- Nutrient enrichment impacts ecosystems globally. Population history, especially past resource environments, of numerically dominant plant species may affect their responses to subsequent changes in nutrient availability. Eutrophication can also alter plant–microbe interactions via direct effects on associated microbial communities or indirect effects on dominant species’ biomass production/allocation as a result of modified plant–soil interactions.
- We combined a greenhouse common garden and a field reciprocal transplant of a salt marsh foundation species (Spartina alterniflora) within a long-term, whole-ecosystem, nutrient-enrichment study to determine whether enrichment affects plant production and microbial community structure differently depending on plant population history. For the greenhouse portion, we collected 20 S. alterniflora genotypes—10 from an enriched creek that had received elevated nutrient inputs for 10 years and 10 from an unenriched reference creek—and reared them in a common garden for 1 year. For the field portion, we conducted a 2-year, fully crossed reciprocal transplant experiment with two gardens each at the enriched and unenriched sites; we examined the effects of source site (i.e. population history), garden site and plant genotype.
- After 2 years, plants in enriched gardens had higher above-ground biomass and altered below-ground allocation compared to plants in unenriched gardens. However, performance also depended on plant population history: plants from the enriched site had decreased above-ground and rhizome production compared to plants from the unenriched site, most notably in unenriched gardens. In addition, almost all above- and below-ground traits varied depending on plant genotypic identity.
- Effects of nutrient enrichment on the associated microbial community were also pronounced. Following 1 year in common garden, microbial community structure varied by plant population history and S. alterniflora genotypic identity. However, at the end of the reciprocal transplant, microbial communities differed primarily between enriched and unenriched gardens.
- Synthesis. Nutrient enrichment can impact plant foundation species and associated soil microbes in the short term. Most importantly, nutrient enrichment can also have long-lasting effects on plant populations and associated microbial communities that potentially compromise their ability to respond to changing resource conditions in the future.
Nutrient enrichment, also known as eutrophication, affects terrestrial and aquatic ecosystems world-wide (Carpenter et al., 1998; Sinha et al., 2017; Smith et al., 2006; Vitousek et al., 1997). Excessive inputs of phosphorus and nitrogen can shift community composition (Avolio et al., 2014; Leff et al., 2015; Payne et al., 2013; Suding et al., 2005) and species diversity (Fujita et al., 2014; Seehausen et al., 1997) across multiple trophic levels via both the random loss of rare species and the non-random loss of specific species or functional groups (e.g. perennials; Stevens et al., 2004; Suding et al., 2005). The effects of eutrophication may be greater for plant species and microbial communities adapted to low nutrient conditions than those from moderate nutrient environments (Ramirez et al., 2012; Vitousek et al., 1997). Conversely, species and communities historically exposed to a eutrophic environment may be more adversely affected by a subsequent decrease in nutrient availability, as ecological optima, life histories and/or allocation trade-offs may have shifted as a result of nutrient-induced selection (Alexander et al., 2016; Frisch et al., 2014; Snell-Rood et al., 2015). Because these changes can, in turn, influence ecosystem function and stability (Hautier et al., 2014; Isbell et al., 2013; Smith & Schindler, 2009), understanding how the effects of eutrophication vary depending on both past and present resource environments is critical for management efforts.
In systems dominated by foundation species, community structure and ecosystem function may depend primarily on the responses of these key species to nutrient enrichment (Altieri & Witman, 2006; Deegan et al., 2012; Hughes et al., 2009; Isbell et al., 2013). For example, fertilization effects on numerically dominant plant species may manifest in the form of (a) increased productivity, with consequent effects on species diversity due to loss of rare species and/or changes in trait optima (Suding et al., 2005); (b) decreased productivity, with the loss of dominant species having a greater effect than random species loss (Isbell et al., 2013); and/or (c) changes in above- and below-ground allocation, which alter physical structure and ecosystem stability (Deegan et al., 2012). In addition, intraspecific variation within (Hairston et al., 1999; Härnström et al., 2011; Tomas et al., 2011) and across (Alexander et al., 2016; Tuckett et al., 2013) populations may also reflect and/or mediate the effects of nutrient enrichment. For example, following a period of eutrophication and harmful cyanobacterial blooms, genotypes of the key aquatic consumer Daphnia evolved increased ability to cope with a diet including cyanobacteria, altering the net effect of eutrophication on the lake community (Hairston et al., 1999). However, relatively little is known about how intraspecific variation within numerically dominant species influences population and community responses to nutrient enrichment since the focus is often on community-level trait shifts as opposed to population-level genotypic and phenotypic changes (Jung et al., 2014; Lepš et al., 2011; Siefert & Ritchie, 2016).
