Northeastern Naturalist 72 T.W. d’Entremont, J.C. López, and A.K. Walker 22001188 NORTHEASTERN NATURALIST V2o5l.( 12)5:,7 N2–o8. 61 Examining Arbuscular Mycorrhizal Fungi in Saltmarsh Hay (Spartina patens) and Smooth Cordgrass (Spartina alterniflora) in the Minas Basin, Nova Scotia Tyler W. d’Entremont1, Juan C. López-Gutiérrez1, and Allison K. Walker1,* Abstract - Saltmarshes are highly productive ecosystems that provide nursery and refuge habitat for animals, buffer storm-wave effects, and stabilize coastlines. Unfortunately, saltmarshes are in decline due to several cumulative stressors. Beneficial root-associated fungi are known to colonize >80% of land plants, but are understudied in intertidal zones. We examined arbuscular mycorrhizal fungi (AMF) in the roots of 2 dominant saltmarsh cordgrasses, Spartina patens (Saltmarsh Hay) and Spartina alterniflora (Smooth Cordgrass) (Poaceae), in the Minas Basin, NS, Canada. We collected 9 sediment cores at the beginning, middle, and end of the 2016 growing season (May–September) for each plant species (n = 54). We examined AMF root colonization using microscopy and fungal-DNA barcoding. Smooth Cordgrass had an AMF root colonization rate of 9%, while Saltmarsh Hay exhibited a higher AMF root colonization rate of 68%. We identified 1 AMF species, Funneliformis geosporum (Glomeraceae), in both host-plant species. We present the first Spartina spp. (cordgrasses) AMF root-colonization data for northeastern North America north of Connecticut, which may aid saltmarsh restoration efforts in Nova Scotia. Introduction Coastal saltmarshes are facing multiple anthropogenic and natural threats including rising sea levels, conversion to waste-disposal areas, and development into agricultural, commercial, or recreational land (Broome et al. 1988). Although some saltmarshes are now protected, many remain vulnerable to toxic spills, dredging, highway construction, and erosion from tidal action and rising sea levels (Broome et al. 1988). These saltmarshes are essential nursery and refuge habitat for juvenile fishes, invertebrates, and birds, which span many trophic levels in both marine and terrestrial food webs (Broome et al. 1988). Saltmarshes also stabilize coastlines, provide a means of storm buffering and nutrient recycling, and are crucial contributors to primary production in marine ecosystems (Broome et al. 1988, Wilson et al. 2015). Restoration of critical saltmarsh habitat has become an interest worldwide due to the threats imposed by current climate-change patterns, especially in areas with extensive coastlines like Nova Scotia (Erwin 2009). Spartina alterniflora (Loisel) (Smooth Cordgrass) and Spartina patens (Aiton) (Saltmarsh Hay) (Poaceae) are the 2 most abundant saltmarsh grasses found along the Atlantic and Gulf coasts of North America (Gessner 1977). These species are found in separate zones within saltmarshes; Smooth Cordgrass grows closer to the 1Department of Biology, Acadia University, Wolfville, NS, Canada B4P 2R6. *Corresponding author - allison.walker@acadiau.ca. Manuscript Editor: Doug Strongman Northeastern Naturalist Vol. 25, No. 1 T.W. d’Entremont, J.C. López, and A.K. Walker 2018 73 tidal interface, due to its ability to oxygenate its roots and rhizosphere in anoxic soils via aerenchyma, and Saltmarsh Hay inhabits the higher areas of the saltmarsh due to lower sediment-salinity levels (Bertness 1991, Daleo et al. 2008, Wilson et al. 2015). This spatial difference is also thought to be the result of the variation in the plants’ ability to compete for nutrients and tolerate the stress of tidal inundation and hypersaline sediment (Daleo et al. 2008). Understanding the factors responsible for the ecological zonation of these 2 species is important for anticipating the effects of tidal action, as well as for improving saltmarsh restoration-project success (Snedden and Steyer 2013). The use of cordgrasses in saltmarsh restoration has been widely applied throughout the US and China due to the species’ extensive root networks that provide infrastructure for saltmarsh sediments (Broome et al. 1988, Hinkle and Mitsch 2005). The ability of both cordgrass species to survive in stressful tidal environments may also be due to symbiotic relationships with arbuscular mycorrhizal fungi (AMF). These fungi colonize the cortical root tissues of most land plants, acting as an extended root network to absorb water and mineral nutrients in exchange for carbohydrates (Cooke and Lefor 1990, Smith and Read 2008). It is currently estimated that 80% of all vascular land plants form symbiotic relationships with AMF (Pirozynski and Malloch 1975, Smith and Read 2008). It is, however, unclear whether some AMF are also limited by the stresses of salinity and submergence experienced in intertidal zones. Existing literature on the halotolerance of AMF is conflicting; some authors