Saltmarsh Sediment Fungal Communities and Arbuscular

Home , Spartina, Spartina patens, Sporobolus alterniflorus

SALTMARSH SEDIMENT FUNGAL COMMUNITIES AND ARBUSCULAR

MYCORRHIZAL FUNGI IN SPOROBOLUS PUMILUS (ROTH) (POACEAE) (SPARTINA

PATENS) OF THE MINAS BASIN, NOVA SCOTIA; IDENTIFICATION, ABUNDANCE

AND ROLE IN RESTORATION

TYLER WADE D’ENTREMONT

BScH Acadia University 2017

Thesis submitted in partial fulfillment of the requirements for the Degree of Master of Science (Biology)

Acadia University Fall Convocation 2019

This thesis by Tyler Wade d'Entremont was defended successfully in an oral examination on 24 June 2019.

The examining committee for the thesis was:

______Dr. Mary Sweatman, Chair

______Dr. Jeremy Lundholm, External Reader

______Dr. Mark Mallory, Internal Reader

______Dr. Allison Walker, Supervisor

______Dr. Juan Carlos López-Gutiérrez, Supervisor

______Dr. Rodger Evans, Head/Director

This thesis is accepted in its present form by the Division of Research and Graduate Studies as satisfying the thesis requirements for the degree Master of Science (Biology).

......

I, Tyler Wade d’Entremont, grant permission to the University Librarian at Acadia University to archive, preserve, reproduce, loan or distribute copies of my thesis in microform, paper, or electronic formats on a non-profit basis. I undertake to submit my thesis, through my University, to Library and Archives Canada and to allow them to archive, preserve, reproduce, convert into any format, and to make available in print or online to the public for non-profit purposes. I, however, retain the copyright in my thesis.

______Author

______Supervisor

______Date

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Table of Contents LIST OF TABLES ...... VI LIST OF FIGURES ...... VII ABSTRACT ...... X ACKNOWLEDGEMENTS ...... XI PRELUDE ...... XII CHAPTER 1: SALTMARSH SEDIMENT FUNGAL COMMUNITIES DIFFER BASED ON SALTMARSH ZONATION AND LOCATION ...... 1 1.1 INTRODUCTION ...... 1

1.1.1 Saltmarsh status ...... 1 1.1.2 Fungi in saltmarshes ...... 2 1.1.3 Environmental DNA ...... 2 1.1.4 Objectives ...... 3 1.2 MATERIALS AND METHODS ...... 4

1.2.1 Study areas ...... 4 1.2.2 Sample collection ...... 5 1.2.3 DNA extraction ...... 6 1.2.4 Analysis of fungal ITS2 metaamplicon sequence data ...... 6 1.3 RESULTS ...... 7

1.3.1 Fungal species composition in Minas Basin, Nova Scotia saltmarshes ...... 7 1.3.2 Species richness among Minas Basin saltmarsh sites and sediment zones ...... 8 1.3.3 Comparison of species composition in saltmarshes of the Minas Basin ...... 9 1.4 DISCUSSION ...... 10

1.4.1 Sediment fungal diversity in saltmarshes of the Minas Basin, Nova Scotia ...... 10 1.4.2 Fungal species richness differs by location and sediment zone ...... 13 1.4.3 Species composition is dependent on the location and sediment collection zone 13 1.4.4 Concluding remarks ...... 14 CHAPTER 2: AMF ARE UBIQUITOUS THROUGHOUT SALTMARSHES OF THE MINAS BASIN, NOVA SCOTIA ...... 15 2.1 INTRODUCTION ...... 15

2.1.1 Saltmarshes ...... 15 2.1.2 Arbuscular mycorrhizal fungi ...... 17 2.1.3 Identification of fungal symbionts ...... 18 2.1.4 Quantification of AMF colonization ...... 19 2.1.5 Objectives ...... 20 2.2 MATERIALS AND METHODS ...... 20

2.2.1 Study Areas ...... 20 2.2.2 Core collection ...... 20 2.2.3 Sample preparation ...... 22 2.2.4 DNA extraction ...... 22 2.2.5 Nested polymerase chain reaction ...... 22 2.2.6 Agarose gel extraction for amplicon purification ...... 23 2.2.7 PCR of isolated DNA amplicons from gel extraction ...... 23 2.2.8 Phylogenetic analysis of DNA sequences ...... 24 2.2.9 Sediment chemical characterization...... 25

2.2.10 Staining of mycorrhizae and root colonization assessment...... 25 2.2.11 Statistical analysis ...... 26 2.3 RESULTS ...... 26

2.3.1 Fungal rDNA analysis...... 26 2.3.2 Sediment chemical characterization ...... 28 2.3.3 Interannual AMF colonization of Sporobolus pumilus roots at the study sites ... 29 2.3.4 Seasonal AMF colonization of Sporobolus pumilus roots ...... 30 2.3.5 Analysis of site-specific AMF colonization ...... 31 2.4 DISCUSSION ...... 32

2.4.1 AMF diversity in Minas Basin, Nova Scotia saltmarshes ...... 32 2.4.2 Saltmarsh sediment chemical characterization at three sites in the Minas Basin, NS 34 2.4.3 AMF colonization by year ...... 35 2.4.4 AMF colonization based on collection period...... 35 2.4.5 AMF colonization differs by collection site ...... 36 2.4.6 Concluding remarks ...... 37 CHAPTER 3: INOCULATION OF RHIZOME DERIVED SPOROBOLUS PUMILUS WITH FUNNELIFORMIS GEOSPORUM MAY BE ESSENTIAL FOR SALTMARSH RESTORATION SUCCESS ...... 39 3.1 INTRODUCTION ...... 39

3.1.1 Restoration efforts ...... 39 3.1.2 Objectives ...... 41 3.2 MATERIALS AND METHODS ...... 41

3.2.1 Seed processing ...... 41 3.2.2 Glomerospore propagation ...... 41 3.2.3 Glomerospore extraction and count ...... 42 3.2.4 Sporobolus pumilus tissue culture initiations ...... 43 3.2.5 Growth trial inoculations ...... 44 3.2.6 Mesocosm growth trial ...... 45 3.2.7 Statistical analysis ...... 47 3.3 RESULTS ...... 48

3.3.1 Simulated saltmarsh trial 1 – rhizome, sterile sand ...... 48 3.3.2 Simulated saltmarsh trial 2 – rhizome, natural sediment ...... 50 3.3.3 Simulated saltmarsh trial 3 – seed, natural sediment ...... 52 3.4 DISCUSSION ...... 54

3.4.1 Growth trial 1 – rhizome, sterile sand ...... 54 3.4.2 Growth trial 2 – rhizome, natural sediment ...... 55 3.4.3 Growth trial 3 – seed, natural sediment ...... 56 3.4.4 Concluding remarks ...... 57 CONCLUSIONS ...... 58 REFERENCES ...... 59

List of Tables

TABLE 1. THE 25 MOST ABUNDANT FUNGAL SPECIES DETECTED AT WOLFVILLE, WINDSOR, AND KINGSPORT,

NOVA SCOTIA SALTMARSHES USING INTERNAL TRANSCRIBED SPACER 2 METABARCODING. SPECIES ARE

ORDERED IN DECREASING ABUNDANCE WITH THEIR DOCUMENTED HABITATS AND PRIMARY GUILDS

REPORTED. PRESENCE = +, ABSENCE = --, AMF = *...... 8

TABLE 2. ARBUSCULAR MYCORRHIZAL FUNGI-SPECIFIC PRIMERS USED IN THE NESTED POLYMERASE CHAIN

REACTIONS (KRÜGER ET AL., 2009)...... 23

TABLE 3. IDENTIFICATIONS USING TYPE SEQUENCES FROM THE NCBI GENBANK DATABASE. SEQUENCES WERE

OBTAINED USING SANGER SEQUENCING AND THE MEGA-BLAST PROGRAM. MULTIPLE IDENTIFICATIONS OF

THE SAME SPECIES WERE LISTED UNDER AN UMBRELLA IDENTIFICATION, ACCESSION NUMBER OF THE BEST

GENBANK MATCH IS REPORTED...... 27

TABLE 4. MEAN VALUES FROM SEDIMENT CHEMICAL CHARACTERIZATION OF THE THREE INVESTIGATED SITES

SURROUNDING THE MINAS BASIN, NOVA SCOTIA (WOLFVILLE, WINDSOR, AND KINGSPORT SALTMARSHES.

(N = 2) ...... 29

List of Figures

FIGURE 1. SPOROBOLUS PUMILUS 2017 SAMPLING SITES LOCATED AT: A) KINGSPORT (45°9'32.42"N,

64°21'36.13"W); B) WOLFVILLE (45°05'42.99"N, 64°21'29.73"W); AND C) WINDSOR, NOVA SCOTIA

(45°0'5.35"N, 64°8'7.13"W)...... 5

FIGURE 2. FUNGAL ALPHA DIVERSITY, AS DETERMINED USING ITS2 METAAMPLICON BARCODING, AT

SALTMARSHES IN KINGSPORT, WINDSOR, AND WOLFVILLE, NOVA SCOTIA SHOWING THE DIFFERENCE IN

SPECIES RICHNESS BETWEEN SITES AND AT DIFFERENT SEDIMENT ZONES, SEPARATED BY ABOVEGROUND

VEGETATION, SPOROBOLUS ALTERNIFLORUS AND SPOROBOLUS PUMILUS...... 9

FIGURE 3. PRINCIPAL COMPONENT ANALYSIS OF SALTMARSH SEDIMENT ITS2 FUNGAL COMMUNITIES SEPARATED

BY COLLECTION SITE (KINGSPORT, WINDSOR, AND WOLFVILLE, NOVA SCOTIA) AND SEDIMENT COLLECTION

ZONE BASED ON THE ABOVEGROUND VEGETATION (SPOROBOLUS ALTERNIFLORUS AND SPOROBOLUS

PUMILUS)...... 10

FIGURE 4. SPOROBOLUS PUMILUS SAMPLING GRID USED AT WOLFVILLE, WINDSOR, AND KINGSPORT, NOVA

SCOTIA TO COLLECT ROOTS, INDICATING THE DISTANCE BETWEEN SAMPLES AND THE LOCATION OF THE

ECOLOGICAL ZONATION. NOTE THAT SPOROBOLUS PUMILUS IS FOUND IN THE HIGH SALTMARSH, WITH

SPOROBOLUS ALTERNIFLORUS BEING CLOSER TO TIDAL WATERS...... 21

FIGURE 5. RELATIVE POSITIONS OF THE FORWARD AND REVERSE AMF-SPECIFIC PRIMERS AND GENERAL FUNGAL

PRIMERS USED FOR PCR OF FUNGAL RDNA EXTRACTED FROM SPOROBOLUS PUMILUS ROOTS (MODIFIED

FROM KRÜGER ET AL., 2009)...... 24

FIGURE 6. NEIGHBOUR-JOINING NETWORK OF THE IDENTIFIED AMF SPECIES FROM THE ROOTS OF SPOROBOLUS

PUMILUS COLLECTED AT WOLFVILLE, WINDSOR, AND KINGSPORT, NOVA SCOTIA. THIS NETWORK USED A P-

DISTANCE MODEL WITH 1000 BOOTSTRAP REPLICATIONS. TYPE SEQUENCES WERE ADDED FROM NCBIS

GENBANK AND CAN BE IDENTIFIED FROM THEIR ACCESSION NUMBER, SEQUENCES DENOTED WITH AN *

WERE PREVIOUSLY IDENTIFIED FROM WOLFVILLE HARBOUR BY D’ENTREMONT ET AL., 2018...... 28

FIGURE 7. MEAN ± SD OF AMF ROOT COLONIZATION AT EACH NOVA SCOTIA SALTMARSH SITE IN 2017 AND

2018. NO SIGNIFICANT DIFFERENCE IN AMF COLONIZATION AT ANY OF THE INVESTIGATED SITES WAS

NOTED BETWEEN YEARS (P = 0.44, Α = 0.05, N = 108)...... 30

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FIGURE 8. POOLED (2017 AND 2018) AMF ROOT COLONIZATION (MEAN ± SD) AT EACH NOVA SCOTIA