Changes in the production and biomass allocation of foundation plant species may further affect, and be affected by, responses of the associated soil microbial community to nutrient enrichment. Understanding how plant–microbial relationships change with nutrient enrichment has important implications for microbial biomass and activity (Kearns et al., 2016; Suding et al., 2008), plant and microbial functional traits (Leff et al., 2015; Sayer et al., 2017) and decomposition rates (Deegan et al., 2012). Studies looking at the effects of nutrient inputs across ecosystems have found relatively consistent changes in microbial community structure, with copiotrophic and oligotrophic taxa increasing and decreasing in abundance respectively (Leff et al., 2015; Ramirez et al., 2010, 2012). However, these shifts may be more nuanced than previously recognized, as eutrophication can also alter the proportion and identity of active versus dormant microbial taxa (Kearns et al., 2016). In addition, soil legacy effects of press and pulse disturbances such as herbivory, drought and extreme precipitation—and potentially nutrient enrichment—can alter the strength of plant–soil interactions both among (Fry et al., 2018; Kaisermann et al., 2017) and within (Crawford & Hawkes, 2020; Kostenko et al., 2012) species, with potential effects on multitrophic interactions (Kostenko et al., 2012), plant species richness (Fry et al., 2018; Kaisermann et al., 2017) and genetic composition (Crawford & Hawkes, 2020). Furthermore, intraspecific variation within plant foundation species (i.e. genotype or lineage) can also play a key role in structuring rhizosphere bacterial communities (Bowen et al., 2017; Schweitzer et al., 2008; Zogg et al., 2018), yet whether and how nutrient enrichment affects this relationship is unknown.
To examine how nutrient enrichment effects on plant responses depend on the interplay of population history (i.e. past nutrient conditions), plant genetic identity and plant–microbe interactions, we conducted a field reciprocal transplant experiment with the salt marsh foundation species Spartina alterniflora within the context of a long-term, whole-ecosystem, nutrient-enrichment experiment (the TIDE project; Deegan et al., 2007). Salt marshes are an ecologically important and economically valuable habitat (Barbier et al., 2011), yet they are in decline globally (Duarte et al., 2008), and eutrophication is a key driver of marsh habitat loss (Pardo et al., 2011). Prior results of the TIDE project demonstrated that marsh collapse along the edge of tidal creeks can result from a combination of elevated microbial respiration and changes in biomass allocation of tall-form S. alterniflora that dominates low marsh elevations (Deegan et al., 2012). However, much less is known about the responses of short-form S. alterniflora and its associated microbial community to nutrient enrichment. Short-form S. alterniflora dominates high marsh elevations that experience relatively infrequent inundation (i.e. only 30% of high tides flood the high marsh; Johnson et al., 2016), resulting in lower nutrient delivery than in the low marsh (Deegan et al., 2007; Johnson et al., 2016).
We tested the effects of the current nutrient environment (enriched vs. unenriched) on short-form S. alterniflora production and soil microbial community structure in the high marsh over the course of 2 years. We compared multiple plant genotypes originally from enriched and unenriched creeks that were transplanted back to both enriched and unenriched sites to test our hypothesis that the effects of nutrient enrichment would vary depending on both plant population history and genotypic identity. This approach allowed us to differentiate between short-term (i.e. duration of the experiment) and long-term (i.e. population history) effects of nutrient enrichment on the foundation species S. alterniflora and its associated microbial community. Specifically, we predicted that transplants from enriched sites would perform worse in unenriched gardens than transplants from unenriched sites, and that transplants from unenriched sites would perform similarly to or better than transplants from enriched sites in enriched gardens. In addition, we expected Spartina traits to vary depending on genotypic identity given previous research that found strong effects of genotype on density, morphology and production (Hughes et al., 2014; Proffitt et al., 2003, 2005; Zerebecki et al., 2017). Lastly, we predicted that both Spartina genotype and nutrient availability would influence associated microbial community structure given earlier studies that identified independent effects of host genetics (Zogg et al., 2018) and environmental conditions (Kearns et al., 2016) on microbial composition and diversity, with a goal of this study being to see how the relative importance of these factors changed over the course of the experiment.
2 MATERIALS AND METHODS
2.1 Study system
The Plum Island Sound Estuary in Massachusetts, USA is the site of the TIDE long-term, whole-ecosystem, nutrient enrichment experiment (Deegan et al., 2007). Briefly, a pair of primary tidal creeks with similar characteristics (i.e. length, volume, physical and chemical properties and plant communities; Deegan et al., 2007, 2012) located in the Rowley River sub-estuary were identified: West Creek was assigned as the reference creek and Sweeney Creek was assigned as the treatment (nutrient-enriched) creek (Figure S1.1a). The ecosystem-scale nutrient treatment was applied by adding nitrogen (in the form of NaNO3) directly to flooding tidal water from mid-May to mid-September starting in 2004; phosphorus (in the form of NaH2PO4) was also added from 2004 to 2012. This method closely mimics anthropogenic nutrient loading and enriches the creek to levels 10–15 and 5–15 times greater than background levels of NO3- and PO4- in the estuary, respectively, corresponding to moderate to high eutrophication (Deegan et al., 2007, 2012).
The plant communities at West and Sweeney consisted primarily of tall-form S. alterniflora in the low marsh along creek banks and short-form S. alterniflora, Spartina patens, and Distichlis spicata on the high marsh platform (Deegan et al., 2007; Johnson et al., 2016). Prior work in this system suggested that effects of nutrient enrichment in the high marsh were relatively weak and inconsistent (Deegan et al., 2007; Johnson et al., 2016), although they did not experimentally manipulate the plants or control for potential differences across sites due to genotypic and/or phenotypic variation.
