Illuminating protist diversity in pitcher plants and bromeliad tanks

Robin S Sleith; Laura A Katz

doi:10.1371/journal.pone.0270913

. 2022 Jul 27;17(7):e0270913. doi: 10.1371/journal.pone.0270913

Illuminating protist diversity in pitcher plants and bromeliad tanks

Robin S Sleith ^1,^*, Laura A Katz ^1,²

Editor: Theodore Raymond Muth³

PMCID: PMC9328516 PMID: 35895712

Abstract

Many species of plants have evolved structures called phytotelmata that store water and trap detritus and prey. These structures house diverse communities of organisms, the inquiline microbiome, that aids breakdown of litter and prey. The invertebrate and bacterial food webs in these systems are well characterized, but less is known about microbial eukaryotic community dynamics. In this study we focus on microbes in the SAR clade (Stramenopila, Alveolata, Rhizaria) inhabiting phytotelmata. Using small subunit rDNA amplicon sequencing from repeated temporal and geographic samples of wild and cultivated plants across the Northeast U.S.A., we demonstrate that communities are variable within and between host plant type. Across habitats, communities from tropical bromeliads grown in a single room of a greenhouse were nearly as heterogeneous as wild pitcher plants spread across hundreds of kilometers. At the scale of pitcher plants in a single bog, analyses of samples from three time points suggest that seasonality is a major driver of protist community structure, with variable spring communities transitioning to more homogeneous communities that resemble the surrounding habitat. Our results indicate that protist communities in phytotelmata are variable, likely due to stochastic founder events and colonization/competition dynamics, leading to tremendous heterogeneity in inquiline microeukaryotic communities.

Introduction

Microbial eukaryotes are critical and ubiquitous members of ecosystems and have profound impacts on natural and human systems [1–3]. These organisms are key components of food webs by serving in diverse roles such as autotrophs (i.e. diatoms, green algae), heterotrophs (i.e. most ciliates, amoebae) and mixotrophs (i.e. many dinoflagellates). Despite their tremendous biodiversity, only a limited number of molecular studies have focused on the biodiversity of plant-associated protist communities (e.g. [4–6]). Here we use a large-scale amplicon sequencing approach to target SAR (Stramenopila, Alveolata, Rhizaria) diversity in phytotelmata (i.e. water filled plant cavities). One of the largest clades of microbial eukaryotes, SAR (Stramenopila, Alveolata, Rhizaria) contains ~3/5ths of eukaryotic diversity (reviewed in [7]). While this clade includes well-studied model organisms such as Plasmodium and Tetrahymena, the diversity and ecology of much of this group remains to be discovered. Members of the SAR lineages are globally distributed, and large-scale marine sequencing projects have greatly expanded our understanding of the ecology of these organisms (e.g. [8]). Much less is known about SAR in freshwater habitats, which has been identified as an area in particular need of additional research [7].

Phytotelmata have evolved for a variety of functions including supplementing nutrients for plants growing in nutrient-poor environments (e.g. the pitcher plants and tank forming bromeliads that are the focus of this work). Two genera of pitcher plants, Sarracenia and Nepenthes, have evolved modified leaves that help them to survive in low nutrient bogs in North America and epiphytic habitats in Southeast Asia, respectively [9]. Species of Nepenthes (also known as tropical pitcher plants) secrete fluid and digestive enzymes that fill their traps whereas the traps of Sarracenia species fill with rainwater [10]. Epiphytic bromeliads are native to the New World, with a high diversity of species and life-history traits [11]. Tank-forming bromeliads are epiphytic bromeliad species adapted to store water and break down leaf litter and trapped organisms [4].

Pitcher plant and bromeliad phytotelmata, the focus of our study, have associated microbiomes that aid in the breakdown of captured prey and detritus [10, 12]. Initial investigations of the communities in phytotelmata of the purple pitcher plant (Sarracenia purpurea) demonstrate that these plants maintain a distinct food web where arthropods, protists, and bacteria interact to break down trapped animals/materials into host-available nutrients [10, 13]. A history of intense study spanning nearly three decades has established Sarracenia purpurea in particular as a model system for experimental ecology [12].

Much work has been done to investigate the bacterial and animal members of pitcher plant and bromeliad associated communities. The bacterial communities within phytotelmata of both Sarracenia and Nepenthes pitchers are heterogeneous in composition but distinct from surrounding aquatic or terrestrial communities [5]. Studies have shown that these bacterial communities have similar core functions (e.g., chitin degradation, protease and other extracellular enzyme production, amino acid metabolism) across populations; these functions are thought to contribute to a mutualistic relationship between plant and inquiline community [6, 14, 15]. The relationship between the bacterial community and the plant has furthermore been shown to be mediated by the larvae of the pitcher-plant mosquito (Wyeomyia smithii), which prey on bactivorous protists, thus increasing bacterial decomposition [13].

Studies of bacterial and animal communities in bromeliads detect communities distinct from surrounding habitats, with highly variable composition within and between host species [16–20]. In both the purple pitcher plant and bromeliad phytotelmata, mosquito larvae have been identified as a major controlling factor in food webs [21]. Gene sequencing and metatranscriptomic approaches have revealed genes associated with chitinases, nitrogenases, and methanogenesis, indicating that these habitats may be important for bacteria involved in global carbon and nitrogen cycling [17, 22].

In contrast to studies of bacteria and animals, few studies focus on the microbial eukaryotic communities in pitcher plants and bromeliads, though fungi have been the subject of several studies [23, 24]. Studies relying on morphological identification of protists have determined that their major role in these systems is as bactivores [25]. Stoichiometric models of these systems show that the relationship between the protists and the plant host can be parasitic [26], although there are conflicting accounts in the literature [27]. Previous studies of protists in phytotelmata typically relied on morphological identification (e.g. [28]), therefore focusing on a handful of identifiable protists and missing a vast portion of cryptic or unidentifiable diversity. The diversity of ciliates within tank forming bromeliads has been extensively studied, as these environments harbor high levels of rare and endemic species [4, 29, 30].

The few studies based on molecular techniques uncover interesting patterns that may further our understanding of the relationship between microbes (eukaryotic and prokaryotic) and the host plant. Satler et al. [31] found that the pattern of genetic diversity of eukaryotic inquiline community members was similar to the pattern of genetic diversity of the host plant (Sarracenia alata), indicating an evolutionary link (co-diversification) between plant and associated microbial community. Bittleston et al. [5] used an amplicon approach with “universal” eukaryotic primers to demonstrate that communities in two highly diverged pitcher plant genera were strikingly similar, and that pitcher plant communities represent a distinct subset of the community found in the background habitat. Simão et al. [4, 32] used a different primer set and found that ciliates were the most abundant protists in bromeliad phytotelmata. These approaches are advantageous in that a large portion of eukaryotic diversity may be captured, but problematic as members of certain eukaryotic lineages (e.g. Diptera) can swamp the signal of less dominant taxa in a sample and these ‘universal’ primers inevitably miss many lineages. Our taxon-focused approach here, an analysis of the SAR clade that represents a tremendous diversity of eukaryotes (Grattepanche et al. 2018), allows us to narrow our focus to comparisons of protist diversity between communities in space and time, and across ecological gradients. This focused approach is needed as few studies examine protist interactions with plants and even fewer draw observations from the natural environment [33].