SALTMARSH SITE BASED ON COLLECTION PERIOD. WOLFVILLE SHOWED SIGNIFICANTLY HIGHER

COLONIZATION AT LATE COLLECTION (P = 0.003, Α = 0.05), BUT NO SIGNIFICANT DIFFERENCE IN AMF

COLONIZATION WAS DETECTED AT WINDSOR OR KINGSPORT SALTMARSH SITES BETWEEN EARLY AND LATE

COLLECTION (P = 0.84 AND 0.21 RESPECTIVELY, Α = 0.05)...... 31

FIGURE 9. POOLED EARLY AND LATE SPOROBOLUS PUMILUS ROOT AMF COLONIZATION DURING THE 2017 AND

2018 GROWTH SEASON AT THREE MINAS BASIN SALTMARSH SITES IN NOVA SCOTIA. EACH SITE HAD EQUAL

SAMPLE NUMBER (N = 36). * DENOTES A SIGNIFICANT DIFFERENCE IN COLONIZATION (P < 0.001, Α = 0.05).32

FIGURE 10. TIDAL MESOCOSM BENCH USED FOR ALL SIMULATED SALTMARSH GROWTH TRIALS. THE TIDAL

INUNDATION WAS SET USING A 3 HR FLOOD TIDE (PERISTALTIC PUMP) FOLLOWED BY A 3 HR EBB TIDE (DUMP

VALVE) AND A HOLD AT LOW TIDE FOR 6 HR 28 MIN...... 47

FIGURE 11. SURVIVAL OF SPOROBOLUS PUMILUS, DERIVED FROM RHIZOME, UNDER SIMULATED SALTMARSH

CONDITIONS DURING THE 42-DAY MESOCOSM GROWTH TRIAL 1. THE INOCULATED GROUP WAS COLONIZED

BY FUNNELIFORMIS GEOSPORUM (64%) AND THE CONTROL GROUP WAS UNINOCULATED (0%) PRIOR TO

ENTERING THE MESOCOSM BENCH AT DAY 0...... 49

FIGURE 12. SHOOT LENGTH OF SPOROBOLUS PUMILUS GROWTH TRIAL 1 UNDER SIMULATED SALTMARSH

CONDITIONS USING A TIDAL MESOCOSM BENCH. BLUE LINE INDICATES THE AVERAGE SHOOT LENGTH

INCREASE FOR EACH GROUP OVER THE DURATION OF THE 42-DAY TRIAL. INOCULATED GROUP WAS TREATED

WITH FUNNELIFORMIS GEOSPORUM (64% AMF COLONIZATION PRE-TRIAL); CONTROL WAS UNINOCULATED

(0% AMF COLONIZATION PRE-TRIAL)...... 50

FIGURE 13. SURVIVAL OF SPOROBOLUS PUMILUS, DERIVED FROM RHIZOME, UNDER SIMULATED SALTMARSH

CONDITIONS FOR 48 DAYS DURING MESOCOSM GROWTH TRIAL 2. THE INOCULATED PLANTS HAD 17%

COLONIZATION BY FUNNELIFORMIS GEOSPORUM WHILE THE CONTROL ONLY HAD 9% AMF COLONIZATION

PRIOR TO ENTERING THE MESOCOSM BENCH AT DAY 0...... 51

FIGURE 14. SHOOT LENGTH OF SPOROBOLUS PUMILUS GROWTH TRIAL 2 UNDER SIMULATED SALTMARSH

CONDITIONS USING A TIDAL MESOCOSM BENCH. BLUE LINE INDICATES THE AVERAGE SHOOT LENGTH

INCREASE FOR EACH GROUP OVER THE DURATION OF THE 48-DAY TRIAL. INOCULATED GROUP WAS TREATED

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WITH FUNNELIFORMIS GEOSPORUM (17% AMF COLONIZATION PRE-TRIAL); CONTROL WAS GROWN IN

NATURAL SALTMARSH SEDIMENT (9% AMF COLONIZATION PRE-TRIAL)...... 52

FIGURE 15. SURVIVAL OF SPOROBOLUS PUMILUS, DERIVED FROM SEED COLLECTED AT WOLFVILLE HARBOUR,

UNDER SIMULATED SALTMARSH CONDITIONS FOR 28 DAYS DURING MESOCOSM GROWTH TRIAL 3. THE

INOCULATED PLANTS HAD 7% COLONIZATION BY FUNNELIFORMIS GEOSPORUM WHILE THE CONTROL HAD

3% AMF COLONIZATION PRIOR TO ENTERING THE MESOCOSM BENCH AT DAY 0...... 53

FIGURE 16. SHOOT LENGTH OF SPOROBOLUS PUMILUS GROWTH TRIAL 3 UNDER SIMULATED SALTMARSH

CONDITIONS USING A TIDAL MESOCOSM BENCH. BLUE LINE INDICATES THE AVERAGE SHOOT LENGTH

INCREASE FOR EACH GROUP OVER THE DURATION OF THE 28-DAY TRIAL. INOCULATED GROUP WAS TREATED

WITH FUNNELIFORMIS GEOSPORUM (7% AMF COLONIZATION PRE-TRIAL); CONTROL WAS GROWN IN

NATURAL SALTMARSH SEDIMENT (3% AMF COLONIZATION PRE-TRIAL)...... 54

Abstract

Saltmarshes are ecosystems of significant ecological importance for their roles in coastal stability and fundamental roles in marine ecosystems. These ecosystems are in decline due to anthropogenic destruction and current climate regimes causing ocean level rise. Coastal restoration efforts are underway worldwide, but the success remains variable, with many failing in long term coastal stability. The fungal communities within saltmarsh sediment are understudied, especially in areas subject to mega-tidal influence such as the Bay of Fundy, Nova Scotia. My study investigates the fungi present in saltmarsh sediment and focuses on arbuscular mycorrhizal fungi (AMF); symbionts of the dominant saltmarsh grass

Sporobolus pumilus (Roth) (Poaceae) (formerly Spartina patens). Internal transcribed spacer region 2 (ITS2) rDNA metaamplicon barcoding revealed many ubiquitous rhizosphere fungi, including AMF, in three saltmarsh sites around the Minas Basin, NS, but the persistent species composition may be based on the sediment characteristics. AMF colonization counts were conducted on S. pumilus roots and differed between the Kingsport saltmarsh and the

Wolfville and Windsor saltmarshes, although the same Funneliformis species was found in all root tissue. Saltmarsh native AMF, Funneliformis geosporum, was propagated and used as a treatment for S. pumilus in three tidal mesocosm growth trials. This salt tolerant fungal symbiont increased host plant survival and growth under simulated saltmarsh conditions.

These data indicate that fungal saltmarsh community establishment, especially AMF, are an essential consideration for improving saltmarsh restoration practices in areas of mega-tidal influence such as the Bay of Fundy.

Acknowledgements

Over the course of the past two years there have been so many people who have been instrumental to the success of this project. First and foremost, I must thank my wonderful supervisors, Allison Walker and Juan Carlos López-Gutiérrez. These two individuals have become like family to me and without them, I couldn’t have finished this project. They have supported all my decisions over the past years and have encouraged me to think outside the box, which ultimately made this project such a huge success.

I also must thank Arthur, Sandra, and Sarah Irving, for their contributions to this project. Without their generosity, support, the K.C. Irving Environmental Centre, and my

Arthur Irving Academy for the Environment MSc Scholarship, the Atlantic Canadian provinces would still be in the dark about our saltmarsh fungal communities and would be at greater risk of coastal loss. This family has shown incredible dedication to the conservation of both coastal and terrestrial habitats around Atlantic Canada.

I would also like to thank the many others who have played a role in this project:

Glenys Gibson, April Muirhead, Samuel Jean, Robin Browne, Jacob Reicker, Zoë

Migicovsky, Eileen Haskett, John Walker, Russell Easy, David Kristie, Deniz Divanli,

Dalhousie IMR, Genome Quebec, NSERC for supporting the Walker Lab, and finally all the members of the Fungal Dream Team. You have all made valuable contributions to this project and I can’t thank you enough for all you have done!

Prelude

In this thesis I examined the sediment fungal communities in three saltmarshes surrounding the Minas Basin, Nova Scotia using internal transcribed spacer region 2 metaamplicon barcoding (Chapter 1). In Chapter 2, I identified arbuscular mycorrhizal fungi

(AMF) in Sporobolus pumilus roots and determined the strength of the symbiosis using

AMF-specific primers, Sanger sequencing, AMF staining by ink/vinegar, and root colonization counts using compound microscopy. In the final chapter of this thesis (Chapter

3), I propagated native AMF Funneliformis geosporum and applied this inoculant as a treatment to test its ability to increase the survival and growth of S. pumilus under simulated saltmarsh conditions using a tidal mesocosm bench. This thesis provides guidance for future saltmarsh restoration in Atlantic Canada as additional climate stress continues to cause saltmarsh ecosystem loss.

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CHAPTER 1: Saltmarsh sediment fungal communities differ based on saltmarsh zonation and location

1.1 Introduction

1.1.1 Saltmarsh status

Healthy saltmarshes are paramount to the sustainability of intertidal ecosystems due to the large amount of carbon biomass produced and stored in sediment from saltmarsh vegetation (Boesch and Turner, 1984). The term ‘blue carbon’ has been used to refer to the carbon stored in these saltmarsh sinks and is reported to be greater, per unit area, than the carbon sinks of terrestrial forests (McLeod et al., 2011). Unfortunately, despite the huge carbon stores in these ecosystems, we continue to lose them at alarming rates, with minimal efforts targeting their conservation in Atlantic Canada (McLeod et al., 2011).

Saltmarsh ecosystems in Atlantic Canada are dominated by two Sporobolus species in a zonal distribution; Sporobolus alterniflorus (Lawyar-Deslongchamps) (Poaceae), formerly

Spartina alterniflora and commonly known as smooth cordgrass, growing close to the tidal interface, and Sporobolus pumilus (Roth) (Poaceae), formerly Spartina patens and commonly known as saltmeadow cordgrass, found further inland (Bertness, 1991). These Poaceae species must endure multiple stressors such as hypersaline sediment and periodic tidal inundation. Although it is known that these species can excrete a large proportion of the salt they uptake, it is unclear how these stressors affect the essential microbial communities that live in the sediment and help these plants survive. Stressors such as pH, water content, and nutrient loads all influence the assemblages of fungi that can survive in an area due to the unique needs of different fungal species (Kendrick, 2017).

1.1.2 Fungi in saltmarshes

Current knowledge of the fungi associated with these plants is primarily on the arbuscular mycorrhizal symbionts (d’Entremont et al., 2018; Wilde et al., 2009), the primary shoot decomposers of these two species (Buchan et al., 2002; Walker and Campbell, 2010), and the Fusarium pathogens (Elmer and Marra, 2011), with only the former focusing on

Atlantic Canada. Saltmarsh sediment fungal communities have been studied in southeastern

Louisiana, USA, as well, but focused on the rhizosphere shifts after the 2010 Deepwater

Horizon oil spill (Lumibao et al., 2018). To our knowledge, no studies on fungal assemblages in saltmarsh sediments have been undertaken in Atlantic Canada. The fungal diversity in these productive ecosystems may provide insight into better saltmarsh restoration practices throughout Atlantic Canada to combat current habitat degradation and loss.

Fungi play essential roles in practically every ecosystem around the globe, with saltmarshes being no different (Kendrick, 2017). They are essential decomposers in nutrient cycles, symbionts of plants and can even be pathogens (Kendrick, 2017). Saltmarsh dieback is a current looming threat to saltmarsh health and stability and is characterized by thinning of saltmarsh vegetation or areas completely devoid of vegetation (Elmer and Marra, 2011).

Although an outright culprit has not been identified, some suggest that Fusarium spp. may play a role in this detrimental phenomenon (Elmer and Marra, 2011). Fusarium spp. have been identified in saltmarshes of the Minas Basin, NS by d’Entremont et al. (2018), possibly indicating that Minas Basin saltmarshes may be at future risk of dieback.

1.1.3 Environmental DNA

The use of environmental DNA (eDNA) allows for a rapid means of biodiversity assessment using target amplicons that can discriminate between species (Thomsen and

Willerslev, 2015). Metabarcoding approaches allow for entire microbial communities to be sequenced and analyzed together to get a snapshot of the biodiversity present in an area at a given time and allows identification of possibly detrimental or essential microbes to species in the area (Thomsen and Willerslev, 2015). Amplification of the internal transcribed spacer region 2 (ITS2) of fungal rDNA, using ITS86F/ITS4 primers, provides species level identification and is commonly used in metabarcoding of fungi from environmental samples

(Krüger et al., 2009; Op De Beeck et al., 2014). Additionally, metabarcoding approaches decrease the bias of previous morphological studies, as these studies are limited to the species reproducing at the collection time and some fungi are not easily cultured under laboratory settings (Prosser, 2002). Data from eDNA work can be used to assess alpha diversity (species richness within a site) and beta diversity (a comparison of diversity among different sites) (Condit et al., 2002). Using beneficial fungi identified from these studies can help prevent saltmarsh restoration practices from failing similarly to the project initiated near the Indian River, Connecticut (Cooke and Lefor, 1990). This project involved the use of infilled sediment and transplanted Sporobolus, but neglected the use of natural, beneficial microbes. Ultimately, this project is suspected to have failed due to the absence of arbuscular mycorrhizal fungi in the sediment (Cooke and Lefor, 1990).

1.1.4 Objectives

The objectives of this study were to: 1) examine the sediment fungal diversity within three saltmarshes bordering the Minas Basin, Nova Scotia to determine if fungal communities were site dependent; and 2) determine whether zonation of sediment collection can influence the fungal diversity recorded using ITS2 rDNA metabarcoding.

1.2 Materials and Methods

1.2.1 Study areas

The first site used for this study was a well-developed, densely vegetated saltmarsh bordering the Minas Basin near Wolfville, Kings County, Nova Scotia (45°05'42.99"N,

64°21'29.73"W) (Figure 1B). This saltmarsh was dominated by S. alterniflorus at lower tidal areas and S. pumilus in higher saltmarsh elevations. Sparse Distichlis spicata was found throughout the S. pumilus zone. At this site, S. alterniflorus was regularly inundated by the semi-diurnal tides, whereas S. pumilus was only flooded on high spring tides. This area was classified as a tidal plain saltmarsh based on the gentle gradient (<2%), fine sediment and

100% vegetation cover in the 48m2 transect plot. For additional history and characteristics of

Wolfville Harbour saltmarsh see Bleakney and Meyer (1979).

The second saltmarsh site was in Windsor, Kings County, Nova Scotia (45°0'5.35"N,

64°8'7.13"W) (Figure 1C). The slope and vegetation in this saltmarsh was similar to the

Wolfville site, however this saltmarsh was much younger, as it was formed by the construction of a causeway in 1970, that obstructed the Avon River near the study site and extended an already existing saltmarsh. Due to the obstruction, tidal waters slowed, and sediments accumulated by settling forming this new saltmarsh. These sediments are primarily clay, silt, and fine sand that were suspended in the tidal waters, but larger cobbles have also been brought from ice-rafting (see review in van Proosdij and Townsend, 2006).

The third site was near Kingsport, Kings County, Nova Scotia (45°9'32.42"N,

64°21'36.13"W) and was dissimilar to the other two sites in terms of sediment composition and vegetation communities (Figure 1A). The site has a steeper pitch and coarse, sandy sediments (Landry, 2016). This site consists of both S. pumilus and S. alterniflorus, but

unlike the other two sites, S. alterniflorus dominates this ecosystem. For additional history on this site, refer to Landry (2016) and Law et al. (2019). All three sites are influenced by the mega-tidal, cold-water regime of the Bay of Fundy, which has a tidal range of 16m, and results in a highly dynamic environment for all intertidal plants present at the sites (Keyser et al., 2016).

