agar (MEA), glucose yeast peptone agar (GYPA), and Zostera marina agar (Zostera). The numbers included on each bar represent the count of plates inoculated for that media recipe with each inoculum source and tissue treatment combination. For example, the first column shows the count of plates inoculated on 1% tryptone from leaves, with eight plates inoculated with rinsed whole leaves.

(TIF)

Click here for additional data file.

Distribution of counts of fungal isolates across media recipes used for isolation.

A histogram representing the number of fungal isolates grouped by order and colored by media recipe used for isolation. The media recipes used included 1% tryptone agar, potato dextrose agar (PDA), potato carrot agar (PCA), palm oil media, lecithin media, malt extract agar (MEA), and glucose yeast peptone agar (GYPA). The numbers included on each bar represent the count of isolates grown on each media recipe.

(TIF)

Click here for additional data file.

Scatterplots showing observed trend between the count of unique media types, salt sources and isolation sources from which a fungal genus was isolated.

Scatter plots representing A) the relationship between the count of unique isolation sources (leaf, root, rhizome, sediment) and the count of unique media types (PDA, palm oil media, lecithin media, MEA), a fungal genus was isolated from (R = 0.86), B) the relationship between the count of unique isolation sources and the count of unique salt sources (no salt, varying amounts of instant ocean [8 g, 16 g, or 32 g], seawater) a fungal genus was isolated from (R = 0.92), and C) the relationship between the count of unique media types and the count of unique salt sources a fungal genus was isolated from (R = 0.87).

(TIF)

Click here for additional data file.

Distribution of counts of fungal isolates across tissue treatments used for isolation.

A histogram representing the number of fungal isolates faceted by inoculum source (leaf, root, rhizome), grouped by order and colored by tissue treatment (tissue wash, crushed tissue, rinsed whole tissue) used for isolation. The numbers included on each bar represent the count of isolates in each Order grown with each inoculum source and treatment combination. For example, the first column shows the count of isolates in the Capnodiales from leaf tissue, with six isolates obtained from crushed leaves, one from leaf washes, and four from rinsed whole leaves.

(TIF)

Click here for additional data file.

Distribution of counts of bacterial isolates across isolation sources and media recipes used for isolation.

Histograms representing the number of bacterial isolates grouped by order and colored by isolation source (A) or media recipe (B). A) When colored by isolation source (leaf, rhizome, root, seawater or sediment), the numbers included on each bar represent the count of isolates obtained from that particular isolation source. B) When colored by media recipe used for isolation (1% tryptone agar, potato dextrose agar [PDA], palm oil media, lecithin media, malt extract agar [MEA], and glucose yeast peptone agar [GYPA]), the numbers included on each bar represent the count of isolates grown on each media recipe.

(TIF)

Click here for additional data file.

A molecular phylogeny of 28S rRNA genes for isolates in the Basidiomycota and Mucoromycota was constructed using Bayesian inference. This alignment was generated using MAFFT (v. 7.402) on the CIPRES Science Gateway web server, trimmed using trimAl (v.1.2) and a phylogenetic tree was inferred on the trimmed alignment with a GTR + I + G model using MrBayes (v. 3.2.2) [75-77,81]. Displayed at each node as a circle in the tree are the Bayesian posterior probabilities (e.g. a black circle represents probabilities greater or equal to 90%, a grey circle represents probabilities greater or equal to 70%, a white circle represents probabilities less than 70%). The names of fungi isolated from Z. marina are shown in green, fungi isolated from other seagrass species are shown in black, and all other fungi are shown in grey. For visualization purposes, selected clades have been collapsed and the number of sequences within that clade is indicated. The GenBank accession numbers of the sequences used to build this phylogeny can be found in Tables 1 and S2–S4.

(TIF)

Click here for additional data file.