We conducted a reciprocal transplant with two gardens at both sites (N = 4 gardens total) to explore the effects of nutrient enrichment in the high marsh using a replicated and controlled experimental design. At each site, we chose two areas in the high marsh dominated by short-form S. alterniflora (hereafter, Spartina), with gardens located along the two primary branches of each tidal creek (West Garden 1: N 42.7382, W 70.8486 and West Garden 2: N 42.7385, W 70.8476—hereafter, unenriched gardens; Sweeney Garden 1: N 42.7223, W 70.8478 and Sweeney Garden 2: N 42.7219, W 70.8488—hereafter, enriched gardens; see Figure S1.1a for a map of sites and gardens). We selected two gardens at each site to better encompass any potential environmental variation within sites, and to decrease the likelihood of erroneously attributing location-specific differences to enrichment effects. During the experiment, we monitored plant species composition and per cent cover, as well as sediment salinity and oxygen, to characterize environmental conditions within and across garden sites. Importantly, a comparison of these characteristics did not identify significant differences between sites (see Appendix S1 for detailed results). We did not monitor herbivore abundance within gardens because grazing is low at these sites (Johnson et al., 2016) and consumer regulation of Spartina is weak in this system (Johnson & Jessen, 2008). In addition, we observed almost no grazing damage on experimental plants, so herbivory is unlikely to differ between sites or among gardens. Lastly, prior work indicates that local-scale variability in salt marsh microbial communities is small (Bowen et al., 2012) and that both microbial community structure (Bowen et al., 2009a; Buchan et al., 2003; Kearns et al., 2016) and bacterial production (Bowen et al., 2009b) within habitats are relatively consistent, further suggesting minimal differences in microbial properties between sites.
2.2 Collection, propagation and identification of Spartina genotypes
In September 2013, we collected a ‘plug’ (~15 cm diameter) of above- and below-ground biomass of short-form Spartina every 50 m along both sides of each 250 m branch of West Creek and Sweeney Creek (Figure S1.1a). This approach allowed us to sample plants that experienced a range of environmental conditions (e.g. differences in tidal inundation frequency and duration) within each marsh and gave us a pool of plants from each creek that spanned a distance of 500 m and originated from both the landward and seaward sides of both branches. We then isolated three unconnected stems (including attached rhizome and root) from each plug in the laboratory to create a stock of unique individuals. Each stem was planted in a 15.2 cm diameter × 14.6 cm height pot containing a 50:50 mix of commercial potting soil (Miracle-Gro Nature's Care organic garden soil) and sand. Plants were propagated in a common greenhouse environment at the Northeastern University Marine Science Center in Nahant, MA (located 30 miles from the Plum Island Sound Estuary), irrigated with freshwater daily, and immersed in seawater once weekly during the growing season (May–October) to approximate tidal inundation frequency at the field site. Plants were split into multiple pots approximately every 3 months to facilitate the production of new stems. To maintain distinct genotypes, plants were checked twice weekly during the reproductive season (July–September) and any flowers were removed.
We selected 20 unique Spartina genotypes from the greenhouse stock for our reciprocal transplant experiment that started September 2014 (see details below). Thus, plants used in the field reciprocal transplant were grown in a common greenhouse environment for 1 year prior to being transplanted to the field. Prior to the start of the reciprocal transplant, we confirmed that all genotypes used in the field experiment were genetically distinct by genotyping each individual using 12 microsatellite loci (see Appendix S2 for details).
2.3 Field reciprocal transplant experiment
In August 2014, the 20 selected Spartina genotypes (10 from the enriched creek and 10 from the unenriched creek) were separated into clusters of three to five interconnected stems of similar sizes (including associated rhizomes and roots) and planted in 10.6 L (15.2 cm diameter) pots containing a 50:50 mix of commercial potting soil and sand. Prior to planting, pot bottoms were removed and pots were lined with non-reactive mesh (i.e. windowscreen) to expose plants to local edaphic conditions in the field while allowing quantification of above- and below-ground biomass of the transplants at the end of the 2-year experiment. We counted stem density and measured stem height of all plants in the greenhouse prior to transplanting to the field gardens; transplants did not differ significantly in density or height between source sites at the start of the experiment.
Two hundred forty plants were transplanted to one of four gardens in the field (two each at the enriched and unenriched sites respectively) in September 2014. Each 4 m × 20 m garden consisted of three rows (i.e. blocks) with 20 plants each; rows were separated by 2 m, and plants within rows were separated by 1 m. Plants were randomly assigned within each row, with one replicate of each genotype included in every row for a total of three replicates per garden (see Figure S1.1b for diagram of experimental design). By controlling for genotypic identity in our design—a factor that can strongly affect plant morphology and primary production (Hughes, 2014; Hughes et al., 2014)—we avoided potentially confounding differences in identity between creeks and increased our ability to detect effects of nutrient enrichment. We measured stem density and stem height (including maximum height and average height of three haphazardly selected stems) monthly during the growing season (May–October) in 2015 and 2016. We also clipped flowers at monthly intervals during the reproductive season to prevent gene flow via sexual reproduction. At the end of the experiment in November 2016, we measured stem density as well as total height and leaf width of 10 haphazardly selected stems; we then harvested plants to determine above- and below-ground biomass. To assess below-ground allocation, we separated roots and rhizomes using a fine mesh (~0.8 mm) sieve. All biomass samples (i.e. above-ground, roots and rhizomes) were dried for at least 72 hr at 60℃ before weighing.
Prior to harvesting, we collected leaf tissue from each plant to assess Spartina elemental composition, including per cent carbon (C), per cent nitrogen (N) and C:N ratio. Samples were stored at −20℃ prior to processing, then rinsed with deionized water, dried for at least 48 hr at 60℃, homogenized using a Retsch MM400 mixer mill and analysed on a Costech ECS 4010 elemental analyser interfaced to a Thermo DeltaPlus Advantage mass spectrometer.