Figure 1. Sporobolus pumilus 2017 sampling sites located at: A) Kingsport (45°9'32.42"N, 64°21'36.13"W); B) Wolfville (45°05'42.99"N, 64°21'29.73"W); and C) Windsor, Nova Scotia (45°0'5.35"N, 64°8'7.13"W). 1.2.2 Sample collection

Sediment cores were collected 31 May and 8 September 2018 from Wolfville,

Windsor, and Kingsport, Nova Scotia in two different sediment zones defined by the aboveground vegetation (Sporobolus pumilus or Sporobolus alterniflorus). Cores were collected using an Eijkelkamp root auger (Hoskin Scientific Ltd.) (operational length = 15 cm; diameter = 8 cm), creating cores of saltmarsh sediment with those dimensions. Sampling for S. pumilus cores was completed in a 48 m2 grid to standardize sampling and provide multiple samples at the same distance from the nearby S. alterniflorus interface. Cores for S.

alterniflorus were collected at 4 m and 8 m from the S. pumilus interface. The interface was determined by the zonation present between the two Sporobolus species. No sampling was conducted within 2m of this species interface to prevent mixing of the two sediment zones.

Nine sediment cores were collected in the S. pumilus zone and two sediment cores were collected in the S. alterniflorus zone at each site for ITS2 rDNA metabarcoding during each sampling date (54 total S. pumilus cores, 12 total S. alterniflorus cores). Cores were stored at

4oC until DNA extraction was conducted.

1.2.3 DNA extraction

DNA was extracted from 200 mg of field collected saltmarsh sediment, collected 5 cm belowground, using a Qiagen DNeasy® PowerSoil Kit (Hilden, Germany) following the manufacturer’s protocol.

1.2.4 Analysis of fungal ITS2 metaamplicon sequence data

DNA from sediment samples was first amplified using ITS86F/ITS4 primer sets to confirm presence of fungal DNA (65/66 were successful). Non-amplified DNA was then sent to the Center for Comparative Genomics and Evolutionary Bioinformatics Integrated

Microbiome Resource (IMR) at Dalhousie University in Halifax, Nova Scotia for ITS2 amplicon sequencing using the Illumina® MiSeq platform. All bioinformatic analyses were conducted using MicrobiomeHelper_amplicon_v0.4 on Oracle VM VirtualBox (Comeau et al., 2017). Initially a metadata file was created for the workflow, which included all parameters of interest. FastQC was then run to manually inspect read quality to ensure it was sufficient and read depth was also checked after trimming off the primer sequences

(Andrews, 2010). The DADA2 workflow was used to stitch sequences and the rare amplicon sequence variants (ASVs) (0.1% of mean sample depth) were filtered out (average sample

depth 17884). QIIME2 (version 2018.6) was used to visualize the data before using R

(version R-3.6.0) create the final alpha (richness) and beta (principal component analysis

(PCA) using unweighted UniFrac distance) diversity plots (R Core Team, 2019). Unweighted

UniFrac distance is a type of PCA used to correlate presence or absence of species to determine how similar sediment communities are based on the species present, not their abundance. Taxonomy was assigned using the UNITE database (Kõljalg et al., 2005), with species level identification and abundance, visualized through QIIME2 view, being the focus of this study.

1.3 Results

1.3.1 Fungal species composition in Minas Basin, Nova Scotia saltmarshes

Metabarcoding taxonomy was assigned using the UNITE database and abundance of species-specific reads were assessed using QIIME2 view. The top 25 species-level identifications were used for the purpose of this study (Table 1). One arbuscular mycorrhizal fungus, Funneliformis geosporum, was found in large abundance at all sites, but reads were also collected for three other AMF species, Claroideoglomus sp., Archaeospora trappei, and

Rhizophagus irregularis, in decreasing abundance.

Table 1. The 25 most abundant fungal species detected at Wolfville, Windsor, and Kingsport, Nova Scotia saltmarshes using internal transcribed spacer 2 metabarcoding. Species are ordered in decreasing abundance with their documented habitats and primary guilds reported. Presence = +, absence = --, AMF = *. Habitat Guild Species Wolfville Windsor Kingsport Marine/terrestrial Saprotrophic Gaeumannomyces graminis + + + Marine/terrestrial Saprotrophic Acremonium breve + + + Terrestrial Saprotrophic Ganoderma lucidum + + + Marine Saprotrophic Halosarpheia japonica + + + Terrestrial Saprotrophic Pholiota adiposa + + -- Marine Saprotrophic Scheffersomyces spartinae + + + Marine Saprotrophic Halomyces littoreus + -- + Marine/terrestrial Mycoparasitic Trichoderma harzianum + + + Terrestrial Saprotrophic Lycoperdon pyriforme + + + Marine Saprotrophic Lignincola laevis + + -- Terrestrial Saprotrophic Coprinopsis atramentaria + + + Terrestrial Saprotrophic Clonostachys rosea + + + Marine/terrestrial Symbiotic Funneliformis geosporum* + + + Marine/terrestrial Saprotrophic Mucor circinelloides -- -- + Terrestrial Saprotrophic Coprinellus micaceus + + + Marine Saprotrophic Phaeosphaeria halima + + -- Marine/terrestrial Saprotrophic Saitozyma podzolica + + -- Marine/terrestrial Mycoparasitic Trichoderma viride + + + Marine/terrestrial Saprotrophic Mortierella alpina + + + Marine Saprotrophic Phaeosphaeria spartinicola + + + Terrestrial Saprotrophic Abortiporus biennis + + -- Terrestrial Saprotrophic Pholiota abieticola + + + Terrestrial Saprotrophic Emericellopsis glabra + + + Terrestrial Saprotrophic Lacrymaria lacrymabunda + + -- Terrestrial Saprotrophic Hericium coralloides -- + +

1.3.2 Species richness among Minas Basin saltmarsh sites and sediment zones

Sediment species richness in Minas Basin saltmarsh sediment were shown to be site specific and dependent on the aboveground vegetation (Figure 2). Alpha diversity was similar between sediment zones at Wolfville and Windsor for sediment collected below

Sporobolus pumilus but was much higher for Windsor for sediment collected below S. alterniflorus. Kingsport had the lowest species richness for both sediment zones compared to

Wolfville and Windsor.

Figure 2. Fungal alpha diversity, as determined using ITS2 metaamplicon barcoding, at saltmarshes in Kingsport, Windsor, and Wolfville, Nova Scotia showing the difference in species richness between sites and at different sediment zones, separated by aboveground vegetation, Sporobolus alterniflorus and Sporobolus pumilus. 1.3.3 Comparison of species composition in saltmarshes of the Minas Basin

Fungal assemblages in Kingsport sediment samples differed from those we documented in Wolfville and Windsor sediment samples (Figure 3), based on our ITS2 metabarcoding data. Wolfville and Windsor saltmarsh species compositions were similar in both sediment collection zones (Figure 3). The PCA plot showed strong separation between

Kingsport and the other sites based on location, but much weaker correlation was present between sediment collection zone (Figure 3). Principal component 2 best differentiated sediment zone, while PC1 best explained location differences.

Figure 3. Principal component analysis of saltmarsh sediment ITS2 fungal communities separated by collection site (Kingsport, Windsor, and Wolfville, Nova Scotia) and sediment collection zone based on the aboveground vegetation (Sporobolus alterniflorus and Sporobolus pumilus).

1.4 Discussion

1.4.1 Sediment fungal diversity in saltmarshes of the Minas Basin, Nova Scotia

The diversity of saltmarsh sediment fungal communities in mega-tidal environments, such as the Bay of Fundy, Canada, have been poorly studied compared to terrestrial soil fungal communities, which are known to provide critical ecosystem services such as decomposition, nutrient cycling and form mutualisms to help plants survive (Dighton, 2016).

These crucial roles form the basis of most food webs and are paramount to nutrient turnover and natural function of these ecosystems (Dighton, 2016).

Our study provides the first sediment fungal eDNA data from Minas Basin, NS saltmarshes. Not surprisingly, many known terrestrial and amphibious fungal species were

found in sediments at all three of our saltmarsh sites. These occurrences are possibly the result of spore deposition from wind, freshwater inputs or ocean currents, and some are unlikely to be metabolically active in these ecosystems, although this remains to be tested.

Ganoderma lucidum and Lycoperdon pyriforme are two examples of terrestrial species which accounted for a high proportion of the reads received through metabarcoding but have no records of being salt-tolerant or inhabiting saltmarsh or marine ecosystems (Freeman and

Ward, 2004; Huss, 1993). Some species such as Mucor circinelloides and Pholiota adiposa were found in high proportion but were not found to be ubiquitous throughout the Minas

Basin, supporting this idea that some of the terrestrial fungi documented in this study may come from neighbouring ecosystems and spore deposition.

Fungi such as Acremonium breve and Halosarpheia japonica have known roles in marine environments (Sterflinger et al., 2001; Abdel-Wahab and Nagahama, 2012).

Acremonium breve has been shown to contribute to biofilms on bryozoans, while H. japonica contributes to the decomposition of woody materials in intertidal ecosystems (Sterflinger et al., 2001; Abdel-Wahab and Nagahama, 2012). Identification of known marine fungi is important for intertidal sediment studies as it shows that the identifications are not due to contamination of the sediment during transport and storage but rather represent fungi that are known to have evolved in the marine environment. Fungal ITS2 metabarcoding is a crucial tool in fungal sediment studies but does not discriminate between species that are active in the area or species that have been brought into the area by an external force (Prosser, 2002).

Additionally, the lysis efficiency of fungal spores and mycelia differ and may alter the true fungal abundance of the sediment (Prosser, 2002).

The sediment used in this study was collected below two different Sporobolus species, Sporobolus pumilus and Sporobolus alterniflorus, the two predominant saltmarsh grass species found along the Minas Basin, NS. Zonation exists between these two species due to competition and tolerance to different levels of tidal inundation (Bertness, 1991; Daleo et al., 2008; Wilson et al., 2015). Phaeosphaeria halima, Phaeosphaeria spartinicola,

Scheffersomyces spartinae, and Funneliformis geosporum all have known co-occurrence with these two Sporobolus species and have been identified in other saltmarsh studies in the USA and Canada (Walker and Campbell, 2010; Filip and Alberts, 1993; Kurtzman et al., 2011; d’Entremont et al., 2018). Phaeosphaeria halima and P. spartinicola have both been isolated from the leaves of S. alterniflorus and play an important role in the decomposition of

Sporobolus litter and recycling nutrients in saltmarsh habitats (Buchan et al., 2002; Walker and Campbell, 2010; Filip and Alberts, 1993). Scheffersomyces spartinae is only known from aquatic environments and it is unknown whether a true interaction between Sporobolus species and S. spartinae exists, or whether S. spartinae occurs due to suspension in the water column; the latter is better supported (Kurtzman et al., 2011).

Funneliformis geosporum forms a mutualistic relationship with both Sporobolus alterniflorus and Sporobolus pumilus in the Minas Basin (d’Entremont et al., 2018).

Funneliformis geosporum is an arbuscular mycorrhizal fungus that colonizes Sporobolus roots in Wolfville, NS, although the strength of the interaction is different for S. pumilus and

S. alterniflorus; the former is more extensively colonized than the latter. Three other undocumented AMF species, from Minas Basin saltmarshes, were also found at low prevalence in this study, Claroideoglomus sp., Archaeospora trappei, and Rhizophagus irregularis. Previous efforts were not able to amplify these AMF from either Sporobolus

species roots (d’Entremont et al., 2018), which may indicate that the do not form a symbiotic relationship with Sporobolus but rather with other saltmarsh plants.

1.4.2 Fungal species richness differs by location and sediment zone

Species richness, as described and measured using alpha diversity, between Wolfville and Windsor was similar for sediment collected below S. pumilus but differed with respect to that of the sediment collected below S. alterniflorus. Windsor had a much higher species richness for this sediment layer which may be the result of ocean current input. Kingsport had the lowest species diversity in both sediment zones compared to the other sites, which may indicate that coarse, sandy sediment at this location is less favourable for intertidal fungi than the clay-rich sediments of Wolfville and Windsor.

For Wolfville and Windsor, the S. alterniflorus sediment zone had higher sediment fungal species richness, which may be due to the frequency of tidal inundation. Kingsport had the opposite trend with the S. pumilus sediment having higher species richness.

Differences in sediment porosity may contribute to this phenomenon but remains untested.

1.4.3 Species composition is dependent on the location and sediment collection zone

Principal component analysis indicated that sediment collected at the Kingsport site contained different fungal communities than the sediments at both Wolfville and Windsor.

This finding indicates that the sediment characteristics at saltmarsh sites may be an important factor in the community makeup. A weaker correlation was found between sediment collection zone indicating that species composition is linked to collection site and aboveground vegetation within a saltmarsh.

Unweighted UniFrac distance was used to assess the differences in community makeup by using presence or absence of a species, without accounting for abundance. As an

exploratory study, we were primarily interested in whether fungal species compositions were similar at our three study locations around the Minas Basin, not fungal abundance.

1.4.4 Concluding remarks

This study provides the first record of saltmarsh sediment fungal diversity in the

Minas Basin, NS and demonstrates that these fungal communities differ based on collection location as well with aboveground vegetation. A large number of terrestrial fungi were also identified in this study, providing an intriguing avenue for future study of ‘amphibious fungi’, in terms of deducing which may be metabolically active in these challenging mega- tidal environments. These data show that marine and terrestrial fungi are present in our valuable coastal saltmarsh ecosystems; some of these species may be essential to their proper function.