In this study, we investigate the diversity of plant associated eukaryotic microbiomes to understand the community assemblage patterns in discrete freshwater ecosystems. We use an amplicon sequencing approach targeting the small ribosomal subunit, and we generate amplicons from RNA extractions to target active community members. In Study 1, we sample broadly across plant/phytotelmata type and landscape to ask how host plant type influences microbial community structure and diversity. This sampling spans natural and built environments, including cultivated tropical pitcher plants and bromeliads in the Lyman Conservatory at the Botanic Garden of Smith College and wild purple pitcher plants from bogs across the Northeast U.S.A. We hypothesize that samples from natural environments will host more species, and that communities will be structured by geography. In Study 2, we sample the same wild purple pitchers three times at a single site over the spring growing season to examine community stability and how seasonal changes lead to shifts in phytotelmata communities. We hypothesize that communities will diverge as the spring season progresses and colonization and dispersal increase.

Materials and methods

Sample collection—Study 1

For Study 1 we examined the influence of plant host on SAR diversity and community structure. Phytotelmata were sampled from three geographic locations across the Northeast, two in Massachusetts: Hawley Bog and the Lyman Conservatory, and one in Maine: Big Heath (see Table 1 for further locality information, permission was obtained at all sites prior to sampling). We sampled two dates in the Lyman Conservatory, repeating the same plants on the second date (3/21). We sampled Hawley Bog twice for a total of 17 samples (separate plants from Study 2) and Big Heath once for a total of 10 samples for Study 1 and sampled Hawley Bog three times for Study 2 (below). The contents of phytotelmata (up to 25mL) were collected using a sterile transfer pipette and either placed in a sterile tube or filtered immediately following the methods described below for Study 2. To sample the biofilm community present in the phytotelmata of pitcher plants and bromeliads we swabbed the interior wall of each with a sterile cotton applicator and swirled the applicator with the collected fluid to mix. Samples collected in tubes were returned to the lab and concentrated on 2μm filters using vacuum filtration. Each filter was placed in 500μl of RLT buffer (Qiagen) and stored at -80°C.

Table 1. Study and sampling information.

Study	Host	Species	Samples	Geography	Dates (all samples from 2019 Month/Day)
1—Host	Bromeliad	Aechmea "Red Ribbon"	3	Lyman Conservatory, MA	2/5; 3/21
1—Host	Bromeliad	Nidularium innocentii	3	Lyman Conservatory, MA	2/5; 3/21
1—Host	Bromeliad	Vriesea fosteriana	3	Lyman Conservatory, MA	2/5; 3/21
1—Host	Bromeliad	Vriesea splendens	3	Lyman Conservatory, MA	2/5; 3/21
1—Host	Tropical Pitcher Plant	Nepenthes maxima	4	Lyman Conservatory, MA	2/5; 3/21
1—Host	Tropical Pitcher Plant	Nepenthes truncata	5	Lyman Conservatory, MA	2/5; 3/21
1—Host	Purple Pitcher Plant	Sarracenia purpurea	17	Hawley Bog, MA	5/16; 6/27
1—Host	Purple Pitcher Plant	Sarracenia purpurea	10	Big Heath, Acadia National Park, ME	5/21
1—Host + 2—Seasonal	Purple Pitcher Plant	Sarracenia purpurea	27	Hawley Bog, MA	5/6 ("Day 1"); 5/29 ("Day 24"); 6/27 ("Day 53")
1—Host + 2—Seasonal	Background Water	NA	9	Hawley Bog, MA	5/6 ("Day 1"); 5/29 ("Day 24"); 6/27 ("Day 53")

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Sample collection—Study 2

For Study 2 we examined how SAR diversity and community structure changes in Sarracenia pitchers over the course of the growing season at Hawley Bog, sampling nine pitchers three times from early May to late June. Pitchers at Hawley Bog emerge throughout the growing season (April—October), and bloom in late May to early June typically after their fifth growing season [12]. At this site, May and June receive an average of 11 cm and 9.5 cm of rain, respectively [34]. For repeated sampling, three rosettes of adult pitchers (likely including some from the previous growing season) were identified at the beginning, middle, and end of an established 26m transect extending from the forest edge to the center of the bog, closest to open water [35]. Three pitchers on each rosette were labelled using permanent marker and sampled on May 6th, May 29th, and June 27th 2019. The diameter of each pitcher opening was measured using calipers to understand the influence of pitcher size and morphology on the SAR community. For each sample, up to 25mL of fluid was removed using a sterile transfer pipette, volume was recorded, and fluid was concentrated on 2μm filters using a syringe and Swinnex Filter Holder. To sample the biofilm community present in the pitchers we swabbed the interior wall of each pitcher with a sterile cotton applicator and swirled the applicator with the collected fluid to mix. The filtrate was replaced in the pitcher to avoid pitchers desiccating prematurely, though we recognize that the amount of fluid in these pitchers varies tremendously throughout the growing season depending on rain levels, heat, cloudiness, and other weather factors. Each filter was placed in 500μl of RLT buffer (Qiagen) and stored at -80°C upon return from the field. To compare pitcher SAR communities to the background communities of the bog, 25mL of water below each rosette was collected and filtered using the methods described above.

Sample processing—Studies 1 and 2

RNA was extracted using the Qiagen RNeasy kit. We removed DNA from the extracted RNA with the TURBO DNA-free™ Kit (Invitrogen, CA, USA), and generated single-strand cDNA using the SuperScript® III First-Strand Synthesis System (Invitrogen) with random hexamer primers (Thermofisher, USA) following the methods of Sisson et al. [36].

We followed the methods of Sisson et al. [36] to generate amplicon libraries from the cDNA using SAR specific primers targeting the V3 region of the small ribosomal subunit (SSU-rRNA). Each PCR was conducted in triplicate then pooled to reduce PCR bias [37, 38]. The University of Rhode Island prepared sequencing libraries from amplicons and performed paired end (2x300bp) Illumina MiSeq High-Throughput Sequencing.

Data analysis

Sequence reads were analyzed following the methods of Sisson et al. [36] with scripts available at https://github.com/jeandavidgrattepanche/Amplicon_MiSeq_pipeline. This pipeline quality filters and merges reads using PEAR [39], builds OTUs with SWARM v2 with the parameter d = 1 [40], and removes non-SAR ‘outgroup’ sequences using a phylogenetic approach in which OTUs are added to alignments based on full length reference sequences from GenBank (though the SAR primers target this group, a small proportion of non-SAR sequences are amplified due to the conserved nature of the SSU-rRNA). We subsampled each amplicon library to 5,000 reads and only included libraries with greater than 2,900 reads in subsequent analyses. We removed OTUs that had fewer than 100 reads in a single sample. The phylogenetic tree was created using RAxML [41] from a Mafft alignment [42] on the CIPRES Science Gateway [43]. We calculated dissimilarity matrices with UniFrac distances [44], both weighted (relative abundance) and unweighted (presence/absence), and performed principal coordinate analysis using R packages phyloseq and vegan [45–47]. Permutational multivariate analyses of variance (function adonis in vegan) were used to test for differences in community composition driven by host plant and sample date. Rarefaction curves were generated using the methods of Hausmann et al. [48]. The network analysis was conducted in R using RAM: R for Amplicon-Sequencing-Based Microbial-Ecology [49].