To assess the structure of the sediment microbial communities, we collected five sediment cores (top 3 cm) from a subset of genotypes in each garden using a sterile 10 cm3 syringe corer. The five sediment cores from each genotype were homogenized in a sterile centrifuge tube and two aliquots from the homogenized pool were flash frozen for subsequent analysis of the microbial community. We collected sediment samples at four times during the study, chosen to identify differences between the first (greenhouse common garden) and second (field reciprocal transplant) phases of the project, as well as changes relative to the first and second growing seasons in the field: (a) August 2014, following 1 year in a common greenhouse environment and prior to being transplanted to the field; (b) May 2015, 8 months after being transplanted to the field and at the start of the first growing season; (c) August 2015, at the end of the first growing season; and (d) August 2016, at the end of the second growing season (see Appendix S3 for details; microbial data are available at NCBI’s Sequence Read Archive—accession number PRJNA631747).
2.4 Data analysis
We did a series of analyses on plant responses (a) over the course of the experiment to determine how quickly effects manifested during the experiment and whether patterns changed across years, and (b) at the end of the experiment to identify the cumulative effects of each treatment on plant survival, traits and biomass production. First, we assessed changes in Spartina density and morphology through time for each year of the experiment (2015 and 2016) separately (given seasonal reset during winter 2015–2016) using linear mixed effect models. We included garden site, source site, garden branch nested in garden site, genotype nested in source site, and time as fixed effects, garden plot ID as a random effect (to account for non-independence of following pots through time), and independent and interactive effects of all fixed factors (see Appendix S4 for detailed results).
We then analysed whether Spartina survival, density, morphology, elemental composition (Spartina %C, %N and C:N, as well as the total C and N pools for each plant, calculated as tissue composition × above-ground biomass; Ritchie et al., 1998) or biomass varied independently or interactively by garden site or source site at the end of the experiment. This approach allowed us to test the potential for local advantage—that is, the relative performance of individuals transplanted back to their source site (e.g. plants from the enriched site in gardens at the enriched site) versus individuals transplanted to a novel site (e.g. plants from the unenriched site in gardens at the enriched site). For survival, we used a generalized linear model with a binomial error distribution and a logit link function. For the other analyses, we used linear models with the same structure as above (except without time as a factor) to look at the cumulative independent and interactive effects of garden site, source site, garden branch (nested in garden site) and genotype (nested in source site) on Spartina responses at the end of the experiment. All models of height included live stem density as a covariate, and all models of biomass included total stem density as a covariate. For the biomass data, we first conducted a multivariate analysis of variance (MANOVA) on all measures of Spartina above- and below-ground biomass using the same model structure described above; because MANOVA identified significant effects of all factors on plant biomass (see Section 3), we then examined their effects on each measure of plant production separately. To meet assumptions of normality and homoscedasticity, we log-transformed the ratios of above- to below-ground biomass and root to rhizome biomass, as well as Spartina %N and N pool. Because garden branches within transplant sites had a similar range of abiotic and biotic conditions (e.g. salinity, redox, and plant species richness, composition and per cent cover; see Appendix S1 for details) and we were interested in site-level effects that accounted for garden-level variation, we focus on the effects of garden site in the results.
To analyse the microbial data (sequence processing and quality control are described in detail in Appendix S3), we first used the Bray–Curtis similarity metric to quantify beta diversity and visualized the resulting similarity matrix using principal coordinates analysis (PCoA) implemented in QIIME (version 1.9.1; Caporaso et al., 2010). We used a nonparametric PERMANOVA to assess whether microbial communities differed significantly (α = 0.01) among garden sites, source sites, genotypes and/or garden site × source site combinations. Because we did not find a significant garden site × source site interaction (p = 0.4), we focus on the independent effects of garden site, source site and genotype in the results. Second, we calculated estimates of mean Shannon diversity derived from rarified data (see Appendix S3 for details) to (a) determine whether microbial alpha diversity changed during the course of the study by comparing Shannon diversity at the end of the greenhouse common garden (August 2014, prior to the field reciprocal transplant) and during the field reciprocal transplant (May 2015, August 2015, August 2016); and (b) examine the independent and interactive effects of garden site, source site and genotype on microbial diversity at each sampling date during the field reciprocal transplant. Third, we tested whether the primary or secondary principal components from the microbial PCoA of the August 2016 sampling were correlated with Spartina biomass measurements at the end of the experiment. Because there was minimal overlap between enriched and unenriched gardens in PCoA1 (see Section 3), we focused on the relationship between microbial PCoA1 and Spartina biomass across garden sites. For PCoA2, we used analysis of covariance (ANCOVA) to assess whether the relationship between microbial PCoA2 (covariate) and Spartina biomass differed between enriched and unenriched gardens (i.e. garden site).
All analyses were conducted in R (version 3.1.2) using the lme4 and lmerTest packages, with the exception of the microbial alpha and beta diversity analyses, which were performed using QIIME (version 1.9.1; Caporaso et al., 2010).
At the end of the 2-year experiment, there was no difference in overall survival between transplant gardens (87% and 86% survival in enriched and unenriched gardens respectively; p = 0.85) or between plants originating from different sites (87% and 86% survival of plants from enriched and unenriched sites respectively; p = 0.85). In addition, there was no evidence of local advantage (garden site × source site: p = 0.18); survival was consistently high, regardless of whether plants were in gardens at their source site or a new site.