CHAPTER 2: AMF are ubiquitous throughout saltmarshes of the Minas Basin, Nova Scotia

2.1 Introduction

2.1.1 Saltmarshes

Tidal saltmarsh vegetation of northeastern North America is primarily composed of halotolerant grasses, such as Sporobolus pumilus (Roth) (Poaceae), formerly Spartina patens, and Sporobolus alterniflorus (Lawyar-Deslongchamps) (Poaceae), formerly Spartina alterniflora, but may also support sedges or other herbaceous angiosperms (Bertness, 1991;

Gessner, 1977). Plants found in these habitats must tolerate high dissolved salt concentrations in sediment as well as regular tidal inundation. Tidal saltmarshes are found in intertidal zones fringing coastlines, primarily surrounding rivers, bays and estuaries that are influenced by tidal regimes (Bertness, 1991; Broome et al., 1988). Saltmarshes vary in size depending on the gradient of the landscape, creating either a narrow skirt of vegetation on steeper coastal terrains or large grasslands when the pitch of the land is subtler (Broome et al., 1988).

Until recently, many of these productive saltmarshes were converted to agricultural or commercial lands due to their fertile soils and prime locations (Broome et al., 1988;

Government of Nova Scotia, 2014). Some of these areas have also been illegal dump sites of biohazardous and household waste (Government of Nova Scotia, 2014). Saltmarsh reclamation was particularly lucrative to the Acadians of Nova Scotia, who constructed dykes to convert saltmarshes to farmland (Department of Natural Resources, 2013). Since the early 1700s, these practices have contributed to the loss of 80% of saltmarshes along the Bay of Fundy and 50% loss across Nova Scotia saltmarsh ecosystems, which provide essential nursery and refuge habitat for juvenile fishes and invertebrates (Broome et al., 1988;

Government of Nova Scotia, 2014). Saltmarshes are also crucial feeding and nesting areas for migratory birds, stabilize coastlines, provide a means of storm buffering, recycle nutrients and are crucial contributors to primary production in marine ecosystems via large carbon stores entering the system through detritivores (Shepard et al., 2011; Wilson et al., 2015).

Although some saltmarshes have recently been protected, others have been lost or remain vulnerable to anthropogenic damage. Construction as well as natural phenomena such as erosion from tidal action and rising sea levels, caused by the warming global climate, contribute to this damage (Shepard et al., 2011). Restoration of critical saltmarsh habitat is underway worldwide, especially in areas where a large proportion of saltmarshes have been lost, such as Nova Scotia (Erwin, 2009; Government of Nova Scotia, 2014). The majority of saltmarsh creation currently underway in Nova Scotia has been from unanticipated establishment from infrastructural development, redistribution of sediments, or conservation efforts by Ducks Unlimited Canada or CB Wetlands & Environmental Specialists (CBWES)

(Bowron et al., 2012). Since 2005, few long-term monitoring programs have been established to assess the success of these restoration projects (Bowron et al., 2012).

Sporobolus pumilus is the most abundant saltmarsh grass found along the Atlantic and Gulf coasts of North America (Gessner, 1977). Sporobolus pumilus is found at higher saltmarsh elevations that are less influenced by tidal regimes and inundation, causing a distinct zonation pattern with a related species Sporobolus alterniflorus, occurring at lower saltmarsh elevations at the tidal interface (Porter et al., 2015). This zonation is dictated by the two species’ abilities to compete for nutrients, as well as tolerate the stresses of tidal inundation by seawater and hypersaline sediments (Daleo et al., 2008). Higher saltmarsh zones are inhabited by S. pumilus because it is thought to be a superior competitor for

nutrients in the less saline environment. Lower saltmarsh habitats are dominated by S. alterniflorus which can oxygenate its roots and the rhizosphere of the anoxic sediment using aerenchyma tissue (Bertness, 1991; Daleo et al., 2008; Wilson et al., 2015). Understanding the factors and interactions responsible for this ecological zonation is important for determining the effects of tidal action, the impacts of anthropogenic change and conservation efforts including restoration (Snedden and Steyer, 2013).

2.1.2 Arbuscular mycorrhizal fungi

The ability of S. pumilus to survive in dynamic tidal environments may be, in part, due to symbiotic relationships with arbuscular mycorrhizal fungi (AMF) (Broome et al.,

1988). AMF colonize the cortical root tissue of many plants and help them establish and survive by providing an extended root network to absorb mineral nutrients and improve plant water status, in exchange for photosynthate from the plant (Cooke and Lefor, 1990; Smith and Read, 2008). New research has shown that these fungi play a fundamental role in the plant microbiome, and current estimates indicate that 80% of all vascular plants form symbiotic relationships with AMF (Parniske, 2008; Smith and Read, 2008). The mutualism between plants and AMF may have been essential for the first species of plants to colonize land, by allowing both parties to gain nutrients from the interaction (Pirozynski and Malloch,

1975). Unfortunately, AMF are understudied in saltmarsh ecosystems and it remains unclear whether AMF species are limited by the stress of these saline environments. Some studies suggest that hypersaline soils and sediments reduce hyphal growth and root colonization by

AMF species (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 a few AMF

species developing higher salinity tolerance over time than others, contributing to the low species diversity that we see in these saline environments (Estrada et al., 2013).

My BScH research was the first to document the presence of AMF in Nova Scotia saltmarsh S. pumilus and S. alterniflorus plants (d’Entremont et al., 2018). We showed root

AMF colonization rates of 68% and 9%, respectively, for these two plant hosts (d’Entremont et al., 2018). Earlier studies discovered S. pumilus mutualisms with mycorrhizal species in

Louisiana and North Carolina, USA, with colonization ranging from 8-52% colonization of its roots (Burcham et al., 2012; Hoefnagels et al., 1993). Colonization may be linked to the inability of S. pumilus to process and excrete the large volume of salt it takes up through transpiration from the saltmarsh sediment (Burke et al., 2003; Hoefnagels et al., 1993;

Burcham et al., 2012). The AMF association with Sporobolus species was largely neglected or thought non-existent in early saltmarsh restoration projects and may have contributed to the failure of many of these projects due to plant loss and subsequent sediment erosion

(Cooke and Lefor, 1990; Burcham et al., 2012).

2.1.3 Identification of fungal symbionts

Prior to molecular techniques, morphological identification of AMF species was difficult due to similar spore morphology among species (Krüger et al., 2009). Since AMF are obligate plant root symbionts and cannot be grown in vitro, identification was based solely on spore morphology, making reliable, species-level, identification nearly impossible.

The development of molecular techniques targeting AMF by using the internal transcribed spacer (ITS) regions of the fungal rDNA, has helped tremendously in accurate identification

(Krüger et al., 2009). The primer cocktails and nested PCR protocol can amplify species- specific barcode sequences, spanning the ITS1 and ITS2 regions of fungal ribosomal DNA.

The ITS1 region is the most highly variable region amplified by the primer set and allows discrimination between different species in conjunction with the slightly more conserved

ITS2 region (Nilsson et al., 2008). Although these primers may amplify a few non-AMF fungal genera, they currently provide our best means of species identification (Krüger et al.,

2009).

Currently, in Nova Scotia, one AMF species has been identified in saltmarsh

Sporobolus roots. We identified Funneliformis geosporum (Glomeraceae) as a symbiont of both S. alterniflorus and S. pumilus roots in a saltmarsh located in Wolfville Harbour, Kings

County, NS (d’Entremont et al., 2018). Funneliformis geosporum has a global distribution, with studies indicating that it is the most halotolerant AMF species known from saltmarsh sediments, as well as sodic and gypsum soils in Europe (Hildebrandt et al., 2001, Landwehr et al., 2002). Eighty percent of AMF spores, by mass, found in saltmarshes can belong to F. geosporum, indicating that it is an important species in these ecosystems (Hildebrandt et al.,

2001). Unfortunately, F. geosporum is understudied and little is known about its role in saltmarshes other than its spore morphology and distribution (Evelin et al., 2009). This species has been documented to naturally colonize species such as Artemisia maritima, Aster tripolium, and Plantago maritima (Landwehr et al., 2002). Funneliformis geosporum may be crucial to the function of saltmarshes and an essential aspect of saltmarsh restoration efforts

(d’Entremont et al., 2018).

2.1.4 Quantification of AMF colonization

Arbuscular mycorrhizal abundance estimates are fundamental to all studies investigating interactions between plants and root endosymbiotic fungi. They tell us the strength of these mutualisms and allow us to estimate the importance of this interaction. One

of the most widely applied methods was proposed by Phillips and Hayman (1970), using

KOH to clear the root tissue and trypan blue to stain the chitinous cell walls of the fungi.

Unfortunately, trypan blue stain is a known carcinogen and teratogen, being linked to birth defects such as encephalocele (Ford and Becker, 1982). Vierheilig et al. (1998) modified the existing technique for staining AMF in plant roots using a 5% v/v ink and vinegar solution to replace the toxic trypan blue as a chitin stain. This method is not only safer, but it also helps to neutralize the basic KOH with an inexpensive acid; household white vinegar.

2.1.5 Objectives

The objectives of this study were to: 1) identify the AMF symbionts of S. pumilus and determine the diversity at each site; 2) identify sediment nutrient differences between three saltmarshes surrounding the Minas Basin, NS; 3) determine whether interannual variation in

AMF colonization exists in S. pumilus at these saltmarsh sites; 4) determine whether seasonal variation in AMF colonization exists in S. pumilus at these saltmarsh sites; and 5) determine whether the AMF colonization of S. pumilus is site specific.

2.2 Materials and Methods

2.2.1 Study Areas

Wolfville, Windsor and Kingsport, Nova Scotia saltmarshes were investigated for this study. Refer to Chapter 1 (1.2.1 Study areas), for comprehensive study site descriptions.

2.2.2 Core collection

Sporobolus pumilus roots were collected early and late in the growing season over a two-year period: on 31 May 2017, 8 September 2017, 15 May 2018, and 8 September 2018 from Wolfville, NS, Windsor, NS, and Kingsport, NS. Cores were collected using an

Eijkelkamp root auger (Hoskin Scientific Ltd.) (operational length = 15 cm; diameter = 8 cm), creating cores of Sporobolus pumilus root tissue and saltmarsh sediment with those dimensions. Sampling was completed in a 48m2 grid to standardize sampling and provide multiple samples at the same distance from the nearby Sporobolus alterniflorus interface to provide more reliable results for colonization counts (Figure 4). The interface was determined by the zonation present between the two Sporobolus species. No sampling was conducted within 2 m of this species interface to prevent sampling of Sporobolus alterniflorus roots. Nine sediment cores containing roots of S. pumilus were collected from each site (3 samples per transect) on each sampling date, with 18 samples collected per site throughout a single growing season (18 samples per year, 36 samples total per site) (protocol modified from d’Entremont et al., 2018).

Figure 4. Sporobolus pumilus sampling grid used at Wolfville, Windsor, and Kingsport, Nova Scotia to collect roots, indicating the distance between samples and the location of the ecological zonation. Note that Sporobolus pumilus is found in the high saltmarsh, with Sporobolus alterniflorus being closer to tidal waters.

2.2.3 Sample preparation

Roots were stored at 4oC within the sediment cores then cleaned as follows: cores were placed in a sieve (mesh size 1mm) and gently washed under cold running tap water (to prevent damage to fine roots) until no sediment remained. Cleaned root samples were then further washed by soaking overnight in dH2O on a rocking table at slow speed to cause agitation. For each sampling event, 100 mg (wet weight) of fine root tissue was taken from a randomly generated subset of 5 samples, from each site, and ground into a fine powder using liquid nitrogen with a sterile autoclaved mortar and pestle prior to DNA extraction. The remaining roots were used for staining and colonization counts.

2.2.4 DNA extraction

DNA was extracted from the root tissue of S. pumilus using a Qiagen DNeasy®

PowerSoil Kit (Hilden, Germany) and following the manufacturer’s protocol.

2.2.5 Nested polymerase chain reaction

PCRs using AMF-specific primer cocktails (containing multiple possible primer sets

[Table 2]) were conducted following the procedure of Krüger et al. (2009). Successful DNA amplification was assessed using a 1% agarose gel electrophoresed at 95 V for 40 min and stained with EtBr. An AXYGEN 100bp DNA Ladder was used as a molecular size reference.

Table 2. Arbuscular mycorrhizal fungi-specific primers used in the nested polymerase chain reactions (Krüger et al., 2009). Primer Primer Region Forward (5’-3’) Reverse (5’-3’) Mix 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 ITS ITS1f SSU CTTGGTCATTTAGAGGAAGTAA ITS4 LSU TCCTCCGCTTATTGATATGC

2.2.6 Agarose gel extraction for amplicon purification

PCR bands present in 1% agarose gels (visualized using a BIO RAD Gel Doc 2000 system) were gel extracted using a Qiagen QIAquick® Gel Extraction Kit following the provided protocol (note that when completing gel extractions, the 1% gel previously checked for positive amplification was re-run with an increased running time of 1 hr to allow for proper separation of multiple DNA bands). Individual DNA bands were excised using a scalpel sterilized with DNAway and 100% ethanol. The only modifications to the protocol were as follows: Buffer PE was left to stand for 5 min in the QIAquick® column before centrifugation to increase DNA yield and 30 µL of final elution Buffer EB was used instead of 50 µL to increase the final DNA concentration (d’Entremont et al., 2018).

2.2.7 PCR of isolated DNA amplicons from gel extraction

As the initial PCR was done with primer mixtures, the gel-extracted DNA amplicons were subsequently amplified with one set of fungal-specific primers prior to Sanger sequencing. The 25 µL PCR reactions contained 12.5 µL of BioRad PCR Master 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. All primers used were at 10 µM concentrations and obtained from

Invitrogen (ThermoFisher). These primers amplify the ITS1-5.8S-ITS2 region of rDNA, with

ITS1f having enhanced specificity for arbuscular mycorrhizal fungi (Gardes and Bruns,

1993). PCR cycling was conducted as follows: 95°C for 3 min, followed by 35 cycles of 95°C for 1 min, 56°C for 45 secs, and 72°C for 1.5 min, and a 10 min final elongation at 72oC.