Results

Amplicon analysis of phytotelmata (Studies 1 and 2)

Based on analyses of 84 samples (54 Sarracenia pitcher plants, 9 Nepenthes pitcher plants,12 bromeliads, 9 background water samples; see Table 1 for sample information), we found 463 OTUs (i.e. rDNA amplicons) represented by 420,116 reads that fall among SAR reference sequences using the phylogenetic approach described in Sisson et al. [36]. After removing samples with fewer than 2,900 reads and low abundance OTUs (i.e. fewer than 100 reads for a single sample, see methods), 135 OTUs represented by 391,044 reads in 81 samples remain for subsequent analyses (Fig 1, S2 File). Raw reads associated with this study are available under NCBI BioProject ID: PRJNA682436. The phylogenetic tree in Fig 1 shows that ciliates make up nearly half the 135 OTUs recovered in this study.

Fig 1 — Branches are colored by major clades, Rhizaria (Rhi, orange) Stramenopila (Str, green) and Alveolata (Alv, blue, with Ci indicating ciliates). The number of reads is indicated in the inner ring, followed by host plant specificity (>90% of reads from a given host), followed by geographic specificity (>90% of reads from a given location).

Study 1: The influence of plant host on OTU richness and community structure

Rarefaction analysis of observed OTUs and Shannon’s diversity index consistently rank samples from background bog water as more diverse than phytotelmata (Fig 2). We investigated whether plant type influences microbial community composition by comparing patterns across three distantly-related plant groups with convergent water trapping morphologies–bromeliads, Nepenthes, and Sarracenia. Although we detect the influence of host plant type on community structure (adonis: R² = 0.23, p = 0.001), we also observe tremendous heterogeneity among samples (Fig 2), with no clear geographic signal, counter to our hypothesis. Principal coordinate analysis based on weighted UNIFRAC distances shows overlap of communities in bromeliads and tropical pitcher plants growing in the Lyman Conservatory with those occurring in Sarracenia purpurea in two bogs across the Northeast U.S.A. (Fig 2). Communities from Sarracenia purpurea are most variable (i.e. occupy the most ordination space), followed by cultivated bromeliads and the pitcher plant Nepenthes. Although Sarracenia pitcher plants displayed the most heterogeneity, communities in bromeliad tanks located in a single room at the Lyman Conservatory occupy nearly as much ordination space (i.e. distance along PC1) as do communities occurring in Sarracenia purpurea in four bogs separated by hundreds of kilometers across the Northeast U.S.A. (Fig 2).

Fig 2 — A. OTU richness (rarefaction plots, error bars are standard deviations) and Shannon diversity (inset beanplots) is highest in samples from the background community (light blue), followed by samples collected from *Sarracenia* (purple), bromeliads (brown), and *Nepenthes* (green). B. Principal Coordinate Analysis based on weighted UNIFRAC demonstrates differentiation of communities by host plant, with tremendous heterogeneity among communities in field-collections from *Sarracenia* (purple dots) and among greenhouse bromeliads (brown triangles; all from the same room in the Lyman Conservatory). Details on site and OTUs can be found in S1 and S2 Files.

Study 2: The influence of season on OTU richness and community structure

We sampled nine pitchers three times at Hawley Bog over the course of the 2019 spring growing season (May 6th ("Day 1"); May 29th ("Day 24"); June 27th ("Day 53")), removing most but not all of the fluid each time. Analyses of the resulting amplicons show that the background samples consistently have more OTUs than the communities sampled in Sarracenia pitchers (S1 Fig). The highest richness estimated by rarefaction is in samples collected from the background water, with samples from the site closest to the forest edge consistently higher than the other sampling sites (i.e. rarefaction curve for site 1 background samples higher than all other curves in S1 Fig).

To determine whether pitcher volume or shape influenced OTU richness, we fit a linear model of observed OTU richness by pitcher diameter and volume (S2 Fig). We do not detect any pattern between the volume or the diameter of the top of pitchers and observed richness at Hawley Bog (p >0.7, S2 Fig). We find that the communities in Sarracenia purpurea at Hawley Bog are a subset of the bog water community as the network analysis shows connections between the OTUs occurring in pitchers and the background samples (Fig 3).

Fig 3 — Connection of OTUs (circles) shared (lines) between pitchers (purple squares) and bog water (blue square) for the rosette of pitchers at Hawley Bog closest to the forest edge, sampled on May 6th ("Day 1"). Taxonomy of OTUs is colored as Rhizaria (orange) Stramenopila (green) and Alveolata (blue).

We again use principle coordinate analysis to determine how time and local habitat influence protist communities. We detect a temporal pattern whereby samples collected in early and late May vary widely across ordination space, while samples collected in late June (Day 53) cluster together on the right side of PC1 (Fig 4A, adonis: R² = 0.18, p = 0.003). The samples collected in late June also fall close to the background bog water samples, indicating that by late June the SAR community in the pitcher plants resembles the community of the background habitat. The OTU ordination in Fig 4B demonstrates that Alveolate taxa are the most abundant in samples from the background and late June, while many rhizarian taxa are abundant in the earlier months (as weighted Unifrac takes relative abundance into account in our ordination).

Fig 4 — A. Principal Coordinate Analysis based on weighted UNIFRAC of communities from repeated sampling along a transect (26m from Forest Edge to Pool Edge) at Hawley Bog (May 6th ("Day 1", green); May 29th ("Day 24", purple); June 27th ("Day 53", yellow)) shows *Sarracenia* SAR communities in late June (Day 53, yellow filled symbols mostly appear on right side of PC1) clustering near the background community (all open symbols). B. Principal Coordinate Analysis based on weighted UNIFRAC of OTUs (same ordination space as 4A), showing that the OTUs that differentiate and correspond to the late June samples (the yellow, Day 53 samples on right side of PC1 in 4A) are primarily Alveolate OTUs (denoted in blue). Details on site and OTUs can be found in S1 and S2 Files.

We also detect temporal shifts in community structure at the highest taxonomic level (i.e. Stramenopila, Alveolata, Rhizaria), within Sarracenia pitchers. Over the three sampling times, OTUs identified as Stramenopila and Rhizaria decreased and OTUs identified as Alveolata increased across the spring season (Fig 5). The Alveolate OTUs with the largest increases across the growing season (OTU22, OTU25) have phylogenetic taxonomic assignment in the ciliate classes Colpodea and Oligohymenophorea, respectively (closest reference genera are Bardeliella and Tetrahymena, with full taxonomic information in S2 File, columns G and H). The Rhizarian OTU with the largest decrease (OTU10 in Figs 1 and 4) is in the Viridiraptoridae, a parasite of algae. The Stramenopile OTU with the largest decrease (OTU6 in Figs 1 and 4) is an autotrophic Chrysophyte. In the background community, OTU8, a ciliate most closely related to Leptopharynx was dominant.