suggest that hypersaline soil reduces hyphal growth and root colonization (Giri et al. 2007, Sheng et al. 2008), while others report no reduction in colonization or growth (Yamato et al. 2008). This discrepancy may be due to certain AMF species having higher salinity-tolerances than others (Estrada et al. 2013). It has been observed that AMF increase the salt tolerance of plant species, by allowing them to maintain higher root and shoot biomass than non-mycorrhizal plants in stressful saline environments (Estrada et al. 2013, Giri et al. 2007), although the mechanism remains unknown (Evelin et al. 2009). The effect of submergence on AMF has greater consensus within the mycological community; studies have shown that the conditions created by flooding are unfavorable for AMF (Hildebrandt et al. 2001, Kumar and Ghose 2008). Tidal inundation creates anoxic microenvironments that make it difficult for both fungi and plants to survive, thus reducing species diversity in these areas. Saltmarsh Hay is a highly mycorrhizal species, likely due to lower tolerance of the hypersaline saltmarsh soil (Burcham et al. 2012, Burke et al. 2003, Hoefnagels et al. 1993). In contrast, the AMF-association status of Smooth Cordgrass has been debated. The work of Hoefnagels et al. (1993) and Cooke and Lefor (1990) support that it is a non-mycorrhizal species, but Burcham et al. (2012) demonstrated a weak root-colonization rate of 3%. According to Burcham et al. (2012), the AMF association with Cordgrasses was largely neglected or thought non-existent in early saltmarsh restoration projects. This oversight may have contributed to the failure of many of these projects due to plant loss and soil erosion (Cooke and Lefor 1990). Northeastern Naturalist 74 T.W. d’Entremont, J.C. López, and A.K. Walker 2018 Vol. 25, No. 1 The morphological identification of AMF species is difficult due to the shared spore morphology among most species (Krüger et al. 2009). As a result, the development of molecular techniques targeting AMF species is a promising approach for identifying AMF from field-collected plants. By targeting the internal transcribed spacer (ITS) region of fungal ribosomal DNA (rDNA), Krüger et al. (2009) developed a set of DNA primers capable of amplifying species-specific sequences of AMF, partially spanning the small subunit (SSU), as well as the complete ITS1, 5.8S, ITS2, and partial large subunit (LSU) regions of AMF rDNA. The SSU, 5.8S, and LSU portions are highly conserved, which reduces the number of primers needed in the PCR reactions, while maintaining the ability for the primer cocktails to amplify all possible AMF species (Krüger et al. 2009). Our goals were: (1) to use molecular and microscopic techniques to investigate whether both Smooth Cordgrass and Saltmarsh Hay are colonized by AMF in the Minas Basin, NS, Canada; (2) to increase our understanding of the role this symbiosis may play in plant survival within the highly dynamic, mega-tidal environment of the Bay of Fundy by comparing colonization rates between the 2 species spatially and temporally; and (3) to identify fungi present in cordgrass root samples to increase our understanding of fungal diversity in Nova Scotia saltmarshes. Field-Site Description The site used for this experiment was a well-developed, densely vegetated saltmarsh bordering the Minas Basin near Wolfville, NS, Canada (45°05'42.99"N, 64°21'29.73"W; Fig. 1). This saltmarsh was composed of Smooth Cordgrass near Figure 1. Location of saltmarsh collection site (Wolfville Harbour) in the Minas Basin, NS, Canada. Northeastern Naturalist Vol. 25, No. 1 T.W. d’Entremont, J.C. López, and A.K. Walker 2018 75 the tidal interface transitioning, predominantly, to Saltmarsh Hay in higher saltmarsh elevations. At this site, Smooth Cordgrass was regularly inundated by the semi-diurnal tides, whereas Saltmarsh Hay was only flooded on strong spring tides. This area was classified as a tidal-plain saltmarsh based on the gentle gradient (less than 2%), fine sediment, and 100% cordgrass vegetation cover in the 96-m2 plot. The site is influenced by the mega-tidal regime of the Bay of Fundy, which has a tidal range of 16 m, and results in a stressful environment for the plants present at the site (Keyser et al. 2016, Langley et al. 2013). Methodology Core collection We collected cordgrass roots 3 times during the 2016 growing season from Wolfville Harbour, NS (Fig. 1). Sampling dates were: early (May 27), middle (July 19), and late (September 5). We employed an Eijkelkamp root auger (operational length = 15 cm; diameter = 8 cm; Hoskin Scientific Ltd., Burlington, ON, Canada) to collect cores of cordgrass root tissue and saltmarsh sediment. We sampled along transects to standardize the distance between samples, as well as to provide multiple samples at the same distance from the species interface to provide more reliable