Amplicons obtained were 500 –1000 bp long (Figure 5) (d’Entremont et al., 2018).

Figure 5. Relative positions of the forward and reverse AMF-specific primers and general fungal primers used for PCR of fungal rDNA extracted from Sporobolus pumilus roots (modified from Krüger et al., 2009).

2.2.8 Phylogenetic analysis of DNA sequences

DNA amplicons obtained from the final fungal ITS barcode PCRs were sent for

Sanger sequencing in the forward and reverse directions at the Genome Québec Innovation

Centre (McGill University, Montreal). Trimmed, consensus DNA sequences were locally aligned to the online nucleotide collection in NCBI’s GenBank using BLAST (Altschul et al.,

1990) to identify fungi present in each of the Sporobolus root samples. A list of species was compiled and a Glomeromycota neighbour-joining tree (p-distance model) was created. A

97% pairwise similarity threshold was used to assign species identities to the sequences; if this threshold was not met by any of the local alignments generated by BLAST, then a sequence was identified to genus only.

2.2.9 Sediment chemical characterization

Two additional sediment cores were collected at each sampling site (Wolfville,

Windsor, and Kingsport) on 15 May 2018 for nutrient analysis by the Nova Scotia

Department of Agriculture, Harlow Institute (Truro, NS). Cores were collected using the same methods as root cores (Section 2.2.2).

2.2.10 Staining of mycorrhizae and root colonization assessment

AMF within collected roots were stained using an ink-vinegar technique modified from Vierheilig et al. (1998) and viewed under 400X magnification using a Nikon

Alphaphot-2 YS2 compound microscope (Figure 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 fully immerse root sections, (3) capping vials and boiling them for 3 min in a hot water bath, (4) straining the resultant cleared roots using cheesecloth then rinsing with distilled H2O, (5) transferring cleared roots into new 20 mL vials and boiling for 3 min in a 5% (v/v) ink/vinegar solution containing Shaeffer® Skrip black ink (Sered, Slovak Republic) and white vinegar (5% acetic acid), (6) straining roots using cheesecloth, and (7) allowing roots to soak in 30 mL of distilled H2O with 3 drops of

5% acetic acid for 20 min to remove stain from the root tissue. After staining the chitinous cell walls of the AMF, lines were drawn 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 the compound microscope at 400X magnification. Stained S. pumilus root sections were placed across these lines and covered, with 10% glycerol and a glass cover slip. For each sample,

100 separate slide transects were analysed to determine the percent AMF colonization (where

AMF-colonized areas of the roots, now stained blue, contacted these lines) (Giovanetti and

Mosse, 1980).

2.2.11 Statistical analysis

A generalized linear model with binomial distribution determined whether 1) AMF colonization was dependent on the year the roots were collected at each location (interannual

AMF colonization) or 2) collection site (Wolfville, Windsor, or Kingsport) caused significant differences in the average AMF colonization (α = 0.05). A Welch’s two-sample t-test was used to determine whether seasonal differences in AMF colonization exist at each saltmarsh site using R software (version R-3.6.0) (R Core Team, 2019).

2.3 Results

2.3.1 Fungal rDNA analysis

Sporobolus pumilus roots were subject to DNA extraction and subsequently targeted

PCR to identify AMF symbionts. One AMF species was identified to genus using Sanger sequencing (Table 3), but a species level identification was not obtained. Three non-AMF species were also amplified and were identified as possible (Table 3).

Table 3. Identifications using type sequences from the NCBI GenBank database. Sequences were obtained using Sanger sequencing and the mega-blast program. Multiple identifications of the same species were listed under an umbrella identification, accession number of the best GenBank match is reported. Sample Identification Ident Accession Literature Reference 3467 Lobulomycetales sp. 97% EF432819 A chytrid fungal order proposed 3468 in 2009, it encompasses species 3472 such as Clydaea vesicula, a species formerly identified in saltmarshes of the Minas Basin, NS (Simmons et al., 2009; d’Entremont et al., 2018) 3469 Funneliformis sp. 95% JQ048769 Species in this AMF genus often 3477 have funnel-shaped spores that are 3478 found in clusters of 1 to 20 spores 3480 (Schüßler and Walker, 2010). 3481 Funneliformis geosporum has 3482 previously been identified to form 3484 symbioses with Sporobolus species in saltmarshes of Wolfville Harbour, NS, as well as saline sites in Germany and Hungary (d’Entremont et al., 2018; Hildebrandt et al., 2001; Landwehr et al., 2002). 3471 Tremellaceae sp. 94% NR159742 A family with cosmopolitan distribution, species may be anamorphic or teleomorphic, the former primarily being yeasts (Liu et al., 2007). Species have a variety of roles including human pathogens. 3476 Naumovozyma sp. 90% HE580273 Ascomycete yeast belonging to the Saccharomyces sensu lato group (Andersson and Cohn, 2017)

Multiple AMF sequences were obtained from Sanger sequencing and were used, along with AMF type material sequences from GenBank, to construct a neighbour-joining network using the p-distance model to assess phylogenetic relationships with related species

(Figure 6).

Figure 6. Neighbour-joining network of the identified AMF species from the roots of Sporobolus pumilus collected at Wolfville, Windsor, and Kingsport, Nova Scotia. This network used a p-distance model with 1000 bootstrap replications. Type sequences were added from NCBIs GenBank and can be identified from their accession number, sequences denoted with an * were previously identified from Wolfville Harbour by d’Entremont et al., 2018.

2.3.2 Sediment chemical characterization

Sediment chemical characterization was performed for each site (Table 4). Wolfville and Windsor sites had similar nutrient levels, but the Kingsport site had much lower levels of most nutrients.

Table 4. Mean values from sediment chemical characterization of the three investigated sites surrounding the Minas Basin, Nova Scotia (Wolfville, Windsor, and Kingsport saltmarshes. (n = 2) Parameter Wolfville, NS Windsor, NS Kingsport, NS Organic Matter (%) 3.75 2.95 1.00 pH 6.76 6.77 6.78

P2O5 (kg/ha) 136 126 57

K2O (kg/ha) 1398 1310 285 Sodium (kg/ha) 11198 8715 2003 Calcium (kg/ha) 1409 1333 199 Magnesium (kg/ha) 2223 2070 372 Copper (ppm) 1.66 2.32 0.83

2.3.3 Interannual AMF colonization of Sporobolus pumilus roots at the study sites

Arbuscular mycorrhizal colonization was analyzed at Wolfville, Windsor, and

Kingsport saltmarshes during the 2017 and 2018 Sporobolus pumilus growing season. The

AMF colonization rates were not significantly different within a single site over the two-year study period (p = 0.44, α = 0.05, n = 108) (Figure 7).

Figure 7. Mean ± SD of AMF root colonization at each Nova Scotia saltmarsh site in 2017 and 2018. No significant difference in AMF colonization at any of the investigated sites was noted between years (p = 0.44, α = 0.05, n = 108).

2.3.4 Seasonal AMF colonization of Sporobolus pumilus roots

AMF colonization of S. pumilus was significantly higher in Wolfville saltmarsh, at late collection in the growing season (p = 0.003, α = 0.05) (Figure 8). There was no significant difference in AMF colonization between early and late season collection at either the Windsor or Kingsport saltmarshes (p = 0.84 and 0.21 respectively, α = 0.05) (Figure 8).

Figure 8. Pooled (2017 and 2018) AMF root colonization (mean ± SD) at each Nova Scotia saltmarsh site based on collection period. Wolfville showed significantly higher colonization at late collection (p = 0.003, α = 0.05), but no significant difference in AMF colonization was detected at Windsor or Kingsport saltmarsh sites between early and late collection (p = 0.84 and 0.21 respectively, α = 0.05). 2.3.5 Analysis of site-specific AMF colonization

Measurements of percent root colonization of Sporobolus pumilus by AMF were conducted at three sites around the Minas Basin, NS. Although AMF colonization did not significantly differ over the 2-year study period for Wolfville (70%) and Windsor (67%), the

Kingsport site had a significantly lower colonization (41%) than the other two sites (Figure

9). Increased branching of AMF hyphae, outside of the roots, were noted for the Kingsport saltmarsh samples, although colonization within the root was lower.

Figure 9. Pooled early and late Sporobolus pumilus root AMF colonization during the 2017 and 2018 growth season at three Minas Basin saltmarsh sites in Nova Scotia. Each site had equal sample number (n = 36). * denotes a significant difference in colonization (p < 0.001, α = 0.05).

2.4 Discussion

2.4.1 AMF diversity in Minas Basin, Nova Scotia saltmarshes

Arbuscular mycorrhizal diversity in Minas Basin, NS saltmarshes has only been investigated by one other study, which documented a single species, Funneliformis geosporum, present in a saltmarsh in Wolfville, NS (d’Entremont et al., 2018). This present study improved the methods of the former study to verify whether the results were representative of other saltmarsh sites surrounding the Minas Basin, NS.

Funneliformis sp. was the only AMF species found in S. pumilus roots in this study, but it was found in high abundance at Wolfville, Windsor, and Kingsport, NS. Funneliformis geosporum is likely the species found within the S. pumilus roots, based on the previous study by d’Entremont et al. (2018) and the identification of F. geosporum from eDNA in

Chapter 1 of this study. Unfortunately, a species level verification could not be obtained from the Sanger sequence data due to noisy chromatograms and mismatching of nucleotides in the consensus alignment, possibly caused by partial DNA degradation or genetic variation within the population. Many AMF individuals can colonize the same root, which can lead to messy data as single amplicons cannot be separated and sequenced.

Non-target fungal sequences were also obtained in this study, similar to the former studies conducted in the Minas Basin, NS (d’Entremont et al., 2018). Improved washing techniques of the root tissue did significantly reduce the number of non-target amplicons, but due to the broad range of the Krüger et al. (2009) primer cocktails used in this study, not all non-target fungi could be excluded. These primers are known to amplify some non-AMF species but remain the most stringent primer set to broadly amplify AMF sequences. These non-target amplicons can also provide insight into the biological processes occurring in saltmarsh ecosystems. Lobulomycetales, identified in this study, have previously been identified from marine habitats, and have been shown to parasitize diatoms, possibly acting as an alternative algal substitute at the base of these food webs (Hassett et al., 2017).

Sequence variability obtained from Sanger sequencing was high for F. geosporum, possibly due to DNA degradation or genetic variation within the population. Samples were left at room temperature during washing and, due to the nature of AMF DNA work, a total of

3 polymerase chain reactions were used as well as a gel extraction, which requires brief exposure to UV light which is a powerful mutagen (Rastogi et al., 2010). Even with the use of consensus sequences, high level identifications were not obtained due to sequence dissimilarity to type material on NCBIs GenBank, possibly indicating that the Funneliformis

sp. found in saltmarshes along the Minas Basin, NS, could be a new species closely related to

F. geosporum.

The sequences obtained were used to construct a neighbor-joining network including reference sequences from NCBI GenBank. There were no site-specific groupings of

Funneliformis sp. sequences from this study, indicating that the strains of Funneliformis at

Wolfville, Windsor and Kingsport are similar but can vary genetically within a single saltmarsh. Arbuscular mycorrhizal fungi are known to be exceptionally variable in the ITS region compared to other fungi, and do not resolve well with closely related species

(Stockinger et al., 2010).

2.4.2 Saltmarsh sediment chemical characterization at three sites in the Minas Basin, NS

Saltmarsh sediment chemical analysis at Wolfville, Windsor, and Kingsport, NS was conducted to determine whether these sites differed in the available nutrients for S. pumilus growth. Visual examination of the three sites indicated that the sediment at the Kingsport site was sandy and coarse compared to Wolfville and Windsor which was primarily composed of fine silt and clay. Wolfville and Windsor showed similar nutrient loads for each of the nutrients tested in this study, yet only showed similarity to Kingsport with respect to pH level. It is worth noting that this study did not test all nutrients required for plant growth, but instead used field soil tests provided by the Department of Agriculture, Harlow Institute,

Truro, NS, and sample size was low (2 pooled samples per site).

Kingsport was lower in all tested nutrients compared to the other two sites, most notably potassium and phosphorus, even though all three sites are influenced by the same tidal nutrient input. In pedology, it is generally accepted that coarser soils have a reduced

ability to hold nutrients, including water, than finer soils (Eash et al., 2016). The increased surface area of particles such as clay, as found at the Wolfville and Windsor saltmarshes, allow more nutrients such as calcium, magnesium, phosphorus, and water to be retained in the sediment, increasing total nutrient content and increasing the availability to plants (Eash et al., 2016).

2.4.3 AMF colonization by year

A comparison of the colonization counts at each site over the two-year sampling period showed that there was no significant change in colonization rates over the sampling period at any of the sites. There was however a marginal increase in AMF colonization in

2018, although the cause of this increase was not determined.

AMF colonization rates of S. pumilus were similar to that reported in a previous study conducted at the Wolfville, NS saltmarsh; d’Entremont et al. (2018) reported a 68% colonization rate, for the same plant host in the growing season of 2016, compared to our numbers for 2017 and 2018; 68% and 72% respectively. Since AMF communities tend to stabilize over time, it can be inferred that each of these sites have well developed AMF communities and colonization is not likely to deviate from the presented rates under normal environmental conditions (Smith and Read, 2008). These data also indicated that this symbiosis is persistent in the saltmarsh environments and may be crucial for S. pumilus to survive under these conditions (d’Entremont et al., 2018).