Fig 5 — The changes in relative abundance of the OTUs from each sampling month separated by sampling type (background bog water (left) vs pitchers (right); OTUs from discussion labeled) shows that the relative abundance of OTUs assigned to Rhizaria declines sharply over the growing season in pitchers, though much less drastically in the background samples. At the same time, the relative abundance of alveolates increases. Stacked barplots represent relative abundance (denoted by bar segment size and shading).

Discussion

The main insights of this study include: 1) inquiline SAR communities contain fewer OTUs than nearby aquatic communities, irrespective of plant host 2) plant host influences SAR community structure, but with tremendous variation 3) at a single site, SAR communities in wild pitcher plants converge and resemble the background communities as the growing season progresses. Together these data highlight the complexity of microeukaryotic communities associated with plants.

Richness

Previous work investigating physiological and environmental drivers of protist richness in pitcher plants relied on morphological identification, which makes comparison with molecular studies difficult as richness may be underestimated in morphological studies [50]. In our study, richness measures indicate that in both bromeliads and pitcher plants (both in the field and greenhouse), the inquiline community typically represents only a fraction of the diversity found in nearby standing water (Fig 2). Previous studies have shown that the fluid of pitcher plants and bromeliads is a specialized environment with strong fluctuations of volume, temperature, pH, and chemical composition (reviewed in [10]). These fluctuations make the inquiline environment distinct from the surrounding habitats and likely explain the lower richness observed in the pitcher and bromeliad tanks. This pattern could also be due to limited dispersal of SAR community members into the modified leaves of pitcher plants and bromeliads. Previous work on fungi demonstrated that dispersal ability was a key driver of metacommunities in purple pitcher plants [24].

As we hypothesized, samples from wild pitchers had higher richness than samples from cultivated tropical pitcher plants and bromeliads growing in the Lyman Conservatory (Fig 2). The high richness in wild Sarracenia may result from the island-like nature of these pitchers and/or the variation in source populations. We recognize though that interpretation of patterns are confounded by the fact that we sample greenhouse and wild pitchers and different genera, although Bittleston et al. [5] used amplicon sequencing methods to show that Nepenthes and Sarracenia pitchers grown together developed similar bacterial and eukaryotic inquiline communities. The indoor habitat may have more limited access to protists compared to pitchers in wild habitats, which were sampled across sites in two states, potentially increasing the ability to detect diversity. We did not detect a pattern of increased richness with increased volume as others have ([28]; S2 Fig).

Study 1: The influence of plant host on community structure

To investigate how host plant characteristic influence protist communities, we sampled from a mix of cultivated and wild phytotelmata forming plants (Table 1). While we detected some differentiation among a subset of samples from bromeliad tanks (i.e. cluster of 7 brown triangles on left side of PC1; Fig 2), there was generally a lack of differentiation between communities from different host species, with samples from bromeliads, Sarracenia, and Nepenthes overlapping in ordination space (Fig 2). The extensive overlap in SAR communities among hosts could be due to a homogeneous species pool of potentially colonizing protists at the scale of northern New England (i.e. the same protists have equal chance of colonizing bromeliads or pitcher plants because the same protist species/propagules exist at each site). Comparisons of the variation within and between host species raise interesting questions. The variability of communities from Sarracenia purpurea from two sites across five dates is understandably high given the range of locations and dates (Fig 2), but contrasts with the almost equal variation detected from bromeliads growing in a single room (i.e. samples from bromeliads span a similar range along PC1 compared to samples from Sarracenia in Fig 2). This could be due to the species level morphological differences of tank forming bromeliads as differences in the shape of the tank likely contribute to dispersal patterns and water flow, perhaps leading to the observed variation in SAR communities. Previous work investigating eukaryotic and bacterial communities from wild bromeliads has demonstrated that extensive variation exists within and between bromeliad species [19, 30, 32], supporting our findings of variation in SAR communities within bromeliads found within 3 feet of one another in the Lyman Conservatory.

Varying source communities within the Lyman Conservatory are insufficient to explain the SAR biodiversity patterns in this study. Samples from the tropical pitcher plant genus Nepenthes grown in the Lyman Conservatory were distinct from bromeliad samples in our PCoA (Fig 2), indicating that within the Lyman Conservatory, host plant type influences SAR community structure, though some Nepenthes were grown in hanging baskets as opposed to shelf top bromeliad pots. This is also supported by the clustering of SAR communities within two species of Nepenthes (Fig 2). In a review of carnivorous pitcher plants, Adlassnig et al. [10] synthesize evidence demonstrating that the Nepenthes pitchers secrete digestive enzymes and actively change the pH of their fluid, which may lead to the lower diversity and homogeneity among samples from this genus that we observe.

Study 2: The influence of season on community structure

The microbial communities present in phytotelmata food webs are useful for understanding ecosystem processes such as colonization and community turnover, as each cavity/pitcher can be considered a unique microcosm that can be sampled at distinct levels including cavities/pitchers within plants and plants within populations [12, 28]. Similar to previous studies of eukaryotes [6], we detect high variability of eukaryotic microbial communities between pitchers on a single rosette and in a single bog. Repeated monitoring of communities over time in our study found that, counter to our original hypothesis, this variability decreased over the course of the summer in Hawley Bog (i.e. SAR communities within pitchers in late June were more homogeneous than samples in early May). This leads to a number of follow-up hypotheses: in early spring, stochastic colonization events from the background community, as well as differences in pitcher age, lead to diverse communities in pitchers but then as summer progresses, communities are homogenized through dispersal (e.g. rain/wind storms) and/or biotic interactions (e.g. competition, predation) or other ecological and environmental phenomena occur such that, by mid-summer the communities are similar in composition.

Previous research has shown a similar pattern for bacteria and protists where seasonal dynamics can lead to community homogenization over time [51–54]. Bacteria in the phyllosphere have clear seasonal patterns dependent on priority effects whereby the success of one early strain influences the outcome of the entire community [51, 55]. Although we remove fluid containing SAR community members throughout the course of the study, we left behind a portion of the community (in both the water column and biofilm), particularly in the tapering base of each pitcher that likely serves as a refugia for microbial species. Moreover, the stochastic nature of the system means that rainstorms, animal movement, and other factors influence the dispersal of organisms into and out of these systems, making our removal of fluid aligned with the natural processes influencing these communities. Furthermore, the communities in pitcher plants on Day 53 were quite similar to the background communities (Fig 4, yellow filled symbols are clustered with open filled background samples), rather than a novel community cluster representing experimental disturbance. Finally, several studies of the pitcher plant mosquito (Wyeomyia smithii) found that densities peaked from late July through September [56, 57] indicating that the phenology of animals in phytotelmata could influence the microbial community temporally. Nevertheless, we recognize that our perturbations may have influenced the observed community changes over time.