A molecular phylogeny of 28S rRNA genes for isolates in the Eurotiomycetes was constructed using Bayesian inference. This alignment was generated using MAFFT (v. 7.402) on the CIPRES Science Gateway web server, trimmed using trimAl (v.1.2) and a phylogenetic tree was inferred on the trimmed alignment with a GTR + I + G model using MrBayes (v. 3.2.2)[75-77,81]. Displayed at each node as a circle in the tree are the Bayesian posterior probabilities (e.g. a black circle represents probabilities greater or equal to 90%, a grey circle represents probabilities greater or equal to 70%, a white circle represents probabilities less than 70%). The names of fungi isolated from Z. marina are shown in green, fungi isolated from other seagrass species are shown in black, and all other fungi are shown in grey. The GenBank accession numbers of the sequences used to build this phylogeny can be found in Tables 1 and S2–S4.

(TIF)

Click here for additional data file.

A molecular phylogeny of 28S rRNA genes for isolates in the Sordariomycetes was constructed using Bayesian inference. This alignment was generated using MAFFT (v. 7.402) on the CIPRES Science Gateway web server, trimmed using trimAl (v.1.2) and a phylogenetic tree was inferred on the trimmed alignment with a GTR + I + G model using MrBayes (v. 3.2.2)[75-77,81]. Displayed at each node as a circle in the tree are the Bayesian posterior probabilities (e.g. a black circle represents probabilities greater or equal to 90%, a grey circle represents probabilities greater or equal to 70%, a white circle represents probabilities less than 70%). The names of fungi isolated from Z. marina are shown in green, fungi isolated from other seagrass species are shown in black, and all other fungi are shown in grey. The GenBank accession numbers of the sequences used to build this phylogeny can be found in Tables 1 and S2–S4.

(TIF)

Click here for additional data file.

A molecular phylogeny of 28S rRNA genes for isolates in the Dothideomycetes was constructed using Bayesian inference. This alignment was generated using MAFFT (v. 7.402) on the CIPRES Science Gateway web server, trimmed using trimAl (v.1.2) and a phylogenetic tree was inferred on the trimmed alignment with a GTR + I + G model using MrBayes (v. 3.2.2)[75-77,81]. Displayed at each node as a circle in the tree are the Bayesian posterior probabilities (e.g. a black circle represents probabilities greater or equal to 90%, a grey circle represents probabilities greater or equal to 70%, a white circle represents probabilities less than 70%). The names of fungi isolated from Z. marina are shown in green, fungi isolated from other seagrass species are shown in black, and all other fungi are shown in grey. The GenBank accession numbers of the sequences used to build this phylogeny can be found in Tables 1 and S2–S4.

(TIF)

Click here for additional data file.

Mean relative abundance of fungal orders isolated in this study across sample types in high throughput sequencing data from Ettinger & Eisen [43].

A histogram representing the mean relative abundance of amplicon sequence variants (ASVs) grouped by order and colored by sample type (leaf, rhizome, root, or sediment). The numbers included on each bar represent the mean relative abundance of the order detected on that particular sample type and only mean relative abundances greater than one percent are shown.

(TIF)

Click here for additional data file.

Collection and media specifications for each microbial isolate associated with Z. marina.

Here we report the specifics of the culture media used to initially grow each isolate including the media recipe used, the salt source and amount, and the inclusion of dehydrated crushed seagrass and of various antibiotics. We also report the collection date of the initial inoculum, whether the inoculum if tissue (e.g. roots, rhizome, leaves) came from tissue washes, crushed tissue or whole tissue, whether the plated inoculum was associated with an individual Z. marina plant from a core or multiple Z. marina plants from a bag, and finally the DNA extraction kit used to extract DNA from each isolate.

(XLSX)

Click here for additional data file.

Fungal sequences used in molecular phylogenies found based on top BLAST matches to Zostera marina associated fungal isolates.

Here we report the GenBank accession number and taxonomic information (Class, Order, Molecular ID) for each fungal 28S rRNA gene sequence obtained based on top BLAST matches to fungal isolates in Table 1 and used here to generate Figs 3–6 and S7–10.

(XLSX)

Click here for additional data file.

Sequences from fungi isolated from seagrasses used in molecular phylogenies.

Here we report information about the fungal 28S rRNA gene sequences used here to generate Figs 3–6 and S7–10 which represent fungal strains previously isolated from seagrasses. We note the seagrass species and tissue material (e.g. leaf, root, matte or rhizomes) the fungus was isolated from, as well as the taxonomic information (Class, Order, Molecular ID, Strain), GenBank accession number and the study of origin for each fungal 28S rRNA gene sequence.