results for colonization counts (Fig. 2). We determined the location of the interface by the zonation present between the 2 cordgrass species; we collected no samples within 2 m of this species interface to prevent sampling of the wrong cordgrass Figure 2. Coresampling grid of Saltmarsh Hay (S. patens) and Smooth Cordgrass (S. alterniflora) showing the distances between the samples of each species. The Saltmarsh Hay/Smooth Cordgrass interface is shown by the bold black line; sampling of each cordgrass species was done away from this interface to reduce the risk of accidental multi-species sampling. We conducted sampling at the same site in May, July, and September of 2016. Northeastern Naturalist 76 T.W. d’Entremont, J.C. López, and A.K. Walker 2018 Vol. 25, No. 1 species. We collected 9 sediment cores containing roots from a single species of cordgrass from each zone (3 samples from each transect) during each sampling event, for a total of 27 samples collected per species throughout the growing period (54 samples total). Sample preparation We cleaned the roots on the day of collection by placing samples in a 1-mm sieve and gently washing under cold, running tap water (to prevent damage to fine roots) until no sediment remained. We then placed the cleaned root samples from each core in paper bags, which we labelled and transferred to a drying oven (45 oC) for 24 hr. For each sampling event, we took 40 mg of dried, fine-root tissue from a subset of 5 samples of each cordgrass species and ground it into a fine powder with an autoclaved mortar and pestle prior to DNA extraction. Staining of mycorrhizae and root colonization assessment We employed an ink–vinegar technique modified from Vierheilig et al. (1998) to stain the roots, which we viewed at 400x magnification under a Nikon Alphaphot-2 YS2 compound microscope (Fig. 3). The staining procedure involved: (1) cutting cleaned roots into 5-cm sections and placing them into clean 20-mL scintillation vials, (2) adding enough 10% KOH to each vial to immerse root sections, (3) capping vials and boiling them for 3 min, (4) straining the resultant cleared roots through cheesecloth and rinsing with distilled H2O, (5) transferring cleared roots into new vials and boiling for 3 min in a 5% (v/v) ink/vinegar solution containing Shaeffer® Skrip black ink (Sered, Slovak Republic) and commercial white vinegar (5% acetic acid), (6) straining the resultant roots through cheesecloth, and (7) soaking them in 30 mL of distilled H2O with 3 drops of 5% acetic acid for 20 min. After staining the chitinous cell walls of the AMF, we drew lines 5 mm apart on the reverse of a glass microscope slide with a fine-tipped permanent marker to create points of interest to analyze under a microscope at 400x magnification. We placed stained cordgrass root sections across these lines and covered them with 10% glycerol and a glass cover-slip. For each sample, we analyzed 100 different transects to determine the Figure 3. Arbuscular mycorrhizal fungi stained with Shaeffer Skrip black pen-ink in root tissue from (a) Saltmarsh Hay and (b) Smooth Cordgrass, viewed under 400x magnification. Northeastern Naturalist Vol. 25, No. 1 T.W. d’Entremont, J.C. López, and A.K. Walker 2018 77 percent AMF colonization; we counted points at which AMF-colonized areas of the roots, now stained blue, contacted these lines. We deposited permanent microscope slides in the E.C. Smith Herbarium, Acadia University, Wolfville, NS, Canada (ACAD 043794 [colonized S. alterniflora root], ACAD 043795 [colonized S. patens root]). DNA extraction We employed a Qiagen DNeasy® Plant Mini Kit (Hilden, Germany) and followed its accompanying protocol to extract DNA from the root tissue of fifteen 40-mg samples of each cordgrass species collected throughout the growing season (5 samples from each sampling event selected via random-generator on Microsoft Excel). We doubled the volumes of buffers AP1 and P3 to accommodate the increased amount of starting tissue compared to the recommended 20 mg. Nested polymerase chain reaction (nested PCR) We followed the procedure of Krüger et al. (2009) to conduct PCRs using AMFspecific primer cocktails containing multiple possible primer pairs (Table 1). We electrophoresed a 1% agarose gel at 95 V for 40 minutes and stained it with EtBr to assess amplification success. A Thermo Scientific GeneRuler 10- bp Plus DNA Ladder (Thermo Scientific, Carlsbad, CA) was used as a molecular -size reference. Agarose gel extraction for amplicon purification We used a Qiagen QIAquick® Gel-Extraction Kit by following its provided protocol to gel-extract positive PCR bands present in 1% agarose gels (visualized using a BIO RAD Gel Doc 2000 system). During the gel extractions, the 1% gel previously checked for positive amplification was re-run with an increased running time of 1 h to allow for proper separation of