For microbial eukaryotes, a competition-colonization model suggests that smaller species will increase population sizes quickly and colonize habitats more rapidly [58–60]. Pitchers in early spring may be dominated by smaller species: indeed, we see Cercozoa (e.g. OTU10, OTU18) dominating, which are likely small flagellate lineages, while larger ciliate species dominate in summer and background samples. Furthermore, as competition increases over the summer, this may lead to higher dispersal that in turn could lead to the homogenizing effect we observe.

Conclusions

In conclusion, our study demonstrates that host plant type does not always strongly influence phytotelmata protist communities as we find considerable variation in SAR communities within bromeliads, Sarracenia purpurea, and Nepenthes pitcher plants. We also detect tremendous heterogeneity across space and time, reflecting the stochastic nature of phytotelmata environments. Within a single site over the course of the spring growing season, seasonal dynamics may decrease heterogeneity and point to the importance of repeated temporal sampling. These findings indicate that protists are an integral part of phytotelmata communities and mirror ecosystem level phenomena such as seasonality and species turnover.

Supporting information

S1 Fig. Rarefaction analysis of samples from Hawley Bog (where repeated sampling occurred) comparing communities from three pitchers on a single rosette at each location in the bog, and background samples below each rosette.

Communities from background water samples (open symbols) are consistently more species rich than from Sarracenia pitchers at Hawley Bog. The background water at the forest edge (light green) is consistently more species rich than the mid bog sites (pink and light blue) along our transect. Error bars are standard deviations.

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S2 Fig. Observed richness (number of OTUs) by pitcher opening diameter and pitcher volume shows no relationship between richness and pitcher morphology.

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S3 Fig. The PCoA from Fig 2 with 95% data ellipses representing the multivariate t-distribution (solid) and multivariate normal distribution (dashed).

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S1 File. Sample SRA and collection detail information.

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Click here for additional data file.^{(11.4KB, csv)}

S2 File. OTU table with phylogenetic taxonomic assignment and read counts per OTU.

(CSV)

Click here for additional data file.^{(31.3KB, csv)}

Acknowledgments

We are grateful to Ayla Say, Berry Williams, Chase Macpherson, Chrisshara Robinson, Lucinda DeBolt, and Tiannettie McKee for fieldwork and lab assistance. We thank Rabindra Thakur and members of the Katzlab for feedback on early drafts. We extend our appreciation to Janet Atoyan at the University of Rhode Island for running the MiSeq sequencing of our samples.

Data Availability

Raw reads associated with this study are available under NCBI BioProject ID: PRJNA682436.

Funding Statement

This work is supported by an NSF grant DEB-1541511 to Laura A. Katz. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

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PLoS One. doi: 10.1371/journal.pone.0270913.r001

Decision Letter 0

Theodore Raymond Muth

13 Jan 2022

PONE-D-21-40078Illuminating Protist Diversity in Pitcher Plants and Bromeliad TanksPLOS ONE

Dear Dr. Sleith,

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Reviewer #1: This is a straightforward report of two small-scale studies analyzing the composition of protist communities in pitcher plant tanks, both in different species and different timepoints. It addresses a novel question, executed in a generally technically sound manner, readable, and is a good fit for PLoS ONE. All of my suggestions could feasibly be addressed without additional wetlab experimentation.

One addition that could improve the paper is some additional information within the main text figures on the identity of the OTUs of interest. The introduction indicates that ciliates are dominant in other studies of bromeliad protist communities using different methods, and that bromeliads harbor rare species, but universal primers miss a lot of diversity that SAR primers could capture- three interesting hypotheses. There isn’t much detail given on the taxonomic resolution of these primers, but the final results paragraph and previous Sisson study cited indicates that the sequences are identifiable to some more specific taxonomic level. This information is not necessary for all the OTUs, but would be helpful to add to the dominant OTU highlighted in Fig. 4 and the specialized OTUs shown in figure 5, to put those initial questions into context. Furthermore, several recent studies of protists living on other types of aerial plant tissues have indicated the presence of a large diversity of Cercozoa esp. dominated by Sarcomonads (Ploch et al. 2016, Flues et al. 2018, Walden et al. 2021, Sun et al. 2021), but a few Ciliophora (Colpodea) have been reported to be universal too (Muller and Muller 1970). Some additional OTU identity could help contextualize which patterns you are observing are common to other plant structures vs. novel or specialized to bromeliads. Given the acidity, one would expect the residents to be very different from that of other plants, and the paragraph starting on line 278 indicates this could be true! Paper dois below:

10.1111/jeu.12314

https://doi.org/10.1111/jeu.12503

https://doi.org/10.1111/1462-2920.15385

https://doi.org/10.2307/2423721

https://doi.org/10.1093/femsec/fiab081

The introduction talks a lot about pitcher plant microbiomes, but could benefit from a little more general information on the pitcher plant environments being studied, to be accessible to researchers not in this specific area. Are the tanks thought to be rain-fed or does the water mainly seep in from the surrounding bog? What type of plant exudates feed the microbiome (i.e., sugars, lipids, insecticidal enzymes, acidity?). When one plant has multiple pitchers, do they usually emerge at the same time or do they have different ages? Is the type and amount of insect prey known to affect the microbiome? Finally, what are the phenotypic differences among the species being studied here (volume, habitat, shape, prey range, lids vs. no lids)? This information would help rationalize your hypotheses about species effects and seasonality.

Also in the introduction, there are some references to there being more studies of bacteria and animal communities than of microbial eukaryotes (line 113), meaning protists, but fungal studies are only mentioned in the discussion. Fungi are technically microbial eukaryotes, so please clarify when you are talking about only protist communities.

Line 126- the genetic diversity of the microbial community was similar to the host plant? Unclear phrasing.

-Line 145 “differentiate active community members from cysts” makes it sound like both parts of the community were measured and compared, it would be better rephrased to say the method primarily targets active protists.

Methods:

-Please provide more information about the sequencing beyond that it was MiSeq. Did you sequence paired end reads? 2x300 or 2x150 length? How many total reads? Were reads filtered for quality or length before entering them in the pipeline? Even if some of the QC is in the cited link, it’s best to say briefly what was done.

-line 201 – meaning of “OTU libraries” is unclear.

In figure 3A it would be helpful to indicate somewhere in the figure or legend that open symbols are background samples. Circles indicating confidence intervals and p-values would greatly help demonstrate the finding that Day 53 communities are significantly different.

I really do not understand what Figure 3B is showing. PCA is typically used to differentiate different sample types, but these are all from the same treatment. If the point is that most OTUs are alveolate at this timepoint, isn’t that already shown in figure 4, and what are the principal components for?

Figure 5 is discussed before Figure 3. Figures should be in the order they are discussed in the text.

There are some places in the results where the verb tense suddenly changes from past to present tense (such as line 268). Please check for verb tense consistency.