(XLSX)

Click here for additional data file.

Non-seagrass associated fungal isolate sequences from the literature used in molecular phylogenies.

Here we report the GenBank accession number and taxonomic information (Phylum, Class, Order, Species, Strain) for each fungal 28S rRNA gene sequence previously used in James et al [73,74] and used here to generate Figs 3–6 and S7–10.

(XLSX)

Click here for additional data file.

R Markdown file of all data analysis performed in R.

An R Markdown file of the code used to generate the figures in this manuscript.

(PDF)

Click here for additional data file.

27 Apr 2020

PONE-D-20-08398

All small things considered: the diversity of fungi, bacteria and oomycota isolated from the seagrass, Zostera marina

PLOS ONE

Dear Ms Ettinger,

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Reviewer #1: GENERAL COMMENTS

The authors present a culture-dependent study of fungal communities associated with Zostera marina resulting in a culture collection of fungi, some bacteria and a few oomycetes, which contributes to our still emerging understanding of marine fungi within seagrass ecosystems. Besides phenotypical characterization of obtained colonies, this research used sequencing and phylogenetic approaches to assess the reliability of the identities assigned to the different colonies, both within the context of seagrasses in general as well as in comparison with a previous culture-independent study of Z. marina. This original research included a variety of classic microbiology methods, high-standard analyses and conclusions that are supported by the data. However, the following general concerns should be considered for the review process to continue. Please also find below my specific comments and suggestions.

1. The rationale behind using three different markers (i.e. ITS-28S, 16S and 28S) should be explained. Why did the authors amplified these if they were pursuing a fungal collection?

2. Even though the methodological approaches are clearly explained, there are missing details that should be included in a revised version of the manuscript.

3. Identification approaches get somehow mixed in the Results section due to the use of confusing/broad terminology (e.g. “molecular”), or just lack of specification (e.g. “Bacterial isolates were identified as belonging to…”). It is not clear whether the authors are talking about visual identifications, or those made based on taxonomic or phylogenetic analyses.

4. The discussion about patterns of distance decay is very interesting but its link with this study needs further explanation. The authors should consider expanding on the evidence they provide that supports a role of habitat specificity and/or dispersal efficiency at a local level.

5. In general, table formatting should be improved.

SPECIFIC COMMENTS

ABSTRACT:

Lines 60-62: It might be worth it to mention that Halophytophthora may also be just saprophytes.

Line 62: This study did not generate a culture collection of bacteria or oomycetes, as these microorganisms were not intendedly isolated, and therefore (as mentioned later in lines 502-504) are not representative of the true diversity of the culturable bacterial/oomycetal community associated with Z. marina.

INTRODUCTION:

Line 75: The Introduction section may benefit from some background information about bacteria and oomycetes associated with seagrasses to support the discussion about these microorganisms, provided later provided in the manuscript (lines 501-521 and 523-532).

Lines 90: “Blue carbon” is not an example of the preceding sentence, which indeed defines what blue carbon is. I suggest changing “e.g.” for “i.e.”.

Lines 111-113: Did the authors find any unidentified isolates? Based on this sentence, one would expect to see at least a proportion of fungi without taxonomic assignments. What about contrasting identities (i.e. different across identification methods)?

METHODS:

Line 130: The authors state they collected “bulk leaf tissue”. Does this mean that tissue from all leaves from a single seagrass shoot was collected or there was a preference for a certain leaf rank?

Lines 132-133: Is there a reason to keep samples in dark (and then incubate plates in dark too, lines 164-165)?

Line 135: Was there any attempt to cultivate anaerobic fungi, which may play important roles within the anoxic conditions typical of seagrass sediments? There are modifications of the plating technique that could help reduce Oxygen conditions (e.g. pour plate method).

Lines 175-179: An explanation of the criteria used to choose different extraction kits is missing. Why PowerSoil was not used for this particular subset of isolates?

Lines 213-215: Were also the LSU, WARCUP and UNITE datasets used for oomycetes?

RESULTS:

Line 313: “A total of 108 fungal isolates were obtained” … From how many initial samples?