2.4.4 AMF colonization based on collection period

AMF colonization, based on collection period, was not shown to be significantly different for two of the three saltmarshes, Windsor and Kingsport. Sporobolus pumilus collected from Wolfville were shown to have significantly higher AMF colonization if

collected toward the end of the growth period. This trend has previously been documented by d’Entremont et al. (2018), who attributed it to an accumulation of AMF in the plant tissue prior to the winter dormancy period.

2.4.5 AMF colonization differs by collection site

Over our sampling period, there was no significant difference in AMF colonization between the S. pumilus communities at Wolfville and Windsor, NS. These sites share sediment characteristics and are inundated by the same tidal waters of the Minas Basin. The resemblance between these sites in terms of sediment chemical composition and inundation patterns suggests that sediment characteristics may be an important driver of AMF community compositions in saltmarsh sediments and may affect the strength of belowground plant-fungal interactions, as has been found in previous terrestrial studies (Alguacil et al.,

2016).

The Kingsport, NS site had significantly lower colonization than the former sites, even though the sediment contained a higher abundance of sand. Sediments highly composed of sand have been shown to increase plant root colonization by AMF, such as Funneliformis geosporum (previously identified in the Minas Basin, NS) (d’Entremont et al., 2018; Martin et al., 2012). Nutrients measured such as sodium, calcium, manganese, copper, P2O5, and

K2O were also much lower in the sediments collected at Kingsport, NS compared with the other investigated sites.

Phosphorus is an important nutrient for plant growth and is known to be one of the valuable nutrients AMF provide plants (Smith and Read, 2008). With lower phosphorus recorded at Kingsport, NS, logic would suggest that AMF colonization would be higher at this site, but the opposite trend was observed. This may be due to increased branching of the

Sporobolus pumilus root networks at this location, which were recorded by observation while conducting this study. These extensive root networks occur due to unstable substrate levels

(larger root networks increase anchorage of plants) and decreased growth-limiting bulk densities of the sand particles (Daddow and Warrington, 1983). Smaller particles such as clay, found in abundance at Wolfville and Windsor, have much higher growth-limiting bulk densities and, once compacted, essentially stop root growth due to resistance (Daddow and

Warrington, 1983). Increased AMF colonization may be used to accommodate this smaller root network, as arbuscular mycorrhizal hyphae are much smaller in diameter than the plant roots and can grow in the small pores between clay particles more easily.

2.4.6 Concluding remarks

We showed that the AMF diversity found within Sporobolus pumilus roots in three saltmarshes bordering the Minas Basin, NS is limited to a single species, likely

Funneliformis geosporum, and there doesn’t appear to be significant difference in the DNA sequences obtained from each site. This important plant root-microbial mutualism, understudied in tidal wetlands, was present over the two-year study period for Windsor and

Kingsport, NS, saltmarshes and over a three-year period for Wolfville, NS saltmarsh.

Although Wolfville and Windsor saltmarsh Sporobolus pumilus had similar AMF colonization rates for this study, Kingsport S. pumilus plants had significantly lower colonization, possibly due to the different sediment type (high sand and low nutrients) present at the site. This study provides a baseline for AMF consideration in saltmarsh restoration projects surrounding the Minas Basin, Nova Scotia.

CHAPTER 3: Inoculation of rhizome derived Sporobolus pumilus with Funneliformis geosporum may be essential for saltmarsh restoration success 3.1 Introduction

3.1.1 Restoration efforts

Sporobolus (formerly Spartina) species have been widely used in Asia, Europe and

North America in saltmarsh restoration to increase bird and fisheries habitat, provide pasture for livestock, reclaim lost saltmarshes, as well as to stabilize coastlines from erosion

(Broome et al., 1988; Cooke and Lefor, 1990; Warren et al., 2002). Sporobolus is ideal for this due to its extensive root networks that provide infrastructure and stability in saltmarsh sediments (Broome et al., 1988). More specifically, Sporobolus pumilus is one of the most commonly used species in saltmarsh restoration, due to increased root branching through far- reaching rhizome growth (Shepard et al., 2011). Unfortunately, S. pumilus has a low seed germination rate, making seedling transplants the method of choice to restore these ecosystems (Cooke and Lefor, 1990). Additionally, little is known about seed predators of this species, which may further decrease the chance of S. pumilus survival via seed stock. In the Bay of Fundy in particular, the mega-tidal dynamics and ice scouring, from harsh winters in Nova Scotia, further add to these challenges. Collectively these variables make efforts of saltmarsh restoration more difficult than other areas which have already shown difficulty in providing long term functioning and conservation of the saltmarsh ecosystems (Callaway,

2005).

One saltmarsh restoration project in the late 1970s along the Indian River in Clinton,

Connecticut, provides an example of a restoration failure (Cooke and Lefor, 1990). After removing contaminated sediment from the Indian River site, it was replaced with clean

sediment fill and planted with a variety of species including S. alterniflorus and S. pumilus

(Cooke and Lefor, 1990). This site was then left to re-establish on its own, via natural sexual and asexual propagation. A re-examination of the site 10 years after the restoration revealed that the restored areas were still largely devoid of vegetation, even though transplants had been used to establish the restored area. Upon analysis of tissue from both native and restored

S. pumilus it was concluded that the restored plants were not forming symbioses with AMF, which restricted establishment and growth (Cooke and Lefor, 1990). The inability to re- establish the mutualism between S. pumilus and AMF is thought to be due to the absence of this native fungus in the infilled sediment, which our project works to remedy. On a local scale, saltmarsh restoration practices have been successful after dyke breaches and culvert expansion, with 100% vegetation cover (primarily Sporobolus alterniflorus) 3 years post breach, but these projects still neglect local sediment AMF (van Proosdij et al., 2010,

Bowron et al., 2011).

Funneliformis geosporum is an AMF species previously identified from a saltmarsh in Wolfville, NS and is thought to be of critical importance to S. pumilus (d’Entremont et al.,

2018). Funneliformis geosporum has been found globally; studies indicate it is one of the most halotolerant AMF species (Hildebrandt et al., 2001; Landwehr et al., 2002).

Funneliformis geosporum may also make up 80% of AMF spores found in saltmarshes

(Hildebrandt et al., 2001). Currently, F. geosporum remains an understudied AMF, but likely plays a crucial role in the Sporobolus root endophytic community in the saltmarshes of

Wolfville Harbour (d’Entremont et al., 2018). Use of this fungus in simulated saltmarsh studies may reveal the importance of this species to saltmarsh ecosystem structure. Methods such as the use of tidal mesocosms, as simulated saltmarshes, allow control of environmental

parameters, such as light intensity and tidal inundation, while still producing seminatural conditions (Callaway et al., 1997). Mesocosms are useful tools for testing factors and restoration techniques and prior to implementation, offering great potential for evaluation

(Callaway et al., 1997).

3.1.2 Objectives

The objectives of this study were to: 1) identify whether inoculation of S. pumilus by

F. geosporum caused increased survival and growth, compared to a sterile control, under simulated saltmarsh conditions (tidal mesocosm); 2) determine if F. geosporum inoculant outperformed the use of natural sediment in improving survival and growth of S. pumilus under simulated saltmarsh conditions; and 3) determine whether AMF inoculated rhizome or seed derived S. pumilus survived better under simulated saltmarsh conditions.

3.2 Materials and Methods

3.2.1 Seed processing

Processing of S. pumilus seed was conducted at the end of the growth season (20

September 2017). Stalks with seeds were collected, from the Wolfville saltmarsh site and placed in brown paper bags. Seeds were processed after the stalks of the plants were dried (7 days at room temperature) to make it easier to remove the surrounding chaff. Final storage was at 4oC in the seed collection at the E.C. Smith Herbarium, Wolfville, NS.

3.2.2 Glomerospore propagation

Trap pots were created in the K.C. Irving Environmental Science Centre greenhouse

(Wolfville, N.S.), using sediment collected under S. pumilus (from the Wolfville Harbour saltmarsh, NS study site), mixed with autoclaved sand in a 1:1 ratio in 2 L pots (Morton et

al., 1993). Untreated corn seeds (Zea mays) (Vesey Seeds) were planted in the trap pots, and were lightly watered, using ground water, every second day, to create water stress (no additional nutrients were added to the soil). Temperature fluctuated between 22-25oC under a

14-hr lighting regime (approx. 500 µmm-2s-1). Stressed conditions allowed AMF spores and mycelia present in the sediment and in the S. pumilus root fragments to colonize the corn roots to alleviate some of the plant nutrient and water stress.

Once mature (8 weeks), the plants were sacrificed by cutting the stem 1 cm above the soil. The soil was then left to dry for 1 week, which allowed the AMF, colonizing the corn roots, to sporulate. Quality of this resulting inoculant was assessed by observing AMF colonization of host plant roots (refer to method described in previous chapter) and AMF spore counts of inoculant.

3.2.3 Glomerospore extraction and count

AMF spores (from primarily Funneliformis geosporum), propagated in the trap pots, were subject to a spore extraction procedure using sieving and a glucose centrifugation technique (Morton et al., 1993). First, 100 g of sediment and root fragments were broken up in a beaker, with a glass stir rod and water, to release the spores from the fungal hyphae. The material was then run through four sieves of 500 µm, 250 µm, 150 µm, and 45 µm mesh size respectively. Cleaned material collected in the bottom two sieves was then added to a centrifugation tube with dH2O and centrifuged at 1750 × g in an Eppendorf centrifuge 5804R for 1 min. The supernatant was then run through a 45 µm sieve then transferred to another centrifuge tube and suspended in a 20/60% glucose gradient. The suspended material was then centrifuged for 3 min at 1000 × g. The supernatant was then transferred to a glass Petri dish and rinsed with tap water to remove all glucose. Spores were manually counted to

determine the spore density using a dissecting microscope at 100X magnification. Final spore concentration was 8 F. geosporum spores per gram of inoculant.

3.2.4 Sporobolus pumilus tissue culture initiations

3.2.4.1 From rhizome fragments

Rhizome fragments of Sporobolus pumilus, collected from the Wolfville Harbour saltmarsh, were placed in mesh tea balls (Kintech International, mesh size <500µm) then wrapped with parafilm, around the seal, to ensure that the balls did not open during the sterilization procedure. The balls were rinsed with reverse osmosis (RO) water, then placed under running tap water for 1 hour. Mesh balls were then transferred to 50% isopropanol for

30 seconds, then immediately rinsed with RO water. Mesh balls were then transferred to pre- sterilized Magenta vessels with forceps inside a laminar flow bench. Twenty percent bleach with two drops of Tween 80 was then added in sufficient quantity to cover the mesh balls. The vessels were agitated for 10 min, then poured off and rinsed 4 times with RO water inside the flow bench. The rhizome fragments were then transferred to media containing 10 ml/L Plant Preservative Mixture (PPM) + ½ Murashige and Skoog (MS) + Ampicillin (A) plates for 3 days, then 1.0 ml/L PPM + ½ MS + A plates for 1-2 weeks. Fragments were then moved to ½ MS + A medium in 25 × 90 mm glass vials, and later to Magenta vessels (77 mm length × 77 mm width × 97 mm height).

3.2.4.2 From seed

Seeds of Sporobolus pumilus, previously collected from the Wolfville Harbour saltmarsh

(20 September 2017) and stored in the seed collection at the E.C. Smith Herbarium, were placed on moist paper towel for 24 hr prior to sterilization. Seeds were sterilized using the same technique as rhizome fragments except the seeds were agitated in 20% bleach with 2

drops of Tween 80 for 15 min instead of 10 min and were transferred to ½ MS + A in 100 ×

15 Petri dish after the 10 ml/L PPM + ½ MS + A treatment.

3.2.5 Growth trial inoculations

Sterile S. pumilus were grown in tissue culture on ½ MS + A medium in Magenta vessels (from rhizome pieces) or Petri dishes (from seed). Sterile rhizome pieces were sectioned multiple times to control the growth of the plants to ensure they would not be too large for the growth trials. Seeds were grown on Petri dishes for 10 days prior to exposure to the fungal inoculant. It was deemed appropriate using a power of t-test, that 30 plants per test group would be sufficient, per group, to detect differences between the populations (delta =

0.73).

3.2.5.1 Trial 1 – rhizome, sterile sand

A group of 48 rhizome derived S. pumilus plants were removed from ½ MS + A medium and planted in a 1:1 mixture of autoclaved sand and Funneliformis geosporum inoculant (8 spores per gram), in a potting tray (26 cm width × 52 cm length × 7 cm deep)

(16 April 2018). Similarly, 48 rhizome derived S. pumilus were planted in autoclaved sand to act as a control. Plants were grown for 21 days under greenhouse conditions with a 14-hour lighting regime, daily moisture monitoring and temperature control between 18-24oC. AMF colonization was assessed using the root staining and microscopy methods described in

Chapter 2, prior to plant introduction to the tidal mesocosm.

3.2.5.2 Trial 2 – rhizome, natural sediment

Groups of 42 rhizome derived S. pumilus were used for both the test and control due to material availability for this trial. The test group was inoculated using a 1:1 mixture of F. geosporum inoculant (8 spores per gram) and natural saltmarsh sediment, collected from the

Wolfville Harbour saltmarsh site (1 June 2018). The control group used a 1:1 mixture of sterile, autoclaved sand and natural saltmarsh sediment. Plants were grown for 21 days under greenhouse conditions with a 14-hour lighting regime, daily moisture monitoring and temperature control between 18-24oC. AMF colonization was assessed using the root staining and microscopy methods described in Chapter 2, prior to plant introduction to the tidal mesocosm.

3.2.5.3 Trial 3 – seed, natural sediment

This trial used seed derived S. pumilus, collected from Wolfville Harbour and reared in tissue culture. Each treatment contained 30 plants and were grown either using a 1:1 mixture of F. geosporum inoculant (8 spores per gram) and natural saltmarsh sediment, collected from the Wolfville Harbour saltmarsh site (test) or 1:1 mixture of sterile, autoclaved sand and natural saltmarsh sediment (control) (8 October 2018). Plants were grown for 21 days under greenhouse conditions with a 14-hour lighting regime, daily moisture monitoring and temperature control between 18-24oC. AMF colonization was assessed using the root staining and microscopy methods described in Chapter 2, prior to plant introduction to the tidal mesocosm.