3.2 Density and morphology
At the end of the second growing season, garden site had a significant effect on Spartina density but not morphology (garden site: density: p = 0.005; all other responses: p > 0.3; Table S5.1), with individuals in enriched gardens having 26% higher stem densities than those in unenriched gardens (Figure 1a). Source site strongly affected both density and morphology (source site: p ≤ 0.005 for all responses except maximum height; Table S5.1): plants originating from the unenriched site had 23% higher stem densities, 16% taller average heights and 8% greater leaf widths than plants originating from the enriched site (Figure 1). In addition, plants in enriched gardens had similar average and maximum heights, regardless of source site, whereas plants in unenriched gardens were taller if they were originally from unenriched gardens (Figure 1b). Genotypic identity also affected stem density, maximum and average height, and leaf width (genotype: p < 0.025 for all responses; Table S5.1; Figure 1d–f).
There were significant independent effects of garden site, source site and genotype on above- and below-ground biomass of S. alterniflora transplants, but no garden site × source site interaction (MANOVA; Table S5.2a). Plants in enriched gardens had 22% greater above-ground biomass than plants in unenriched gardens (garden site: p < 0.001; Table S5.2b; Figure 2a). While there was no effect of garden site on total below-ground biomass (Table S5.2b), plants in unenriched gardens had 21% greater root biomass than plants in enriched gardens (Figure 2d), whereas plants in enriched gardens had 21% greater rhizome biomass than plants in unenriched gardens (garden site: root: p = 0.035; rhizome: p < 0.001; Figure 2c). Garden site also influenced both above- versus below-ground allocation (p < 0.001) and root versus rhizome allocation (p < 0.001), with plants in enriched gardens having a higher ratio of above- to below-ground biomass (Figure 2b) and a lower ratio of root to rhizome biomass (Figure 2e) than plants in unenriched gardens.
Spartina biomass also depended on plant origin (i.e. source site). For example, plants originating from the enriched site had 20% less above-ground and 10% less total biomass than plants originating from the reference site (source site: above-ground biomass: p < 0.001; total biomass: p = 0.056; Figure 2a,f). There was no effect of source site on total below-ground biomass (Table S5.2b), but rhizome biomass was 12% greater for plants that came from the unenriched site than for plants that came from the enriched site (source site: p = 0.006; Figure 2c). Plants originating from the unenriched site also had a higher ratio of above- to below-ground biomass and a lower ratio of root to rhizome biomass than plants originating from the enriched site (source site: above to below: p = 0.021; root to rhizome: p = 0.036; Figure 2b,e). However, while plants originating from the unenriched site had similar above-ground and rhizome biomass across gardens, plants originating from the enriched site had significantly lower above-ground and rhizome biomass when planted in unenriched gardens compared to enriched gardens (Figure 2a,c).
There was also a strong independent effect of genotypic identity on all above- and below-ground measures, including above- and below-ground biomass, root and rhizome biomass and total plant biomass (genotype: p < 0.05 for all responses; Table S5.2b; Figure S5.1), with individual genotypes from both sites displaying a range of phenotypes across gardens.
3.4 Plant elemental content
There were significant independent effects of garden site and of source site on Spartina stoichiometry, but no garden site × source site interaction. Plants in unenriched gardens had higher %C and %N, but lower quality overall (i.e. higher C:N) than plants in enriched gardens (garden site: p < 0.001 for all responses; Table S5.3; Figure 3a–c). In addition, plants originating from the unenriched site had consistently higher %C than plants originating from the enriched site (source site: p = 0.047; Figure 3a). There was also a significant effect of genotypic identity on C content, N content and C:N ratio (genotype: p < 0.005 for all responses; Figure S5.2a–c). In addition to the effects of garden site on tissue %C and %N, analysis of the total pool of C and N for each plant (calculated as per cent composition × above-ground biomass; Ritchie et al., 1998) demonstrated that plants originating from the unenriched site had a larger pool of C and N than plants originating from the enriched site (Figure 3d,e; Table S5.3). There was also a significant effect of garden site on C pool but not N pool (garden site: C pool: p = 0.002; Figure 3d; Table S5.3), with plants in enriched gardens having a larger C pool than plants in unenriched gardens. Finally, C and N pools differed across genotypes (genotype: p < 0.05 for both responses; Figure S5.2d–e; Table S5.3).
3.5 Microbial community composition
In the greenhouse prior to planting in the field, there was a significant effect of source site on microbial community similarity: plants originating from the enriched site had distinct microbial communities from plants originating from the unenriched site, despite having been grown in a common environment for 1 year (PERMANOVA: F15,33 = 1.98, p < 0.01; Figure 4b). Genotype also influenced microbial community similarity after 1 year in the greenhouse (PERMANOVA: F15,33 = 1.98, p < 0.01; Figure S3.1). However, there was no effect of source site or genotype on microbial community similarity following transplant to the field (p > 0.05 for all sampling dates). Rather, there was an effect of garden site at the end of the experiment (PERMANOVA: F2,20 = 2.77, p < 0.01; Figure 4c).