Lines 315-317: “Combined” samples are not mentioned in the Methods section.

Lines 338-341: Confusing sentence. Suggest re-phrasing.

Lines 345-346: An explanation of why this happens is required, since the use of antibiotics is specified in the Methods. The (close) phylogenetic relationship or other similarities between fungi and oomycetes should also be mentioned.

Line 351: Where these bacterial isolates identified by sequencing, phylogenetic analysis or visual characterization of colonies? This should be clearly stated here and in similar sentences throughout the manuscript (e.g. line 360, etc.).

Lines 391-392: Where these two genera rare members of the microbiome?

DISCUSSION:

Lines 414-418: Missing a period in between these two sentences. Change “,” before “Members of the Sordariomycetes…”.

Lines 432-439: This paragraph might benefit from a brief discussion on anaerobic fungi and the limitations of the study in this regard. Please refer to my comment on line 135 below.

Lines 449-451: Explain further. Which results suggest such a role?

Lines 523-524: There are good examples of plant pathogenic oomycetes in the literature.

Lines 528-521: Please correct the reference at the beginning of the sentence (it is missing the year) and delete the one at the end (not necessary).

FIGURES AND TABLES:

Figure 1: The legend should specify which plates correspond to the different taxonomical groups (i.e. bacteria, fungi or oomycetes).

Reviewer #2: This study aimed at characterizing the cultivable mycobiota of Zostera marina from a sampling site in Bodega Bay, California. The authors also report some bacteria that were obtained in spite of the antibiotics used against their growth and two conspecific oomycetes obtained by chance.

Q1: Does the title of your MS fully reflect the merit of your study? Does an a priori biased bacterial spectrum resistant to antibiotics + two accidentally obtained conspecific oomycetes (typically parasites) warrant especially the first part of the title, which reads “All small things considered”? Not least, there are many other “small things” associated with seagrass surfaces (Bryozoa, Cyanobacteria, Foraminifera, microalgae, etc.), so I would suggest choosing a more conservative/informative title for your manuscript.

The authors “opportunistically” collected “individual” plants of this seagrass during “several sampling trips between summer 2017 and summer 2018”. They also collected “bulk leaf tissue” and seawater.

Q2: It seems like this MS represents random results of some (actually, how many?) random (“opportunistically”) collections during one year rather than a targeted study aimed, e.g., at the seasonal effect on the respective mycobiota or the effect of different plant tissues. What is then the reason of publishing these apparently random data in the current form? If the aim was to provide reference material for further studies, i.e., in this case nrDNA sequences, I suggest their deposition in the INSD would do the same job. If the aim was to establish some reference collection of fungi, then these must be first somehow characterized taxonomically (=at the species level) and deposited in a way that they are available to other researchers – even this, however, does not need to be connected with this type of publication. If the aim was to compare the cultivable community with your previous NGS data – then I suggest the authors use at least the same way of sampling (i.e., “at three timepoints 2 weeks apart spanning July to August”), otherwise any such comparisons make little sense to me.

Q3: Considering the sampling protocol, I am not sure whether the authors were actually able to distinguish “individual” Z. marina plants. If yes: why did you collect “bulk leaf tissue”? How many individual plants were actually collected during each sampling, how many in total? These are important data that may help to understand the diversity and abundance of the mycobiota per one such “individual” plant.

For isolation of the mycobionts, the authors used several cultivation media. For obvious reasons this is a good idea, despite that the available seagrass studies indicate only a little effect of this variable. However, it is not easy to evaluate the different media effect in this study because some important information is missing.

Q4: Did you always use the whole range of these media, with the same number of replicates? How many replicates (per seagrass individual, tissue, sediment, seawater, etc.) did you actually establish per sampling?

The authors did not surface-sterilize the investigated tissues (rinsing them in autoclaved water is definitely not enough to get rid of the plethora of surface-dwelling organisms, including their resting stadia). This approach prevents recognition of organisms living inside these tissues (intra- or intercellularly) from the only randomly associated ones.

Q5: Was there any particular reason for omitting surface sterilization? One can ask a simple question, is the mycobiota detected in your study indeed somehow specific to Z. marina or is it just a random spectrum that could be detected +- on any other submerged surface at your sampling site?