multiple DNA bands. We excised individual DNA bands with a scalpel sterilized with DNA Away and 100% Table 1. Arbuscular mycorrhizal fungi-specific primers used in the nested polymerase chain reactions (Krüger et al 2009). Primer Mix Primer Region Forward (5'–3') Reverse (5'–3') SSUmAf1-2 SSUmAf1 SSU TGGGTAATCTTTTGAAACTTYA SSUmAf2 TGGGTAATCTTRTGAAACTTCA LSUmAr1-4 LSUmAr1 LSU GCTCACACTCAAATCTATCAAA LSUmAr2 GCTCTAACTCAATTCTATCGAT LSUmAr3 TGCTCTTACTCAAATCTATCAAA LSUmAr4 GCTCTTACTCAAACCTATCGA SSUmCf1-3 SSUmCf1 SSU TGCGTCTTCAACGAGGAATC SSUmCf2 TATTGTTCTTCAACGAGGAATC SSUmCf3 TATTGCTCTTNAACGAGGAATC LSUmBr1-5 LSUmBr1 LSU DAACACTCGCATATATGTTAGA LSUmBr2 AACACTCGCACACATGTTAGA LSUmBr3 AACACTCGCATACATGTTAGA LSUmBr4 AAACACTCGCACATATGTTAGA LSUmBr5 AACACTCGCATATATGCTAGA Northeastern Naturalist 78 T.W. d’Entremont, J.C. López, and A.K. Walker 2018 Vol. 25, No. 1 ethanol. The only modifications to the protocol were that we allowed Buffer PE to stand for 5 min in the QIAquick® column before centrifugation to increase DNA yield and used 30 μL instead of 50 μL of final elution Buffer EB to increase the final concentration of the DNA; this solution was left to incubate for 4 min at room temperature to increase DNA yield. PCR of isolated DNA amplicons from gel extraction The initial PCR was done with primer mixtures; thus, the gel-extracted DNA amplicons were subsequently amplified with 1 set of fungal-specific primers prior to Sanger sequencing. The 25-μL PCR reactions contained 12.5 μL of Amresco Ready PCR Mix (2X), 9.5 μL of dd’H2O, 10 pmol of forward (ITS1F) primer, 10 pmol of reverse (ITS4) primer, and 1 μL gel-purified DNA. These primers amplify the ITS1-5.8S-ITS2 region of rDNA, with ITS1F having enhanced specificity for fungi (Gardes and Bruns 1993). We carried out PCR cycling as follows: 95 °C for 3 min, followed by 35 cycles of 95 °C for 1 min, 56 °C for 45 sec, and 72 °C for 1.5 min, and a 10-min final elongation at 72 °C. The length of amplicons obtained were 500–1000 bp. Phylogenetic analysis of DNA sequences We sent DNA amplicons obtained from the final fungal ITS barcode PCRs to the Genome Québec Innovation Centre (McGill University, Montreal, QC, Canada) for Sanger sequencing in the forward and reverse directions. We generated consensus DNA sequences from raw nucleotide reads using MEGA7 (Kumar et al. 2015), locally aligned them to the online nucleotide collection in NCBI’s GenBank using BLAST (Altschul et al. 1990) to identify fungi present in each of the cordgrass root samples, and compiled a species list. We chose a 96% pairwise-similarity threshold to assign identities to the consensus sequences; if this threshold was not met by any of the local alignments generated by BLAST, then we identified that sequence only to genus. We deposited all novel sequences presented herein in GenBank under the accession numbers MF409258–MF409264. Statistical analysis Spatial trends in AMF colonization. We performed a Kruskal–Wallis test to compare the AMF colonization present at each transect in the early, middle, and late growing season, as well as the collective colonization at each transect throughout the entire growth season. All statistical tests were performed at α = 0.05. Temporal trends in AMF colonization. We employed a Kruskal–Wallis test to determine whether a significant difference in AMF colonization rates was present among the early, middle, and late growing-season sampling events. Comparison of AMF colonization between the 2 cordgrass species. We compared AMF root-colonization rates for both cordgrass species using Welch’s t-test to determine whether the AMF colonization rates differed significantly for the 2 species across all sampling events. Northeastern Naturalist Vol. 25, No. 1 T.W. d’Entremont, J.C. López, and A.K. Walker 2018 79 Results AMF colonization of Saltmarsh Hay We examined transects for spatial trends in AMF colonization of Saltmarsh Hay. We conducted one-way ANOVAs on ranks to examine these trends and determine if the AMF-colonization rates at the different transects for Saltmarsh Hay roots were statistically different. There was no statistically significant difference in the AMFcolonization rates for the 2-m, 4-m, or 6-m transects in the early (P = 0.2429; Fig. 4a), middle (P = 0.3643; Fig. 4b), or late (P = 0.2964; Fig. 4c) growing-season transects. When we combined the data for the transects throughout the entire growing season, there was no significant difference in the AMF-colonization of the transects, similar to what was observed previously (P = 0.4496; Fig. 4d). We also analyzed Saltmarsh Hay root AMF-colonization rates to detect temporal variation throughout the growing season and determine whether any trends could be delineated. We detected a statistically significant difference (P = 0.0015) among the early-, middle-, and late-season AMF colonization rates. Multiple comparisons showed