3.2.6 Mesocosm growth trial

1200 L of seawater was obtained from the Minas Basin, NS, which borders all sites investigated in this study, and was diluted to a salinity range of 18-22 ppt using groundwater.

The experiment was performed in a tidal mesocosm bench using a natural tidal cycle, to match the Minas Basin, and create natural inundation of the plants by tidal waters. The mesocosm bench was custom designed by Groupe DHB Inc. for the K.C. Irving

Environmental Science Centre (Wolfville, NS). The stainless-steel bench was 181 cm wide,

312 cm long, and 45 cm deep (Figure 10). The program consisted of a 3 hr flood tide followed by a 3 hr ebb tide and a hold at low tide for 6 hr 28 min, to advance the tide each cycle. Tides were simulated using a peristaltic pump, for high tide, or a dump valve for low tide.

Natural saltmarsh sediment (collected from Wolfville Harbour) was placed in custom

1.5 cm thick plexiglass pots (10 cm diameter, 10 cm height). The bottom of the pipe was sealed with Nitex (75 µm) mesh, and a plastic grill to facilitate water exchange without losing sediment (Haskett, 2018). Plexiglass racks were built to suspend the pots 6 cm above the bottom of the bench, with the maximum inundation of the cores (at high tide) covering the cores (< 1 cm).

A 14-hour photoperiod was used to increase the light exposure to approximately 500

µmm-2s-1. Salinity was monitored daily using an Oakton Salt 6+ handheld salinity probe.

Shoot length and survival of the plants was monitored weekly throughout the 2018 growing season until one treatment had reached approximately 50% mortality. At the end of each trial, all plants were sacrificed for wet weight, dry weight, and shoot length to determine whether the effects of AMF inoculation on the growth of the S. pumilus plants were statistically significant.

Figure 10. Tidal mesocosm bench used for all simulated saltmarsh growth trials. The tidal inundation was set using a 3 hr flood tide (peristaltic pump) followed by a 3 hr ebb tide (dump valve) and a hold at low tide for 6 hr 28 min. 3.2.7 Statistical analysis

3.2.7.1 Pre-growth trial

A Welch’s two-sample t-test (α = 0.05) was used to compare shoot length of S. pumilus in the two treatment groups after the 21-day treatment period prior to entering the tidal mesocosm bench.

3.2.7.2 Post-growth trial

A Welch’s two-sample t-test (α = 0.05) was used to compare the final shoot length of the two S. pumilus treatment groups after the duration of the growth trial in the tidal mesocosm bench.

3.2.7.3 Growth rate

The final shoot length, post-growth trial, was subtracted from the initial shoot length, after the treatment, for each sample to determine the average shoot length increase for each group. A Welch’s two-sample t-test (α = 0.05) was then used to compare the inoculated and control group growth rate to determine if the rate of growth over the trial duration was significantly different.

3.3 Results

3.3.1 Simulated saltmarsh trial 1 – rhizome, sterile sand

At the end of the 21-day treatment period, the inoculated S. pumilus had 64% root colonization by F. geosporum, while the control had 0% AMF colonization. Growth trial 1 was run for 42 days with the F. geosporum inoculated group showing higher survival under the simulated saltmarsh conditions (85% survival compared to 44%) (Figure 11).

Figure 11. Survival of Sporobolus pumilus, derived from rhizome, under simulated saltmarsh conditions during the 42-day mesocosm growth trial 1. The inoculated group was colonized by Funneliformis geosporum (64%) and the control group was uninoculated (0%) prior to entering the mesocosm bench at day 0.

Shoot length of AMF inoculated S. pumilus plants was significantly higher than that of control plants both immediately after the 21-day treatment period (p < 0.001, α = 0.05) and overall after the 42-day growth trial (p < 0.001, α = 0.05) (Figure 12). The shoot length increase over the 42-day growth trial was also significantly greater for the inoculated group compared to the control (p = 0.034, α = 0.05) (Figure 12).

Figure 12. Shoot length of Sporobolus pumilus growth trial 1 under simulated saltmarsh conditions using a tidal mesocosm bench. Blue line indicates the average shoot length increase for each group over the duration of the 42-day trial. Inoculated group was treated with Funneliformis geosporum (64% AMF colonization pre-trial); control was uninoculated (0% AMF colonization pre-trial). 3.3.2 Simulated saltmarsh trial 2 – rhizome, natural sediment

At the end of the 21-day treatment period, the inoculated S. pumilus had 17% root colonization by F. geosporum, while the control had 9% AMF colonization. Growth trial 2 was run for 48 days with the F. geosporum inoculated group showing higher survival under the simulated saltmarsh conditions than the lower colonized control (90% survival compared to 57%) (Figure 13).

Figure 13. Survival of Sporobolus pumilus, derived from rhizome, under simulated saltmarsh conditions for 48 days during mesocosm growth trial 2. The inoculated plants had 17% colonization by Funneliformis geosporum while the control only had 9% AMF colonization prior to entering the mesocosm bench at day 0.

Shoot length of AMF inoculated S. pumilus was significantly higher than that of control plants both immediately after the 21-day treatment period (p = 0.004, α = 0.05) and overall after the 48-day growth trial (p < 0.001, α = 0.05) (Figure 14). The shoot length increase over the 48-day trial was not significantly different for this trial between the inoculated and control group (p = 0.60, α = 0.05) (Figure 14).

Figure 14. Shoot length of Sporobolus pumilus growth trial 2 under simulated saltmarsh conditions using a tidal mesocosm bench. Blue line indicates the average shoot length increase for each group over the duration of the 48-day trial. Inoculated group was treated with Funneliformis geosporum (17% AMF colonization pre-trial); control was grown in natural saltmarsh sediment (9% AMF colonization pre-trial).

3.3.3 Simulated saltmarsh trial 3 – seed, natural sediment

At the end of the 21-day treatment period, the inoculated S. pumilus had 7% root colonization by F. geosporum, while the control had 3% AMF colonization. Growth trial 3 was run for 28 days with the F. geosporum inoculated group showing higher survival under the simulated saltmarsh conditions than the lower colonized control (63% survival compared to 40%) (Figure 15).

Figure 15. Survival of Sporobolus pumilus, derived from seed collected at Wolfville Harbour, under simulated saltmarsh conditions for 28 days during mesocosm growth trial 3. The inoculated plants had 7% colonization by Funneliformis geosporum while the control had 3% AMF colonization prior to entering the mesocosm bench at day 0.

Shoot length of AMF inoculated S. pumilus plants was not significantly different than that of the control plants after the 21-day treatment period (p = 0.10, α = 0.05). The shoot length of the inoculated group was significantly higher after the 28-day growth trial (p <

0.001, α = 0.05) (Figure 16). The shoot length increase over the 28-day trial was significantly higher for the inoculated group (p = 0.018, α = 0.05) (Figure 16).

Figure 16. Shoot length of Sporobolus pumilus growth trial 3 under simulated saltmarsh conditions using a tidal mesocosm bench. Blue line indicates the average shoot length increase for each group over the duration of the 28-day trial. Inoculated group was treated with Funneliformis geosporum (7% AMF colonization pre-trial); control was grown in natural saltmarsh sediment (3% AMF colonization pre-trial).

3.4 Discussion

3.4.1 Growth trial 1 – rhizome, sterile sand

The first growth trial resulted in the greatest survival and growth difference between the AMF inoculated S. pumilus plants and the uninoculated control group. After the 21-day inoculation period, evidence of AMF influence was already present, with a significantly greater overall shoot length of S. pumilus prior to simulated saltmarsh exposure in the mesocosm. One other factor that could have contributed to this difference in shoot length, prior to mesocosm exposure, was the sediment used to produce the inoculant. As shown in

Chapter 2 (Table 4), the sediment from Wolfville saltmarsh had a large amount of nutrients

present, which may be much higher than the autoclaved sand used for the control, although the chemical composition was not tested for the sand. Heating soils increases the nutrient availability of the substrate (Badia and Marti, 2003), but the additional nutrients provided by the saltmarsh sediment derived inoculant may provide growth benefits for the plants or other sediment microbiota. Conversely, it also introduces possible stressors such as salts, which are naturally present in the sediment used to create the inoculant. The AMF present in the inoculant may remedy the increase in salt, while maintaining the benefits of the added mineral nutrients, although the exact mechanism is not known (Estrada et al., 2013).

Survival of inoculated S. pumilus was much higher during the first trial, indicating that F. geosporum may offset the shock of the highly saline saltmarsh conditions on the plants. Previous research has shown improved salt stress tolerance by plants when colonized by AMF (Giri et al., 2007). Early survival is fundamental to healthy plant community establishment and sustainability (Cooke and Lefor, 1990). Better early survival reduces the density in which S. pumilus would have to be out-planted in saltmarsh restoration projects, saving both time and money. Additionally, this trial showed that inoculation increased the growth rates and final length of S. pumilus under simulated saltmarsh conditions, which are also factors that contribute to the successful establishment of saltmarsh ecosystems; slowing tidal waters leading to increased sedimentation and saltmarsh growth (Fagherazzi et al.,

2006).

3.4.2 Growth trial 2 – rhizome, natural sediment

This trial compared our AMF trap pot inoculant against the use of natural saltmarsh sediment containing the regular level of nutrients and microorganisms surrounding S. pumilus roots. The mixture of natural sediment and autoclaved sand (in a 1:1 ratio) was used

to give the control treatment the same texture as the inoculant. As it was shown in the previous chapters, sediment characteristics, including porosity, may play a role in the colonization of S. pumilus roots by AMF, therefore similar sediment types were a necessity to make this trial representative of testing natural sediment against the propagated inoculant.

As in trial 1, the shoot lengths of AMF inoculated S. pumilus were significantly higher than the control group despite similar nutrient levels and lower AMF colonization than the previous trial (17% for the inoculated and 9% for the control). The significantly larger size of the inoculated S. pumilus persisted throughout the 48-day saltmarsh trial, although the growth rates of the two groups were not significantly different.

3.4.3 Growth trial 3 – seed, natural sediment

Trial 3 was the only trial to use seed derived S. pumilus as opposed to rhizome derived plants. Conditions of trial 2 were repeated to determine if the same trends would be found for seed-derived plants, as for rhizome-derived plants. Although approaches of restoration in Nova Scotia have not focused on active planting, many saltmarsh restoration projects to date have focused on transplanting S. pumilus derived from collected seed stock due to its easy collection and low cost, but it has been shown that many of these saltmarsh restoration projects fail (Cooke and Lefor, 1990). Thus, a comparison between seed and rhizome was completed to determine if one method of saltmarsh plant propagation was superior to the other for restoration purposes.

The inoculated group had better survival than the control group, even though AMF colonization, prior to the mesocosm growth trial, was low for both groups (7% and 3% respectively, as assessed 21-days after treatment). Additionally, this trial could only be conducted for 28 days, due to the rapid death of both S. pumilus groups. There was no

difference in S. pumilus size after the 21-day inoculation period, but after the 28-day growth trial the inoculated group had grown significantly larger due to the increased growth rate.

Comparison of the survival of seed derived S. pumilus (trial 3) to the rhizome derived plants (used in trial 1 and 2) showed that rhizome derived plants had better survival rates, possibly due to differences in the AMF colonization abilities of adult tissue to juvenile tissue, although this was not tested. Previous research on the seagrass (submerged angiosperm)

Cymodocea nodosa showed that vegetative progeny from adult clones had a better chance of substrate colonization than seed derived C. nodosa (Balestri and Lardicci, 2012). These results are mirrored by our experiment which showed higher survival for rhizome derived S. pumilus compared to seed derived plants, indicating that future restoration should move toward using inoculated rhizome-derived S. pumilus instead of seed-derived.

3.4.4 Concluding remarks

Collectively these three simulated saltmarsh growth trials demonstrated that survival was consistently lower for noninoculated Sporobolus pumilus plants. Sporobolus pumilus rhizome treated with F. geosporum inoculant grew larger than either of their respective control groups, although the growth rate was not always higher for the inoculated group under simulated saltmarsh conditions. Comparison of the survival of seed derived S. pumilus

(trial 3) to the trial 1 and 2 rhizome derived plants showed that rhizome derived plants had greater survival overall, indicating that it may be beneficial to use rhizome over seed in saltmarsh restoration.

Conclusions

We demonstrated fungal sediment communities differ and are dependent on the collection location as well as the aboveground vegetation in saltmarshes. These fungal communities may be tied to sediment type and the nutrients available. Metaamplicon sequencing revealed a high proportion of terrestrial fungi present which opens an avenue for investigating whether these presumably ‘amphibious fungi’ are metabolically active in saltmarshes.

AMF diversity within S. pumilus of the Minas Basin, NS saltmarshes was limited to one species, likely F. geosporum. Seasonal variation in AMF colonization was shown for

Wolfville, NS indicating there may be an advantage of storing AMF over the winter dormancy period at this site. AMF colonization was similar for Wolfville and Windsor but was significantly lower for Kingsport, indicating that the sediment characteristics may play a role in the strength of this mutualism.

Inoculation of S. pumilus with saltmarsh native F. geosporum resulted in higher survival under simulated saltmarsh conditions in all growth trials. These AMF inoculated S. pumilus also grew larger than the control groups in each trial. Rhizome derived S. pumilus showed greater survival under simulated saltmarsh conditions than seed derived plants, indicating they should be the preferred plant stock used for saltmarsh restoration.

These findings, taken together, provide guidance for future saltmarsh restoration in

Atlantic Canada as additional climate stress is predicted to increase in the coming years.

References

Abdel-Wahab, M.A. and T. Nagahama. 2012. Halosarpheia japonica sp. nov.