I would suggest limiting the discussion to be no longer than the length of the results text, which should be enough to synthesize how the results match initial expectations stated in the introduction and extend upon previous work. The recent Walden et al. paper on phenology of protists in aerial plant tissues may be good to mention for context. Also it seems that the findings here (seasonal successional shifts, origin from surrounding environmental microbiota, and specialized/reduced diversity from surrounding microbiota) align well with other emerging paradigms from the leaf and floral ecology field, which may be worth a mention (good overview by Britt Koskella here: https://doi.org/10.1016/j.cub.2020.07.037)

Reviewer #2: In their manuscript “Illuminating Protist Diversity in Pitcher Plants and Bromeliad Tanks”, the authors present a study that used a molecular approach to characterizing eukaryotic microbes (specifically SAR protists) in pitcher plants (Sarracenia and Nepenthes) and tank bromeliads. In the study, the authors sampled inquiline communities from their respective hosts over varying spatial and temporal scales. They found that host plant species affects SAR community structure and that there is a large amount of variability between communities within the same host plant. They also found that Sarracenia pitcher plants from the same bog are more diverse and heterogenous earlier in the growing season than later in the growing season. I enjoyed reading this manuscript immensely. The methods are rigorous and stand out from previous work that characterizes protists in phytotelma primarily based on morphology. Overall, I believe that the manuscript has merit for publication following a round of revisions from the authors. I have made some specific suggestions below.

Thank you for a fun read,

Erica Holdridge

NSF Postdoctoral Research Fellow

Boise State University

1. Although the authors have provided a nice table (Table 1), I still have a hard time understanding the sampling structure of their study. This could use quite a bit of clarification in the text of the manuscript. In particular, there are 3 samples from each bromeliad species taken across 2 dates. Are these from 3 different plants? How many were sampled on each date? etc. The same questions apply for the Nepenthes sampled. There were also 17 Sarracenia samples from Hawley Bog separately from the temporal study. Were these from a separate part of the bog than the Sarracenia that were used in the temporal study?

2. I can’t help but wonder if your Sarracenia samples are so variable (e.g. Figure 2B) because (a) you simply have more samples taken over a wider range of time points and (b) they’re taken from the field rather than a greenhouse.

3. Lines 341-346: Are Nepenthes at Lyman Conservatory grown in hanging baskets? If so, this may be one reason why bromeliads and Nepenthes from the same greenhouse have distinct communities (although I do believe that most of it does have to do with host identity).

4. Do you notice more mosquitoes (particularly W. smithii) in Hawley bog later in the growing season? It’s possible that they are aiding in dispersal and could explain some of the homogeneity you see later in the growing season. You do mention “animal movement” on lines 371-372, which may be a reference to this idea.

Minor comments:

• lines 77-78: “Much less is known about freshwater habitats” to “Much less is known about SAR in freshwater habitats”

• lines 92-93: “host available” to “host-available”

• line 163: “was collected” to “were collected”

• line 203: “SWARM v2 d = 1” to “SWARM v2 with the parameter d = 1”

• line 212: “weighted and unweighted (for relative abundance)” to “weighted (relative abundance) and unweighted (presence/absence)”

• lines 231-232: How many OTUs exactly?

• lines 253-255: The first sentence of this section is a fragment.

• line 267: You discuss Figure 5 before Figure 4; they should probably be reordered so they are organized by the order in which they are discussed

• line 286: OTU8 seems to be most abundant in the background bog water (Figure 4) – what is the taxonomic assignment of that OUT?

• line 342: This is the first place where you clarify that you mean Nepenthes when you say “tropical pitcher plants”. I know that this is the common name but other readers may not, so this should be clarified earlier in the manuscript.

• line 355: This would be a great place to site Aaron Ellison and Nick Gotelli’s new book “Scaling Ecology with a Model System” (2021)

Reviewer #3: The manuscript by Sleigh and Katz investigates the eukaryotic diversity within the inquiline communities of pitcher plants by using a taxon-focused approach. The technique they used is very timely and needed because eukaryotic diversity is understudied in general and especially in pitcher plant systems. When eukaryotic diversity is studied, it is usually with broad primers that can underestimate our knowledge about protist diversity because of the overabundance of insect DNA that can also be captured. Overall, the manuscript was very well written and there was a very indepth literature search. My main comments are that more information is needed in the methods section, and I am not fully comfortable with considering this a study that addresses Succession. With most of the inquiline community removed at each sampling point, this study seems to address more the resilience of the community after a disturbance, and the role of stochastic events and dispersal, instead of internal biotic interactions, on protist diversity throughout time, than a proper assessment of Succession. The study did definitely investigate the difference in protist composition across the different locations in the bog, but their question about Succession may need to be re-thought.

Main comments in Method Section:

In the Methods section of Study 2, additional information is needed in order for a better understanding of what was done. Specifically,

1) Were new pitchers selected on each of the rosettes at the beginning of the field season? This would allow for an accurate assessment of early succession.

2) It is unclear in the Methods if the three pitchers were sampled on each of the three sampling dates, or if one pitcher represented a specific sampling date and the replicate was on the other two rosettes in the section (i.e., Pitcher 1 = May 6, Pitcher 2 = May 29, Pitcher 3 = June 27). This information is stated in the Results section (Line 253-255) and should be moved up to the Methods for clarity.

3) For a better assessment of the methods and results, it would also be helpful to know how much volume was in each pitcher during each sampling time and the distance between each plant in Forest edge versus open bog, for example.

Main comment regarding Succession:

On Line 179-181, the authors state that a maximum of 25mL of fluid was removed and fluid was concentrated onto filters. If each pitcher was sampled three times (the three time periods), removing up to 25mL on each time period would be an exhaustive sampling, which would not capture protist succession through time if the community was removed almost completely during each sampling and needed to reset. To follow succession properly, it would be better to either exhaustively remove the volume of one pitcher on each of 3 replicate rosettes. This would represent one time period, replicated 3 times within 1 section of the bog, or, to take a smaller volume from each pitcher during the three sampling events to prevent a disturbance.

My concern about the large volume that was removed each time continues in Lines 361-365 of the Discussion. These follow-up questions are indeed important, but I just cannot see how biotic interactions will play a role in the authors’ sampling method if they removed most of the volume in the leaf each time. The internal biotic interactions were basically reset at each time point, making it instead a study that addressed the role of stochastic events and dispersal at different time points throughout the growing season, and a study that tests the resilience of the community after such a large disturbance each time. (I am happy to see though that the authors recognize this potentially large perturbation of removing almost all the volume during each sampling point (Line 374-375) and their results do indeed match what is found with microscopic counts of protists in the Sarracenia system, with flagellates dominating at the beginning of succession and ciliates being more abundant later in the season).

Additional comments:

- Why were OTUs used instead of ASVs?

- Figure 2. It would be helpful to visualize which of the Sarracenia samples came from which site.

- Figure 3A. Labelling the Date as a circle is confusing with the Forest Edge circle. Additionally, it is confused what “Pool Edge” means.