that the significant difference was weighted between the middle- and lateseason AMF-colonization rates (P = 0.0011; Fig. 5). AMF colonization of Smooth Cordgrass We examined transect data for spatial trends in AMF colonization of Smooth Cordgrass. We conducted one-way ANOVAs on ranks to examine these trends and Figure 4. Mean ± SEM for arbuscular mycorrhizal fungi colonization of Saltmarsh Hay for each transect in the (a) early (n = 9), (b) middle (n = 9), (c) late (n = 9), and (d) entire growing season (n = 27). Northeastern Naturalist 80 T.W. d’Entremont, J.C. López, and A.K. Walker 2018 Vol. 25, No. 1 determine if the AMF-colonization rates at the different transects were statistically different. There was no statistically significant difference in AMF-colonization rates at the 2-m, 4-m, or 6-m transects in the early (P = 0.9929; Fig. 6a), middle (P = 0.3607; Fig. 6b), or late (P = 0.2786; Fig. 6c) growing-season transects. We detected no significant difference in the combined AMF-colonization rates of any transect, mirroring the results of the sample-period transect analyses (P = 0.3384; Fig. 6d). We also analyzed Smooth Cordgrass root AMF-colonization rates for temporal variation throughout the growing season to determine whether any trends could be Figure 5. Temporal comparison of the AMF colonization of Saltmarsh Hay (mean ± SEM; n = 9). *denotes a statistically significant difference in AMF colonization rates (P = 0.0015). Figure 6. Mean ± SEM for arbuscular mycorrhizal fungi colonization of Smooth Cordgrass for each transect in the (a) early (n = 9), (b) middle (n = 9), (c), late (n = 9), and (d) entire growing season (n = 27). Northeastern Naturalist Vol. 25, No. 1 T.W. d’Entremont, J.C. López, and A.K. Walker 2018 81 delineated. Statistical testing revealed that the early, middle, and late season AMFcolonization rates were statistically different (P = 0.0306). Multiple comparisons showed that this significant difference was weighted between the early and midgrowing- season sampling events (P = 0.0302; Fig. 7). Comparison of AMF colonization of Saltmarsh Hay and Smooth Cordgrass We calculated total AMF-colonization rates for the 2 cordgrass species examined, using all colonization data obtained from the 3 sampling events. Our analysis showed that the total AMF-colonization rates of the 2 cordgrass species were significantly different (P = less than 0.0001) at our study site, with Saltmarsh Hay having a significantly higher colonization rate (68%) than Smooth Cordgrass (9%) (Figs. 5, 7). Phylogenetic analysis One AMF species, Funneliformis geosporum (Glomeraceae), was amplified from both Saltmarsh Hay and Smooth Cordgrass roots. We also identified 5 other fungal species, representing 3 phyla (Table 2). We amplified fungal DNA from 17 of the 30 root DNA samples analyzed, with AMF sequences comprising 18% of all root fungi sequences obtained. Figure 7. Temporal comparison (May– September 2016) of AMF colonization of Smooth Cordgrass (mean ± SEM; n = 9. *denotes a statistically significant difference in AMF colonization rates (P = 0.0302). Table 2. ITS rDNA-sequence identifications of fungi amplified from cordgrass root tissue during this study, with known ecologies. Isolate No. Identification Known ecology 1 Papiliotrema aurea MarineA and terrestrialB yeast found globally 2 Tremella foliacea Commonly found fruiting on branches and decaying wood in Northern temperate regionsC 3 Vishniacozyma carnescens An endophytic yeast found in leaf galls of many different plant speciesD 4 Fusarium acuminatum Pathogen of plants such as Oats and Barley, capable of making T-2 mycotoxin, which is harmful if consumedE 5 Funneliformis geosporum Halotolerant AMF found in saltmarshesF and other types of saline, sodic, and gypsum soilsG 6 Chytridiomycete sp. Morphologically simple aquatic fungi that have a posterior flagellum and reproduce via zoospores H AGao et al. 2007, BTakashima et al. 2003, CRoberts 1999, DGlushakova and Kachalkin 2017, ERabie et al. 1986, FHildebrandt et al. 2001, GLandwehr et al. 2002, HJames et al. 2000. Northeastern Naturalist 82 T.W. d’Entremont, J.C. López, and A.K. Walker 2018 Vol. 25, No. 1 Discussion Spatial trends Statistical analysis of the 3 transects (2 m, 4 m, and 6 m) sampled for each cordgrass species showed no significant spatial differences in AMF colonization for the 3 sampling events in 2016. Many plant species may be more heavily colonized by AMF when experiencing stressful conditions to offset stress-induced losses in plant productivity; AMF species act as an extended root network and provide limiting nutrients to the plant, which can reduce biomass loss (Al-Karaki 2000). We hypothesised that we would find higher AMF colonization rates in locations experiencing more tidal inundation, where plants are most stressed by high salinity levels and may rely more heavily on the AMF