Microbial community composition shifted substantially during the experiment (PERMANOVA: date F4,124 = 44.5, p < 0.01; Figure 4a). The microbial community in the greenhouse prior to planting in the field was distinct from the microbial community 8 months after the start of the experiment (May 2015), and composition continued to shift during the first (August 2015) and second (August 2016) growing seasons (Figure 4a). In addition, microbial diversity (calculated as Shannon diversity index; see Appendix S3 for details) increased following transplanting from the greenhouse to the field, but then decreased by the end of the reciprocal transplant experiment (Figure 5a): compared to after 1 year in the greenhouse common garden (Greenhouse 2014), microbial diversity was higher in the first year (May and August 2015), but lower in the second year (August 2016) of the field experiment (time: p < 0.001). In the middle of the first growing season (August 2015), there was also a significant effect of garden site on microbial diversity, with associated soil communities having greater diversity in unenriched gardens than enriched gardens, regardless of plant origin (garden site: p = 0.03; Figure 5b).
In addition, we found strong correlations between (a) the principal coordinate axes (PCoA1 and PCoA2) from the ordination of Bray–Curtis similarity within the microbial community, and (b) Spartina biomass and above-ground:below-ground allocation at the end of the field experiment. Microbial PCoA1 was negatively associated with the ratio of above-ground to below-ground biomass (p = 0.06, R2 = 0.19; Figure 4d). In contrast, microbial PCoA2 was positively associated with Spartina above-ground:below-ground biomass (p = 0.03, R2 = 0.25; Figure 4e), as well as with Spartina above-ground biomass (p = 0.02, R2 = 0.27; Figure 4f).
We found significant independent effects of garden site and source site on Spartina morphology and production, suggesting that both short-term and long-term exposure to nutrient enrichment affect plant responses. As expected, Spartina originating from the enriched site performed relatively poorly when planted in unenriched gardens, whereas contrary to our prediction, Spartina originating from the unenriched site performed similarly across gardens. Our experimental design controlled for potential differences in genotypic richness and identity between sites, which can strongly influence plant traits and overall production (Hughes, 2014; Hughes et al., 2014; Proffitt et al., 2003, 2005; Zerebecki et al., 2017), as well as bacterial community structure (Bowen et al., 2017). In fact, we found significant effects of Spartina genotype on almost all response variables. The presence of such strong variation across genotypes in this system may have inhibited detection of nutrient enrichment effects on short-form Spartina in prior TIDE studies (e.g. Johnson et al., 2016), reinforcing that consideration of genetic variation can be important for both field and laboratory studies (Milcu et al., 2018).
Despite the relatively moderate levels of eutrophication in our study (Deegan et al., 2007) and the fact that only 30% of high tides flood the high marsh (Johnson et al., 2016), plants in enriched gardens had greater stem height in both years of the experiment, and higher stem density and above-ground biomass in the second year of the experiment, compared to plants in unenriched gardens (Figure S4.1). Thus, our results match findings of positive effects of nutrient enrichment on Spartina above-ground production (Deegan et al., 2007; Levine et al., 1998; McFarlin et al., 2008; Valiela et al., 1978) and plant production more generally (Elser et al., 2007). Interestingly, only below-ground allocation—not total below-ground biomass—differed between garden sites. Plants in enriched gardens had lower root to rhizome ratios (Figure 2e), consistent with observed patterns in terrestrial plants exposed to elevated nutrients (Tilman & Wedin, 1991) and in other marsh nutrient enrichment studies (Wigand et al., 2015). These responses suggest decreased investment in nutrient acquisition (roots) and increased investment in storage and vegetative growth (rhizomes) in enriched gardens. Furthermore, the reduced rhizome biomass of plants from enriched sites in unenriched gardens (Figure 2c) may be due to decreased nutrient uptake efficiency as a result of long-term exposure to a nutrient rich environment (Tilman & Wedin, 1991). Alternatively, this may be due to observed changes in Shannon diversity and/or composition of the associated microbial community (e.g. Dean et al., 2014): in our study, diversity of Spartina-associated soil microbial communities was higher in unenriched than enriched gardens after the first growing season, and decreased across gardens by the end of the experiment, similar to the effects of nutrient enrichment on alpine tundra plants and microbes (Dean et al., 2014). In salt marshes, the effects of fertilization on below-ground biomass are equivocal (Darby & Turner, 2008; Graham & Mendelssohn, 2016); our results suggest that whether below-ground biomass increases (Dai & Wiegert, 1996; Smart & Barko, 1978), decreases (Deegan et al., 2012; Turner et al., 2009) or remains the same (Anisfeld & Hill, 2012; this study) in response to nutrient enrichment may depend on differences in plant allocation to roots and rhizomes (Tilman & Wedin, 1991; Valiela et al., 1976).
While many studies report a positive relationship between fertilization and plant nitrogen content (Yuan & Chen, 2015), lower leaf tissue C and N content in enriched gardens in our study may be due to differences in allocation to above- and below-ground production that can in turn affect root and shoot composition (Mack et al., 2004). Alternatively, greater above-ground production may result in an overall increase in the total pool of above-ground C with enrichment, as in this study, despite a decrease in per leaf percent composition. A decrease in nutrient use efficiency and/or resorption rate in response to long-term fertilization may also impact percent and/or total C and N (Vitousek, 1982; Yuan & Chen, 2015). Correspondingly, we found that plants that had experienced elevated nutrient conditions for 10 years had consistently lower C and N pools than plants that had experienced ambient conditions (Figure 3d,e), similar to the effects of 10 years of fertilization on C and N pools of plant communities in the arctic tundra (Mack et al., 2004). Interestingly, while nutrient levels in this experiment were ‘only’ elevated to moderately high levels (15-fold and fivefold increase in and respectively; Deegan et al., 2012), plants with unique population histories (i.e. from different source sites) still exhibited distinct traits across garden sites. The persistent effects of population history on plant phenotype in our study complement emerging evidence that nutrient availability can alter life-history traits (e.g. elevated nutrients can relax selection on typical life-history trade-offs such as growth versus reproduction and offspring quantity versus quality; Snell-Rood et al., 2010, 2015; Wilson et al., 2006), with consequent effects including altered genetic variation within populations and/or accelerated evolutionary processes among populations (e.g. population divergence; Snell-Rood et al., 2015).