The isolation experiment lasted only 4 weeks which is definitely a very short period as many marine fungi are notoriously slow growing (some need many months to start to emerge from the seagrass tissues). I understand it may be difficult to wait one year for the results but your sampling actually spanned over one year so at least the first isolations could have been kept for a longer time (perhaps at least 2-3 months).

The authors obtained 108 fungal isolates but again, some important info is missing to fully understand the meaning of this number and its context.

Q6: How many replicates did not yield any isolate? How many isolates did you obtain in total, how many of them were non-axenic? How many did you attempt to identify, in how many cases you failed (low quality signal, chimeric sequences – perhaps a sign of non-axenic isolates)? Indeed, it seems that not all isolates were eventually sequenced (L. 171: “from the majority of isolates”) so one could ask why?

One of the ways how to distinguish randomly associated surface dwellers and spores from, e.g., true endophytes would be to compare results from the plated tissue segments and the wash liquid (L. 135-147 in Materials and Methods). The latter is, however, not stated in Results.

Q7: Why there are no results mentioned for the wash liquid? It is hard to believe that the wash liquid would have produced no fungi.

Q8: The Result section includes data for “combined leaf and root tissues” and “combined root and rhizome tissues” – what do these “combined” treatments mean/represent? They are not mentioned in Materials and Methods.

Most of the isolates obtained (n=59) belonged to Penicillium sp., which are ubiquitous fungi occurring practically everywhere especially in the form of spores – this is probably the answer to the question the authors asked themselves (L. 550-551): “For our study here we could ask - are many of the proposed habitat generalists actively growing in the seagrass ecosystem or merely passing through as spores?”. Does it make sense to attribute these specifically to Z. marina?

Q9: What does it mean when you state that you did not “attempt to control for effort or isolation frequency in this study”? Please be specific.

There was a little overlap between the fungi isolated here and in other studies and one of the explanations was that this could be “the result of the limits of culture-dependent studies generally or the use of different media recipes and methods for isolating fungi in other studies.”

Q10: What are the “general limits of culture-dependent studies” that could have caused these differences? Is there any “general limit” in you study that differs from other studies? I cannot find any particular...

Q11: Ad different media recipes, here is what you state in L. 311-313: “…we used a variety of media types including several which had been used previously to isolate fungi from seagrasses (e.g. PDA (Sakayaroj et al., 2010; Vohník et al., 2016; Supaphon et al., 2017), GPYA (Panno et al., 2013), MEA (Torta et al., 2015)).”. Did you detect any similarities at least for these media (PDA, GPYA, MEA)? What other methods of isolation did the other studies use? As far as I know they were actually very similar to your methodology…

The authors wrote that “The fungal taxa that did have close relatives that were associated with other species [in other studies] included Penicillium sp….”/“Penicillium sp. and Cladosporium sp. were some of the only fungi in this study which were found to have close relatives associated with different seagrass species”. Indeed, Penicillium is everywhere and for example in culture-based studies of endophytes, these fast growing sporulating isolates are typically discarded as contaminants. At the same time, the authors note that “Penicillium sp., Cladosporium sp. and Ramularia sp. are drivers of the positive relationship observed previously between the number of tissue types and number of media types a fungal genus was isolated from”…

The main result of this study seems to be the establishment of the collection of the 108 fungal isolates “expanding understanding of the diversity of Z. marina associated fungi”. The collection actually comprises ca. 25 different fungal “species”, ca. 7 of these (more than 70 isolates) are ubiquitous sporulating fungi and 11 are represented by only one isolate – I suggest the authors use a more conservative/modest expression than “to expand”.

Minor comments:

Line 48: endophytes are typically defined as symptomless plant endosymbionts and while I acknowledge that they may have secondary beneficial effects on their hosts, they can be to the same extent harmful, e.g., when environmental conditions change, and typically function along the so-called mutualism-parasitism continuum. An easy way how to solve this might be deleting “beneficial” and perhaps also listing mycorrhizal fungi (which are not endophytes, e.g., see Wilson 1995 Oikos 73:274–276)

L. 54: Penicillium sp., etc. – “sp.” not in italics

L. 58-60: Please refrain from the remarks about “interesting secondary metabolites” and “nitrogen cycling in the seagrass ecosystems” – these claims cannot be supported by any results presented in your manuscript.