to reduce this stress. Temporal trends We observed a temporal trend in AMF-colonization rates. We showed that, in Smooth Cordgrass, the highest AMF-colonization rates occurred in the early growing season, and decreased significantly in the middle and late growing season at our test site. To our knowledge, ours is the first study to investigate temporal changes in AMF-colonization rates in Smooth Cordgrass; some researchers believe it does not form AMF associations (Cooke and Lefor 1990, Hoefnagels et al. 1993). Studies of related cordgrass species show a similar trend, with higher colonization rates occurring during periods of rapid vegetative growth, as is observed in the early growing season. Rapid-growth periods are known to be energetically costly for plants, and increase their need for essential nutrients to build cellular structures (Welsh et al. 2010). Saltmarsh Hay exhibited the opposite trend in AMF colonization, with significantly higher colonization rates occurring in the late growing season, prior to senescence. This result differs from the findings of Welsh et al. (2010) in their similar study on Saltmarsh Hay in Texas, which showed higher AMF-colonization rates during the early growing period, coinciding with periods of rapid growth. Saltmarsh Hay is known to reproduce primarily vegetatively through its rhizomes; thus, the higher AMF-colonization rate during the late growth period may be the result of accumulating AMF for storage prior to the winter dormancy period, allowing it to become more rapidly colonized in the spring once growth resumes. Although this hypothesis must be tested, similar symbiotic behavior has been observed in fungal endophyte species, which colonize plant leaves and excrete defensive compounds and, in turn, can begin decomposing the leaf as soon as it dies (Saikkonen et al. 2010). Cordgrass species comparison When comparing the 2 plant species, the AMF-colonization rates between Saltmarsh Hay and Smooth Cordgrass were significantly different for the populations we sampled at Wolfville Harbor. Saltmarsh Hay had an overall AMF-colonization rate of 68% which corresponds to that of other North American studies conducted on this species (Burcham et al. 2012, Burke et al. 2003, Hoefnagels et al. 1993). The high colonization rate is thought to be caused by an inability to cope with the highly Northeastern Naturalist Vol. 25, No. 1 T.W. d’Entremont, J.C. López, and A.K. Walker 2018 83 saline environment and low soil-nitrogen levels (Burcham et al. 2012, Burke et al. 2003, Hoefnagels et al. 1993, Welsh et al. 2010). Salinity stresses Saltmarsh Hay, and the species relies on tidally imported nitrogen; thus, AMF can increase mineral nutrient and water availability, aiding growth and reducing biomass loss (Welsh et al. 2010). In contrast, Smooth Cordgrass had a lower AMF colonization rate of 9%. This difference may be due to the higher salinity level of lower saltmarsh zones, possibly reducing fungal survival (Pennings et al. 2004). Smooth Cordgrass may also have less reliance on AMF because it is located closer to the tidal interface, and thus, is more frequently inundated by tidal waters, which replenish nitrogen to the sediment (Welsh et al. 2010). The regular flooding of the sediment in which Smooth Cordgrass grows may also decrease the AMF abundance due to sediment anoxia (Kumar and Ghose 2008). Although Smooth Cordgrass had a lower colonization rate than Saltmarsh Hay, it is worth noting that, to the best of our knowledge, our study documents the highest recorded AMF-colonization rate for Smooth Cordgrass; Burcham et al. (2012) reported a 2.4% colonization rate for that species in Louisiana saltmarshes. Fungal identities Analysis of DNA extracted from the roots of the 2 cordgrass test species revealed a diversity of fungi, including 1 AMF species. Fungi from 4 of the main 5 fungal phyla were amplified through PCR, sequenced, and identified through comparison with the reference database GenBank. During the nested PCR, we used the primers developed by Krüger et al. (2009) to discriminate against non-AMF species. Most amplicons from our study were non-AMF, which may be minimized in future studies by adding a surface-sterilization protocol. Krüger et al. (2009) recognized the primer specificity as being problematic against chytridiomycete species, one of which was amplified in our study, but many other genera were also amplified during this experiment, indicating a need for the development of a more selective AMF primer set for environmental studies. One AMF species was amplified from DNA extracted from both cordgrass species sampled in this study: Funneliformis geosporum (T.H. Nicolson & Gerd.) C. Walker & A. Schüßler (previously Glomus geosporum). Funneliformis geosporum has been found globally, with studies indicating that it is one of the most halotolerant AMF species (Hildebrandt