- Table 1. Minor detail, but signify that the dates are Month/Day.

**********

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Reviewer #2: Yes: Erica Holdridge

Reviewer #3: No

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PLoS One. 2022 Jul 27;17(7):e0270913. doi: 10.1371/journal.pone.0270913.r002

Author response to Decision Letter 0

7 Mar 2022

We thank the reviewers for their helpful comments, which we have addressed in detail below. We believe the manuscript is now substantially strengthened and more accessible to the broad readership.

****

Reviewer 1

REPLY: Thank you for the suggestion, we agree that we could do more to teach about the biology of the organisms. To this end, we have added some additional information to the results and point to the OTU table in the supplement which has all taxa identified to species with the corresponding GenBank number of the closest reference taxon in the tree-based taxonomy assignment.

REPLY: We have added additional information to the introduction that explains more pitcher biology and ecology. While we would love to address all of the questions raised here we believe this would be beyond the scope of the introduction to a paper investigating microbial communities and instead point to the literature and some helpful review papers.

REPLY: We have clarified where fungi fit into the phytotelmata literature.

Line 126- the genetic diversity of the microbial community was similar to the host plant? Unclear phrasing.

REPLY: We have reworded the sentence to clarify.

Line 145 “differentiate active community members from cysts” makes it sound like both parts of the community were measured and compared, it would be better rephrased to say the method primarily targets active protists.

REPLY: We have rephrased this sentence to more accurately reflect our methods.

Please provide more information about the sequencing beyond that it was MiSeq. Did you sequence paired end reads? 2x300 or 2x150 length? How many total reads? Were reads filtered for quality or length before entering them in the pipeline? Even if some of the QC is in the cited link, it’s best to say briefly what was done.

REPLY: We have added the requested information to the methods.

line 201 – meaning of “OTU libraries” is unclear.

REPLY: We have added language to clarify the sentence.

REPLY: The key of 3A indicates (background/rosette) for open vs. filled and we have added text to the legend. The addition of confidence intervals makes the figure too difficult to interpret but the Adonis test reported in the results demonstrates that the relationship is significant.

REPLY: This is the same PCoA as Figure 3B but instead of plotting samples, we are plotting the taxa (OTUs) in the same PCoA space, which demonstrates which taxa (or groups of taxa) are driving the differentiation of samples we see in 3A. We have added text to the legend that helps explain this better.

Figure 5 is discussed before Figure 3. Figures should be in the order they are discussed in the text.

REPLY: Apologies, we have fixed the figure order.

There are some places in the results where the verb tense suddenly changes from past to present tense (such as line 268). Please check for verb tense consistency.

REPLY: We have checked verb tense consistency.

REPLY: Thank you, we have received other feedback that requested more discussion, and will work with the editor to reconcile this. We appreciate the suggestions and have incorporated these references.

Reviewer 2

REPLY: We thank the reviewer for this question and we have added text to the methods clarifying the sampling design.

REPLY: In the discussion we mention the high variation of Sarracenia and expect this high variation given the samples are coming from different locations and dates. Also, these pitchers serve as islands in their environments, leading to high variability. What we did not expect was that the variation of bromeliads was nearly as great as Sarracenia. We have clarified wording in the discussion regarding this.

REPLY: Some Nepenthes are in hanging baskets and some are in pots on shelves at the same height as the bromeliads. This is a possible difference and we have added it to the discussion.

REPLY: Unfortunately, we do not have quantitative counts of mosquito larvae from these samples, but yes, this is a possibility. We have added a discussion of mosquito activity that incorporates this idea.

lines 77-78: “Much less is known about freshwater habitats” to “Much less is known about SAR in freshwater habitats”

REPLY: We have made this correction.

lines 92-93: “host available” to “host-available”

REPLY: We have made this correction.

line 163: “was collected” to “were collected”

REPLY: We have made this correction.

line 203: “SWARM v2 d = 1” to “SWARM v2 with the parameter d = 1”

REPLY: We have made this correction.

line 212: “weighted and unweighted (for relative abundance)” to “weighted (relative abundance) and unweighted (presence/absence)”

REPLY: We have made this correction.

lines 231-232: How many OTUs exactly?

REPLY: We have added OTU number to this sentence.

lines 253-255: The first sentence of this section is a fragment.

REPLY: We have made this correction.

line 267: You discuss Figure 5 before Figure 4; they should probably be reordered so they are organized by the order in which they are discussed

REPLY: We have fixed the figure order.

line 286: OTU8 seems to be most abundant in the background bog water (Figure 4) – what is the taxonomic assignment of that OUT?

REPLY: We have added information on OTU8 to the results.

line 342: This is the first place where you clarify that you mean Nepenthes when you say “tropical pitcher plants”. I know that this is the common name but other readers may not, so this should be clarified earlier in the manuscript.

REPLY: We have made this correction.

line 355: This would be a great place to site Aaron Ellison and Nick Gotelli’s new book “Scaling Ecology with a Model System” (2021)

REPLY: Thank you for the suggestion, we have added the reference

REVIEWER 3

REPLY: We appreciate the point regarding succession vs. disturbance and have further addressed the difficulty in untangling succession and disturbance in the discussion.

1) Were new pitchers selected on each of the rosettes at the beginning of the field season? This would allow for an accurate assessment of early succession.

REPLY: Pitchers die back after each season and only “new growth” pitchers were selected. That being said, the pitchers were already open by the start time of the study.

REPLY: We have moved this information to the methods section for clarity.

REPLY: We have added the volume of fluid in each pitcher to the Supplementary Table S1. The rosettes were spaced along a 26m transect with approximately 8 meters between rosettes.

REPLY: We realize that removing liquid will influence the community and impact the measurement of succession, and we discuss this in lines 400-410. We note that the dynamics we observe are in line with other studies of microbial community succession. We are careful to mention disturbance and argue that our results are put in proper context of our sampling design.

REPLY: The morphology of these pitcher plants is such that it was impossible to remove all the fluid from the plant. While removal of the liquid is certainly a disturbance, the size of the organisms being studied meant that many cells likely remained, and likely in similar proportions as the rest of the fluid. These plants are also subject to rain (possibly flooding/diluting the community) and wind (loss of volume), and so their natural state is quite stochastic.

Why were OTUs used instead of ASVs?

REPLY: OTUs were used to align this study with previous work using the SAR specific primers. We agree that ASVs have utility but can be difficult to interpret for groups such as ciliates that have intra-individual variability of rDNA. Moreover, unpublished data from our lab have shown substantial problems with DADA2 when applied to microeukaryotes: it is not parameterized to capture the length variation found among microeukaryotes.

Figure 2. It would be helpful to visualize which of the Sarracenia samples came from which site.

REPLY: We did try this initially but the many shapes made the figure too difficult to interpret. We now point our reader to supplement that includes these details.

Figure 3A. Labelling the Date as a circle is confusing with the Forest Edge circle. Additionally, it is confused what “Pool Edge” means.