(Halosphaeriales, Ascomycota) from marine habitats in Japan. Mycological Progress

11:85-92.

Alguacil, M., M.P. Torres, A. Montesinos-Navarro, and A. Roldán. 2016. Soil characteristics

driving arbuscular mycorrhizal fungal communities in semiarid Mediterranean soils.

Applied and Environmental Microbiology 82:3348-3356.

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.

Andersson, A.K. and M. Cohn. 2017. Naumovozyma castellii: An alternative model for

budding yeast molecular biology. Yeast 34:95-109.

Andrews, S. 2010. FastQC: A quality control tool for high throughput sequence data. URL:

http://www.bioinformatics.babraham.ac.uk/projects/fastqc/

Badia, D. and C. Marti. 2003. Plant ash and heat intensity effects on chemical and physical

properties of two contrasting soils. Arid Land Research and Management 17:23-41.

Balestri, E. and C. Lardicci. 2012. Nursery‐propagated plants from seed: A novel tool to

improve the effectiveness and sustainability of seagrass restoration. Journal of

Applied Ecology 49:1426-1435.

Bertness, M.D. 1991. Zonation of Spartina patens and Spartina alterniflora in New England

salt marsh. Ecology 72:138-148.

Bleakney, J.S. and K.B. Meyer. 1979. Observations on saltmarsh pools, Minas Basin, Nova

Scotia 1965-1977. Proceedings of the Nova Scotian Institute of Science 29:353-371.

Boesch, D.F. and E.R. Turner. 1984. Dependence of fishery species on salt marshes: The role

of food and refuge. Estuaries 7:460-468.

Bowron, T., N. Neatt, D. van Proosdij, J. Lundholm, and J. Graham. 2011. Macro-tidal salt

marsh ecosystem response to culvert expansion. Restoration Ecology 19(3): 307-322.

Bowron, T.M., N. Neatt, D. van Proosdij, and J. Lundholm. 2012. Tidal Marsh Restoration.

Washington: Island Press/Center for Resource Economics, 191-209.

Broome, S.W., E.D. Seneca, and W.W. Woodhouse Jr. 1988. Tidal salt marsh restoration.

Aquatic Botany 32:1-22.

Buchan, A., S.Y. Newell, J.I.L. Moreta, and M.A. Moran. 2002. Analysis of internal

transcribed spacer (ITS) regions of rRNA genes in fungal communities in a

southeastern U.S. salt marsh. Microbial Ecology 43:329-340.

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 salt marsh grass

Spartina patens, arbuscular mycorrhizal fungi and sediment bacteria during growing

season. Soil Biology & Biochemistry 35:501-511.

Callaway, J.C. 2005. The challenge of restoring functioning salt marsh ecosystems. Journal

of Coastal Research SI40:24-36.

Callaway, J.C., J.B. Zedler, and D.L. Ross. 1997. Using tidal salt marsh mesocosms to aid

wetland restoration. Restoration Ecology 5:135-146

Comeau, A.M., G.M. Douglas, and M.G.I. Langille. 2017. Microbiome Helper: a custom and

streamlined workﬂow for microbiome research. mSystems 2:e00127-16.

Condit, R., N. Pitman, E.G. Leigh Jr., J. Chave, J. Terborgh, R.B. Foster, P. Núñez, S.

Aguilar, R. Valencia, G. Villa, H.C. Muller-Landau, E. Losos, and S.P. Hubbell.

2002. Beta diversity in tropical forest trees. Science 295:666-669.

Cooke, J.C. and M.W. Lefor. 1990. Comparison of vesicular-arbuscular mycorrhizae in

plants from disturbed and adjacent undisturbed regions of a coastal salt marsh 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 salt-marsh plant zonation depending

on nutrient supply. Journal of Ecology 96:431-437.

Daddow, R.L. and G.E. Warrington. 1983. Growth-limiting soil bulk densities as influenced

by soil texture. Watershed Systems Development Group, USDA Forest Service. d’Entremont, T.W., A.K. Walker and J.C. López-Gutiérrez. 2018. Examining arbuscular

mycorrhizal fungi in saltmarsh hay (Spartina patens) and smooth cordgrass (Spartina

alterniflora) in the Minas Basin, Nova Scotia. Northeastern Naturalist 25:72-86.

Department of Natural Resources. 2013. Fundy dykelands and wildlife. URL:

https://novascotia.ca/natr/wildlife/habitats/dykelands/.

Dighton, J. 2016. Fungi in Ecosystem Processes. Boca Raton: CRC Press.

Eash, N.S., T.J. Sauer, D. O’Dell, and E. Odoi. 2016. Soil Science Simplified. Hoboken:

John Wiley & Sons, Inc..

Elmer, W.H. and R.E. Marra. 2011. New species of Fusarium associated with dieback of

Spartina alterniflora in Atlantic salt marshes. Mycologia 103:806-819.

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.

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.

Fagherazzi, S., L. Carniello, L. D’Alpaos, and A. Defina. 2006. Critical bifurcation of

shallow microtidal landforms in tidal flats and salt marshes. PNAS 103:8337-8341.

Filip, Z. and J.J. Alberts. 1993. Formation of humic-like substances by fungi epiphytic on

Spartina alterniflora. Estuaries 16:385-390.

Ford, R.J. and F.F. Becker. 1982. The characterization of trypan blue-induced tumors in

Wistar rats. The American Journal of Pathology 106:326-331.

Freeman, J. and E. Ward. 2004. Gaeumannomyces graminis, the take-all fungus and its

relatives. Molecular Plant Pathology 5:235-252.

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.

Giovanetti, M. and B. Mosse. 1980. An evaluation of techniques for measuring vesicular

arbuscular mycorrhizal infection in roots. New Phytologist 84:489-500.

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.

Government of Nova Scotia. 2014. Historic wetland loss in Nova Scotia. URL:

https://novascotia.ca/nse/wetland/historic-wetland-loss-ns.asp.

Haskett, E.C. 2018. Response of intertidal benthic invertebrate assemblages to high and low

concentrations of suspended sediment. Acadia University Thesis; Access date: 18

May 2019.

Hassett, B.T., A.L. Ducluzeau, R.E. Collins, and R. Gradinger. 2017. Spatial distribution of

aquatic marine fungi across the western arctic and sub-arctic. Environmental

Microbiology 19:475-484.

Hildebrandt, U., Janetta, K., Ouzaid, F., Renne, B., Nawrath, K., and Bothe, H. 2001.

Arbuscular mycorrhizal colonization of halophytes in central European salt marshes.

Mycorrhiza 10:175.

Hoefnagels, M.H., S.W. Broome, and S.R. Shafer. 1993. Vesicular-arbuscular mycorrhizae

in salt marshes in North Carolina. Estuaries 16:851-858.

Huss, M.J. 1993. Spatial distribution among mycelial individuals of Lycoperdon pyriforme

occurring on decaying logs. Mycological Research 97:1119-1125.

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.

Kendrick, B. 2017. The Fifth Kingdom. Indianapolis: Hackett Publishing Company, Inc.

Kõljalg, U., K‐H. Larsson, K. Abarenkov, R.H. Nilsson, I.J. Alexander, U. Eberhardt, S.

Erland, K. Høiland, R. Kjøller, E. Larsson, et al. 2005. UNITE: a database providing

web‐based methods for the molecular identification of ectomycorrhizal fungi. New

Phytologist 166:1063-1068.

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.

Kurtzman, C.P., J.W. Fell, and T. Boekhout. 2011. The Yeasts, A Taxonomic Study. San

Diego: Elsevier.

Landry, B. 2016. Biodiversity of marine fungi in Kingsport, Nova Scotia, seawater and their

oil degradation potential. Acadia University Thesis; Access date: 18 May 2019.

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.

Law, B.A., P.S. Hill, T.G. Milligan, and V. Zions. 2019. Temporal and spatial changes in

grain size on a macro-tidal channel-flat complex: Results from Kingsport, Nova

Scotia, Bay of Fundy. Ocean Dynamics 69:239-252.

Liu, C., W. Qu, and W. Zhang. 2007. Analyses of Tremella aurantialba (Tremellaceae) and

its analog species inferred from ITS sequences. Acta Botanica Yunnanica 29:237-

242.

Lumibao, C.Y., S. Formel, V. Elango, J.H. Pardue, M. Blum, and S.A. Van Bael. 2018.

Persisting responses of salt marsh fungal communities to the Deepwater Horizon oil

spill. Science of the Total Environment 642:904-913.

Martin, S.L., S.J. Mooney, M.J. Dickinson, and H.M. West. 2012. The effects of

simultaneous root colonisation by three Glomus species on soil pore characteristics.

Soil Biology and Biochemisty 49:167-173.

Mcleod, E., G.L. Chmura, S. Bouillon, R. Salm, M. Björk, C.M. Duarte, C.E. Lovelock,

W.H. Schlesinger, and B.R. Silliman. 2011. A blueprint for blue carbon: Toward an

improved understanding of the role of vegetated coastal habitats in sequestering CO2.

Frontiers in Ecology and the Environment 9:552-560.

Morton, J.B., S.P. Bentivenga, and W.W. Wheeler. 1993. Germ plasm in the International

Collection of Arbuscular and Vesicular- Arbuscular Mycorrhizal Fungi (INVAM)

and procedures for culture development, documentation, and storage. Mycotaxon 48:

491-528.

Nilsson, R.H., Kristiansson, E., Ryberg, M., Hallenberg, N., and Larsson, K-H. 2008.

Intraspecific ITS variability in the kingdom fungi as expressed in the international

sequence databases and its implications for molecular species identification.

Evolutionary Bioinformatics 4:193.

Op De Beeck, M., B. Lievens, P. Busschaert, S. Declerck, J. Vangronsveld, and J.V.

Colpaert. 2014. Comparison and validation of some ITS primer pairs useful for fungal

metabarcoding studies. PLoS ONE 9:e97629.

Parniske, M. 2008. Arbuscular mycorrhiza: The mother of all plant root endosymbioses.

Nature Reviews Microbiology 6:763-775.

Phillips, J.M. and D.S. Hayman. 1970. Improved procedures for clearing roots and staining

parasitic and vesicular-arbuscular mycorrhizal fungi for rapid assessment of infection.

Transactions of the British Mycological Society 55:158-161.

Pirozynski, K.A. and D.W. Malloch. 1975. The origin of land plants: A matter of

mycotrophism. BioSystems 6:153-164.

Porter, C., J. Lundholm, T. Bowron, B. Lemieux, D. van Proosdij, N. Neatt, and J. Graham.

2015. Classification and environmental correlates of tidal wetland vegetation in Nova

Scotia. Botany 93:825-841.

Prosser, J.I. 2002. Molecular and functional diversity in soil micro-organisms. Plant and Soil

244:9-17.

Rastogi, R.P., Richa, A. Kumar, M.B. Tyagi, and R.P. Sinha. 2010. Molecular mechanisms

of ultraviolet radiation-induced DNA damage and repair. Journal of Nucleic Acids

2010:1-32.

R Core Team. 2019. R: A language and environment for statistical computing. R Foundation

for Statistical Computing, Vienna, Austria.

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.

Shepard, C.C., C.M. Crain, and M.W. Beck. 2011. The protective role of coastal marshes: A

systematic review and meta-analysis. PLoS ONE 6:e27374.

Schüßler, A, and C. Walker. 2010. The Glomeromycota. A species list with new families and

new genera. Edinburgh & Kew, UK: The Royal Botanic Garden; Munich, Germany:

Botanische Staatssammlung Munich; Oregon, USA: Oregon State University.

Simmons, D.R., T.Y. James, A.F. Meyer, and J.E. Longcore. 2009. Lobulomycetales, a new

order in the Chytridiomycota. Mycological Research 113:450-460.

Smith, S. and D. Read. 2008. Mycorrhizal Symbiosis. San Diego: Academic Press.

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.

Sterflinger, K., M. Hain, J. Scholz, and K. Wasson. 2001. Fungal infections of a colonial

marine invertebrate: diversity and morphological consequences. Facies 45:31-37.

Stockinger, K., M. Krüger, and A. Schüssler. 2010. DNA barcoding of arbuscular

mycorrhizal fungi. New Phytologist 187:461-474.

Thomsen, P.F. and E. Willerslev. 2015. Environmental DNA – an emerging tool in

conservation for monitoring past and present biodiversity. Biological Conservation

183:4-18. van Proosdij, D. and S.M. Townsend. 2006. Spatial and temporal patterns of salt marsh

colonization following causeway construction in the Bay of Fundy. Journal of Coastal

Research SI39:1859-1863. van Proosdij, D., J. Lundholm, N. Neatt, and T. Bowron. 2010. Ecological re-engineering of

a freshwater impoundment for salt marsh restoration in a hypertidal system.

Ecological Engineering 36:1314-1332.

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.

Walker, A.K. and J. Campbell. 2010. Marine fungal diversity: A comparison of natural and

created salt marshes of the north-central Gulf of Mexico. Mycologia 102:513-521.

Warren, S.R., P.E. Fell, R. Rozsa, H.A. Brawley, A.C. Orstead, E.T. Olson, V. Swamy, and

W.A. Niering. 2002. Salt marsh restoration in Connecticut: 20 years of science and

management. Restoration Ecology 10:497-513.

Wilde, P., A. Manal, M. Stodden, E. Sieverding, U. Hildebrandt, and H. Bothe. 2009.

Biodiversity of arbuscular mycorrhizal fungi in roots and soils of two salt marshes.

Environmental Microbiology 11:1548-1561.

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 salt marsh

macrophytes. Ecology 96:840-849.

Yamato, M., Ikeda, S., and Iwase, K. 2008. Community of arbuscular mycorrhizal fungi in a

coastal vegetation on Okinawa Island and effect of the isolated fungi on growth of

Sorghum under salt-treated conditions. Mycorrhiza 18:241-249.