et al. 2001, Landwehr et al. 2002). Hildebrandt et al. (2001) reported that up to 80% of AMF spores found in saltmarshes may belong to F. geosporum. Funneliformis geosporum remains an understudied species, but our study shows that it may be a crucial part of the cordgrass rhizosphere in the Wolfville Harbour saltmarsh. Conclusion Our study identified 1 Spartina root-associated AMF species, F. geosporum, which may play a role in the survival of Smooth Cordgrass and Saltmarsh Hay in the Minas Basin, NS, Canada. To our knowledge, we present the first AMF rootcolonization rates for Cordgrass from Atlantic Canada. AMF colonization rates Northeastern Naturalist 84 T.W. d’Entremont, J.C. López, and A.K. Walker 2018 Vol. 25, No. 1 of both Cordgrass species did not differ significantly with respect to their spatial distribution within our study plot, indicating that the difference in salinity levels at each transect may have little influence on AMF colonization. We detected a temporal difference in AMF colonization across the 2016 growing season and the trend was opposite for the 2 cordgrass species, possibly indicating different survival strategies. Our findings demonstrated that Saltmarsh Hay was more reliant on AMF than Smooth Cordgrass, which may be due to decreased tolerance for the saline environment or to less available nitrogen for growth, although we did not test the latter. Our work provides a foundation for further investigating the ecological roles of AMF in the highly dynamic and tide-dominated environments of Atlantic Canada. Such information has the potential to help increase the success of future saltmarsh restoration projects in Atlantic Canada by testing if this identified fungal symbiont can increase cordgrass growth and establishment. Healthy cordgrass plants are crucial contributors to successful saltmarsh restoration efforts because they stabilize saltmarsh sediment and aid in sedimentation Acknowledgments We thank our lab technician Brent Robicheau for assistance with this project. We are grateful to Arthur and Sandra Irving, the Arthur Irving Academy for the Environment, Dr. David Kristie, the Blomidon Naturalists Society, the Acadia University Research Fund, the Acadia University Raddall Research Fund in Biology, and Genome Québec Innovation Centre, Mc- Gill University. Without their generous support, this project would not have been successful. Literature Cited Al-Karaki, G.N. 2000. Growth of mycorrhizal tomato and mineral acquisition under salt stress. Mycorrhiza 10:51–54. Altschul, S.F., W. Gish, W. Miller, E.W. Myers, and D.J. Lipman. 1990. Basic local-alignment search tool. Journal of Molecular Biology 215:403–410. Bertness, M.D. 1991. Zonation of Spartina patens and Spartina alterniflora in a New England saltmarsh. Ecology 72:138–148. Broome, S.W., E.D. Seneca, and W.W. Woodhouse Jr. 1988. Tidal saltmarsh restoration. Aquatic Botany 32:1–22. Burcham, A.K., J.H. Merino, T.C. Michot, and J.A. Nyman. 2012. Arbuscular mycorrhizae occur in common Spartina species. Gulf of Mexico Science 2012:14–19. Burke, D.J., E.P. Hamerlynck, and D. Hahn. 2003. Interactions between the saltmarsh grass Spartina patens, arbuscular mycorrhizal fungi, and sediment bacteria during growing season. Soil Biology and Biochemistry 35:501–511. Cooke, J.C., and M.W. Lefor. 1990. Comparison of vesicular–arbuscular mycorrhizae in plants from disturbed and adjacent undisturbed regions of a coastal saltmarsh in Clinton, Connecticut, USA. Environmental Management 14:131–137. Daleo, P., J. Alberti, A. Canepuccia, M. Escapa, E. Fanjul, B.R. Silliman, M.D. Bertness, and O. Iribarne. 2008. Mycorrhizal fungi determine saltmarsh plant zonation depending on nutrient supply. Journal of Ecology 96:431–437. Erwin, K.L. 2009. Wetlands and global climate change: The role of wetland restoration in a changing world. Wetlands Ecology and Management 17:71–84. Northeastern Naturalist Vol. 25, No. 1 T.W. d’Entremont, J.C. López, and A.K. Walker 2018 85 Estrada, B., R. Aroca, J.M. Barea, and J.M. Ruiz-Lozano. 2013. Native arbuscular mycorrhizal fungi isolated from a saline habitat improved maize antioxidant systems and plant tolerance to salinity. Plant Science 201:42–51. Evelin, H., R. Kapoor, and B. Giri. 2009. Arbuscular mycorrhizal fungi in alleviation of salt stress: A review. Annals of Botany 104:1263–1280. Gao, L., Z. Chi, J. Sheng, X. Ni., and Wang, L. 2007. Single-cell protein production from Jerusalem Artichoke extract by a recently isolated marine yeast Cryptococcus aureus G7a and its nutritive analysis. Applied Microbiology and Biotechnology 77:825–832. Gardes, M., and T.D. Bruns. 1993. ITS primers with enhanced specificity for basidiomycetes: Application to the identification of mycorrhizae and rusts. Molecular Ecology 2:113–118. Gessner, R.V. 1977. Seasonal occurrence and distribution of fungi associated with Spartina alterniflora from a Rhode Island estuary. Mycologia 69:477–491. Giri, B., K. Rupam, and K.G. Mukerji. 