L. 81: Comments as for L. 48

L. 135: Please state, at least approximately, the time interval between the sample collection and the tissue plating (the length of this interval may significantly affect the obtained symbiont spectra)

L. 136-7: Please detail the rinsing of tissues in autoclaved water – how many washes, for how much time, etc. In case someone wants to compare your protocol with other studies…

L. 140: Perhaps use 1 ml instead

L. 166-7: This implies that some of the obtained isolates were non-axenic; how did you recognize these and how/on what basis could you be “confident” that the final cultures were axenic?

L. 314: Fig. 1 is nice but redundant and since it is practically the same as Fig. 1 in Amend et al. 2019 I suggest omitting it

L. 359: Z. marina not maraina

L. 426: Mediterranean not mediterranean

L. 427-428: “associated with changes in root hair morphology” – not true, please read carefully the paper by Borovec and Vohnik 2018

References must be doublechecked – for example, there is no journal for Rodriguez 2008 (L. 730), L. 788 Kolařík not Kolarˇík, etc.

Table 1: The column “Genus detected in ITS amplicon data” has a very little information value; for example, Penicillium contains more than 300 species with varying biology so detecting “Penicillium” in both data sets tells nearly nothing…

Figure 4: The 45 Penicillium isolates seem to be conspecific – does it make sense to include all the 45 (practically indentical) sequences in the tree? The same holds true for other non-singleton isolates (perhaps better to establish some representative MOTUs?) and also for reference isolates from other studies (the trees are very large – one representative sequence would be enough).

Reviewer #3: The manuscript by Ettinger and Eisen describes a non-targeted cultivation approach to isolate fungi from Zostera marina. The authors identify the strains using standard techniques and compare the data to their previously published ITS amplicon data. As by-catch, some bacteria and Oomycetes were cultured. The manuscript is straightforward albeit without clear directions or hypotheses that are being tested. Large parts of the data presentation (tables and figures) do not have sufficient quality for publication in their present form.

Comments:

1) Tables 1 and 2 are not in a form that is suitable for publication in a paper. I cant event read some of the column headers. Also, how were these calls made? What were the sequence identity cutoffs, etc. What is a ‘Molecular ID’?

2) Tables 2/3: Top BLAST match. I think the authors can do much better to provide useful information on the ID of the bacteria. E.g., sequence similarity to the most closely related described species, not to a genbank entry with potential dubious history

3) Figure 1: While this looks pretty, it is hardly scientific. The authors should indicate the strain names and why this subset of strains is shown here

4) Figure 4,5,6: Isolates that are identical/nearly identical should get clustered

5) Fig. 7: should show which genera were detected by amplicon sequencing or isolation only. Also, again more detail on how these calls were made is necessary. E.g. are the amplicons 100% matches to the isolates? Same applies to the 'yes' or 'no' for detection of the respective 'genera' in the amplicon data

6) Line 908 and Figure S2: It’s one genus, genera is plural…

7) L. 220: analyzed

8) ‘sp.’ like in Penicillium sp. is not in italics as it’s not a species name. Please correct throughout the paper

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Reviewer #1: No

Reviewer #2: No

Reviewer #3: No

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8 Jun 2020

We have addressed all comments in this revision. Please see the uploaded file titled, "Response to Reviewers", for our point by point comments.

Submitted filename: Fungal Collection Response to Reviewers.docx

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30 Jun 2020

Fungi, bacteria and oomycota opportunistically isolated from the seagrass, Zostera marina

PONE-D-20-08398R1

Dear Dr. Ettinger,

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Reviewer #2: All comments have been addressed

Reviewer #3: All comments have been addressed

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Reviewer #3: Yes

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Reviewer #3: Yes

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Reviewer #3: The authors have addressed my concerns appropriately. While the design of the study is not ideal, it does add information to the field.

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6 Jul 2020

PONE-D-20-08398R1

Fungi, bacteria and oomycota opportunistically isolated from the seagrass, Zostera marina

Dear Dr. Ettinger:

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