Nutrient enrichment can also alter soil microbial communities (Bulseco et al., 2019; Graves et al., 2016; Kearns et al., 2016; Ramirez et al., 2010, 2012), which in turn affect microbial respiration rates, plant–microbial interactions, and key biogeochemical pathways. Interestingly, microbial community composition in the greenhouse (prior to the field experiment) varied by source site, despite the fact that plants had been in a common garden for 1 year (Figure 4b). Given that Spartina genotypes were collected up to 500 m apart and on both sides of the creeks, it is unlikely that this effect is solely due to local-scale environmental heterogeneity. Instead, it could be the result of long-term nutrient enrichment. This pattern is similar to differences between bacterial and fungal communities associated with a terrestrial plant species that had evolved for multiple generations in non-drought versus drought conditions (terHorst et al., 2014), suggesting that population history has legacy effects on plant conditioning of soil communities for diverse species and different environmental stressors. We also found substantial variation in microbial community composition among Spartina genotypes in the greenhouse, which is consistent with studies of Phragmites australis (a perennial grass; Bowen et al., 2017), Arabidopsis thaliana (an annual flowering plant; Micallef et al., 2009), Boechera stricta (a perennial mustard; Wagner et al., 2016), Populus spp. (deciduous tree species; Madritch & Lindroth, 2011; Schweitzer et al., 2008) and various cultivars (Aira et al., 2010; Edwards et al., 2015; Mahoney et al., 2017).
However, the effects of Spartina genotypic identity and population history (i.e. source site) on microbial responses were quickly overwhelmed by the local environment once the plants were transplanted to the field, which is consistent with local edaphic conditions strongly influencing microbial community assembly (Fierer & Jackson, 2006). While these results contrast a study of the terrestrial plant Brassica rapa that found persistent interactive effects of evolutionary history and ecological context on soil microbes (terHorst et al., 2014), a key difference is the use of mesocosms (their study) versus field sites (our study) for the reciprocal transplant, suggesting that the relative importance and potential interaction of past population/evolutionary history and current ecological setting for plant–microbe interactions merits further investigation, especially in field reciprocal transplants. In our study, there was a strong temporal component to microbial community composition in the field, complementing prior studies documenting the importance of seasonality in structuring bacterial communities (Crump et al., 2003; Gilbert et al., 2010; Kearns et al., 2016). There was also a significant effect of garden site at the end of the experiment (Figure 4c), providing further evidence that nutrient enrichment may be influencing microbial communities in the high marsh. Finally, we found lower Shannon diversity in the microbial community of enriched gardens compared to unenriched gardens during the first growing season, similar to other studies documenting a decrease in microbial diversity at fertilized sites in a variety of systems (Dai et al., 2018; Kearns et al., 2016; Zhang et al., 2017). The effects of enrichment on the overall microbial community in this experiment are in contrast to results previously reported from the TIDE project (Bowen et al., 2011; Kearns et al., 2016) that showed some resistance to perturbation in the microbial community. This apparent contradiction may be explained by the fact that those studies did not account for genetic variation within the plant populations, and/or that they did not examine short-form Spartina. The more strongly reducing conditions, lower redox potential and higher sulphide concentrations typically associated with high marsh sediment and short-form Spartina (DeLaune et al., 1983) may amplify the importance of the elevated nitrogen (which was added as NaNO3) as an electron acceptor (Bowen et al., 2020), leading to a stronger response of the associated microbial community to nutrient enrichment. Although we had insufficient replication among genotypes to robustly test the Spartina genotype × nutrient enrichment interaction, the variance in the microbial community along the first principle coordinate was always lower in enriched communities (both origin and garden) than in unenriched communities. This is consistent with prior results indicating that nutrient additions result in homogenization of the active microbial community that erases effects of both season and habitat (Kearns et al., 2016) and suggests that while there may be an effect of genotype on the microbial community in the unenriched garden, that effect is minimized with the addition of nutrients.
Effects of nutrient enrichment on plant production and microbial community responses occurred in the same timeframe (i.e. after only 2 years), providing further evidence that plant–soil feedbacks may be operating in this system. In fact, we found a strong association between microbial community structure (represented by both PCoA axes) and components of Spartina biomass and allocation at the end of the field reciprocal transplant (Figure 4d–f). Plant performance is integrally linked to microbial biomass, activity and community composition, so changes in associated microbial communities due to altered nutrient conditions may feedback to affect plant growth and reproduction (Dean et al., 2014; Suding et al., 2008). Conversely, plant phenotypic variation, particularly in below-ground traits, can impact associated microbial communities (Bowen et al., 2017; Micallef et al., 2009; Schweitzer et al., 2008) via effects on ecosystem processes such as nutrient cycling (Bardgett et al., 2013; Madritch & Lindroth, 2011; Sayer et al., 2017). It is not possible to disentangle the direct effects of nutrient enrichment on Spartina production and microbial community structure from the indirect effects of nutrient enrichment on plants and microbes mediated by feedbacks in our study. However, our results indicate that both the short- and long-term effects of nutrient enrichment on plant and microbial communities may be extensive, even in areas experiencing relatively moderate and infrequent exposure to elevated nutrients. Greater understanding of the direct and indirect effects of fertilization on plants and microbes may be particularly important for predicting the capacity of these communities to respond to changing environmental conditions in both the short and long term.