2007. Improved tolerance of Acacia nilotica to salt stress by arbuscular mycorrhiza, Glomus fasciculatum, may be partly related to elevated K/Na ratios in root and shoot tissues. Microbial Ecology 54:753–760. Glushakova, A.M., and A.V. Kachalkin. 2017. Endophytic yeasts in leaf galls. Microbiology 86:250–256. Hildebrandt, U., K. Janetta, F. Ouzaid, B. Renne, K. Nawrath, and H. Bothe. 2001. Arbuscular mycorrhizal colonization of halophytes in Central European salt marshes. Mycorrhiza 10:175–183. Hinkle, R.L., and W.J. Mitsch. 2005. Saltmarsh vegetation recovery at salt-hay farm wetland- restoration sites on Delaware Bay. Ecological Engineering 25:240–251. Hoefnagels, M.H., S.W. Broome, and S.R. Shafer. 1993. Vesicular–arbuscular mycorrhizae in saltmarshes in North Carolina. Estuaries 16:851–858. James, T.Y., D. Porter, C.A. Leander, R. Vilgalys, and J.E. Longcore. 2000. Molecular phylogenetics of the Chytridiomycota supports the utility of ultrastructural data in chytrid systematics. Canadian Journal of Botany 78:336–350. Keyser, F.M., J.E. Broome, R.G. Bradford, B. Sanderson, and A. Redden. 2016. Winter presence and temperature-related diel vertical migration of Striped Bass (Morone saxatilis) in an extreme high-flow passage in the inner Bay of Fundy. Canadian Journal of Fisheries and Aquatic Sciences 73:1777–1786. Krüger M., H. Stockinger, C. Krüger, and A. Schüßler. 2009. DNA-based species-level detection of Glomeromycota: One PCR-primer set for all arbuscular mycorrhizal fungi. New Phytologist 183:212–223. Kumar, S., G. Stecher, and K. Tamura. 2015. MEGA7: Molecular Evolutionary Genetics Analysis version 7.0. Molecular Biology and Evolution 33(7):1870–1874. Kumar, T., and M. Ghose. 2008. Status of arbuscular mycorrhizal fungi (AMF) in the Sundarbans of India in relation to tidal inundation and chemical properties of soil. Wetlands Ecology and Management 16:471–483. Landwehr, M., U. Hildebrandt, P. Wilde, K. Nawrath, T. Tóth, B. Biró, and H. Bothe. 2002. The arbuscular mycorrhizal fungus Glomus geosporum in European saline, sodic, and gypsum soils. Mycorrhiza 12:199–211. Langley, J.A., T.J. Mozdzer, K.A. Shepard, S.B. Hagerty, and J.P. Megonigal. 2013. Tidalmarsh plant responses to elevated CO2, nitrogen fertilization, and sea-level rise. Global Change Biology 14:1495–1503. Pennings, S.C., M.B. Grant, and M.D. Bertness. 2004. Plant zonation in low-latitude saltmarshes: Disentangling the roles of flooding, salinity, and competition. Journal of Ecology 93:159–167. Northeastern Naturalist 86 T.W. d’Entremont, J.C. López, and A.K. Walker 2018 Vol. 25, No. 1 Pirozynski, K.A., and D.W. Malloch. 1975. The origin of land plants: A matter of mycotrophism. BioSystems 6:153–164. Rabie, C.J., E.W. Sydenham, P.G. Thiel, A. Lübben, and W.F.O. Marasas. 1986. T-2 toxin production by Fusarium acuminatum isolated from Oats and Barley. Applied and Environmental Microbiology 52:594–596. Roberts, P. 1999. British Tremella species II: T. encephala, T. steidleri, and T. foliacea. Mycologist 13:127–131. Saikkonen, K., S. Saari, and M. Helander. 2010. Defensive mutualism between plants and endophytic fungi? Fungal Diversity 41:101–113. Sheng, M., M. Tang, H. Chen, B. Yang, F. Zhang, and Y. Huang. 2008. Influence of arbuscular mycorrhizae on photosynthesis and water status of Maize plants under salt stress. Mycorrhiza 18:287–296. Smith, S., and D. Read. 2008. Mycorrhizal Symbiosis. Academic Press, San Diego, CA. 800 pp. Snedden, G.A., and G.D. Steyer. 2013. Predictive occurrence-models for coastal-wetland plant communities: Delineating hydrologic-response surfaces with multinomial logistic regression. Estuarine, Coastal, and Shelf Science 118:11–23. Takashima, M., T. Sugita, T. Shinoda, and T. Nakase. 2003. Three new combinations from the Cryptococcus laurentii complex: Cryptococcus aureus, Cryptococcus carnescens, and Cryptococcus peneaus. International Journal of Systematic and Evolutionary Microbiology 53:1187–1194. Vierheilig, H., A.P. Coughlan, U. Wyss, and Y. Piché. 1998. Ink and vinegar, a simple staining technique for arbuscular mycorrhizal fungi. Applied and Environmental Microbiology 64:5004–5007. Welsh, A.K., D.J. Burke, E.P. Hamerlynck, and D. Hahn. 2010. Seasonal analyses of arbuscular mycorrhizae, nitrogen-fixing bacteria, and growth performance of the saltmarsh grass Spartina patens. Plant Soil 330:251–266. Wilson, A.M., T. Evans, W. Moore, C.A. Schutte, S.B. Joye, A.H. Hughes, and J.L. Anderson. 2015. Groundwater controls ecological zonation of saltmarsh macrophytes. Ecology 96:840–849. Yamato, M., S. Ikeda, and K. Iwase. 2008. Community of arbuscular mycorrhizal fungi in coastal vegetation on Okinawa Island and effect of the isolated fungi on growth of Sorghum under salt-treated conditions. Mycorrhiza 18:241–249.