It is important to note that although we had multiple gardens at each site, our study only included one enriched site and one unenriched site—a common feature of many whole-ecosystem field experiments that yield valuable insights into different processes testable only at larger scales and assess relative effect strengths in a real-world context despite limited replication (Carpenter, 1998; Schindler, 1998). By measuring plant species richness and community composition as well as porewater salinity and sediment oxygen for multiple gardens at each site, we were able to assess finer scale variation within sites and assess additional factors that may differ between the enriched and unenriched sites. A comparison of garden-level characteristics did not identify any pronounced differences between sites (see Appendix S1 for details), which parallels the results of an in-depth comparison of these sites using a suite of characteristics measured prior to the start of nutrient enrichment in 2003, including temperature, salinity, plant community composition, and nutrient, phytoplankton and total suspended solid concentrations (Deegan et al., 2007). While there are additional abiotic and biotic characteristics that were not quantified in our experiment, a ubiquitous and consistent difference between these sites is nutrient availability, specifically and concentrations (Deegan et al., 2007, 2012; Johnson et al., 2016). Thus, while we acknowledge that nutrient enrichment may not be the only factor contributing to the patterns observed, we argue it is the most parsimonious explanation.
The higher ratio of above- to below-ground biomass at the enriched site after only 2 years of nutrient enrichment (Figure 2b) has important implications for marsh stability, indicating that the negative effects of nutrient enrichment on creek geomorphology (Deegan et al., 2012) and marsh elevation (Turner et al., 2009, but see Anisfeld & Hill, 2012) are not limited to the low marsh. Rather, enrichment impacts the capacity of the entire marsh to respond and adapt to sea level rise. Furthermore, the effects of nutrient enrichment on below-ground production may be exacerbated if the magnitude and/or duration of fertilization increases. Thus, while we observed increased growth below-ground to exploit unused resource space in the short term (e.g. greater allocation to rhizome production in enriched gardens), this may precede decreased investment in below-ground standing crop in the long-term (Graham & Mendelssohn, 2016). We predict that long-term effects of nutrient enrichment on total below-ground production in the high marsh may be even more pronounced than short-term effects, with both root and rhizome biomass decreasing relative to above-ground production—similar to patterns observed in the low marsh (Deegan et al., 2012).
How decreased below-ground production concomitant with fundamental shifts in microbial community composition in response to long-term nutrient enrichment (Kearns et al., 2016; Leff et al., 2015; Rinnan et al., 2007) impact biodiversity and ecosystem function and stability merits further investigation in terrestrial and estuarine systems, as well as in agriculture (Dai et al., 2018). Pronounced reductions in almost all responses of Spartina originating from the enriched site when planted in unenriched gardens suggest that the long-term effects of nutrient enrichment may also include decreased overall production of plants accustomed to high resource environments. Thus, population history—in this case, long-term nutrient enrichment—can strongly affect both plant production and associated microbial communities, which may differentially impact ecosystem function under future environmental conditions.
We thank L. Deegan and the Plum Island Ecosystems LTER for access to the TIDE project and field sites. We gratefully acknowledge D. Behringer, J. Bloomberg, A. Bulseco, S. Burrell, T. Davenport, S. Fischer, T. Fish, J. Gladstone, M. Greenwood, A. Halverstadt, A. Knott, J. Lesser, C. Lynum, M. Olney, J. Renner, F. Schenck, A. Weiss, M. Willert and R. Zerebecki for help in the field and greenhouse, and assistance processing samples in the laboratory. In addition, we thank T. Gouhier for statistical advice, and B. Silliman and two anonymous reviewers for helpful feedback that improved this manuscript. Funding was provided by the National Science Foundation (DEB-1556738 to A.R.H.). Funding for the base nutrient enrichment experiment came from numerous awards, including DEB-1353140 to J.L.B., as well as DEB-0924287, DEB-0923689, DEB-0213767, DEB-1354494, DEB-1719621 and DEB-1902712. Additional support was provided by the Plum Island Ecosystems LTER (NSF OCE-1673630, OCE-0423565, and OCE-1058747). This is contribution 417 from the Northeastern University Marine Science Center.
CONFLICT OF INTEREST
The authors have no conflict of interest to declare. A. Randall Hughes is an Associate Editor of Journal of Ecology, but took no part in the peer review and decision-making processes for this paper.
T.C.H. and A.R.H. conceived and designed the experiment and collected the plant data; J.L.B. and P.J.K. collected the microbial data; T.C.H., A.R.H. and J.L.B. analysed the data; T.C.H. wrote the first draft of the manuscript. All authors contributed to subsequent drafts and gave approval for publication.
The peer review history for this article is available at https://publons.com/publon/10.1111/1365-2745.13756.
DATA AVAILABILITY STATEMENT
Plant data are available from Northeastern University's Digital Repository Service: http://hdl.handle.net/2047/D20412423 (Hanley et al., 2021). Microbial data are available from NCBI’s Sequence Read Archive, accession number PRJNAs631747 (https://www.ncbi.nlm.nih.gov/bioproject/PRJNA631747/).
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