REPLY: We have added text to the legend to clarify the shapes and the meaning of pool edge.

Table 1. Minor detail, but signify that the dates are Month/Day.

REPLY: We have made this correction.

Attachment

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PLoS One. doi: 10.1371/journal.pone.0270913.r003

Decision Letter 1

Theodore Raymond Muth

13 Apr 2022

PONE-D-21-40078R1Illuminating Protist Diversity in Pitcher Plants and Bromeliad TanksPLOS ONE

Dear Dr. Sleith,

In the revised (2nd revision) manuscript, please address the continued concerns of reviewer three. There remains a need for clarification on the age of the pitcher plants sampled and the possible sampling from pitchers that lasted over the winter from an earlier season. If there is a reason that the pitcher plants in this study are maturing earlier than expected by the third reviewer, please make that clear. Please address the question regarding dipteran larvae remaining in the pitcher plants.

Please submit your revised manuscript by May 28 2022 11:59PM. If you will need more time than this to complete your revisions, please reply to this message or contact the journal office at plosone@plos.org. When you're ready to submit your revision, log on to https://www.editorialmanager.com/pone/ and select the 'Submissions Needing Revision' folder to locate your manuscript file.

Please include the following items when submitting your revised manuscript:

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We look forward to receiving your revised manuscript.

Kind regards,

Theodore Raymond Muth

Academic Editor

PLOS ONE

Journal Requirements:

Please review your reference list to ensure that it is complete and correct. If you have cited papers that have been retracted, please include the rationale for doing so in the manuscript text, or remove these references and replace them with relevant current references. Any changes to the reference list should be mentioned in the rebuttal letter that accompanies your revised manuscript. If you need to cite a retracted article, indicate the article’s retracted status in the References list and also include a citation and full reference for the retraction notice.

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Reviewers' comments:

Reviewer's Responses to Questions

Comments to the Author

1. If the authors have adequately addressed your comments raised in a previous round of review and you feel that this manuscript is now acceptable for publication, you may indicate that here to bypass the “Comments to the Author” section, enter your conflict of interest statement in the “Confidential to Editor” section, and submit your "Accept" recommendation.

Reviewer #1: (No Response)

Reviewer #2: All comments have been addressed

Reviewer #3: (No Response)

**********

2. Is the manuscript technically sound, and do the data support the conclusions?

Reviewer #1: Yes

Reviewer #2: Yes

Reviewer #3: Partly

**********

3. Has the statistical analysis been performed appropriately and rigorously?

Reviewer #1: Yes

Reviewer #2: Yes

Reviewer #3: Yes

**********

4. Have the authors made all data underlying the findings in their manuscript fully available?

Reviewer #1: Yes

Reviewer #2: Yes

Reviewer #3: Yes

**********

5. Is the manuscript presented in an intelligible fashion and written in standard English?

Reviewer #1: Yes

Reviewer #2: Yes

Reviewer #3: Yes

**********

6. Review Comments to the Author

Reviewer #1: I feel that the authors responded satisfactorily to the previous comments and that the manuscript meets PLoS ONE review criteria.

Reviewer #2: The authors have revised their manuscript, “Illuminating Protist Diversity in Pitcher Plants and Bromeliad Tanks”, following comments from myself and two other reviewers. The revised manuscript adequately addresses all of my previous comments and I do not have any additional comments. Great job handling all of the (sometimes conflicting) comments and thanks for a fun read!

Reviewer #3: Thank you very much for the revisions to the manuscript. I commend the authors for the work they did. Overall, I find the study interesting and timely. Eukaryotic diversity is understudied in general, and especially in pitcher plant systems. I still, however, have concerns about the methods in Study 2. My concerns do not mean that there is a problem with the data, but rather in the question that is being addressed (Succession). My reasons for this are detailed below:

1) Standardization of pitcher leaf age:

Sampling in the beginning of May in Massachusetts seems very early in the growing season for Sarracenia purpurea at this latitude. It is thus surprising that there were new leaves/pitchers available on the rosettes (especially 3 new leaves per rosette). More surprising is that the new leaves that were used in the study were open and filled with large volumes of water (as shown in the supplemental data), unless several large rainstorms occurred just before sampling. I just can’t imagine that the new pitchers (from that season) would be large enough to hold such a large volume at this time period.

Not all pitchers die back each year. Instead, they overwinter, with the snow protecting them. There will, of course, be some old pitchers from several seasons in the past that have died back (the pitchers closer to the ground), but the middle of the rosette will have pitchers from last season that have survived the winter as well as the new pitchers that are growing in the current season. It can sometimes be difficult to determine last season’s pitchers from new growth pitchers if the new growth pitchers are ~4 weeks or older. I am therefore uncertain if the authors used pitchers from that season or overwintered pitchers. For the study to examine Succession, the age and opening time of the pitchers needed to be better standardized. This can be done by selecting very new (fresh, flexible green leaves that had just opened) pitchers. The first sampling event should have then been done several weeks after they opened, so that there was time for the pitchers to fill with water.

Study 2 still informs us about the inquiline community that inhabits pitchers, but without the initial standardization, it acts more of a snapshot at each point in the season, and not succession.

Apologies if my phenology is incorrect. Addition of field site information (flowering time, new leaf production, precipitation) should be added to the manuscript.

2) Dipteran larvae in the inquiline community:

The large volume of water that was removed from each pitcher during sampling would also remove any of the dipteran larvae that are essential for dynamics of the inquiline community. Some (the midge and some of the mosquito larvae) could be down at the bottom of the leaf near the decomposing insects, so it is likely that those ones could remain in the community after each sampling. It is unfortunate that the authors did not record midge and mosquito larvae during the sampling so that the authors have an idea of how many were lost from the community during each sampling. Were there any mosquito larvae stuck on the filter after the inquiline community was filtered?

Along those lines, a better reference than Rango 1999 for seasonal Wyeomyia dynamics would be to use the Harvard Forest Data Archives. These archives contain data from the authors’ field site and/or sites in the region and have information about the seasonal dynamics of the inquiline mosquito larvae in the system. https://harvardforest1.fas.harvard.edu/exist/apps/datasets/showData.html?id=hf193

It very well might be that the mosquito larvae densities do not peak until mid-summer, and therefore are not a part of the May-June sampling that occurred in this study. In the lower latitude part of Sarracenia purpurea’s distribution (e.g., Florida) the mosquito larvae are the most abundant as soon as the leaf opens. If that is the case in this study site as well, then the mosquito density would be greatly impacted. If overwintered leaves were indeed used for this study, there could be mosquito larvae present because they were in diapause in the leaves during the winter.

3) Disturbance by removing large volumes of water during sampling: a large volume was taken from each community during each sampling time period in Study 2. The authors do replace the volume they removed with the supernatant, but this is still a large, open niche space. I agree that there will be some microbes that remain at the bottom of the leaf during each sampling time period, but the community is undergoing a great disturbance during each sampling period, with diversity potentially being diluted each time.

In summary, I do think this study has accomplished a lot of things and is interesting, it is just that there are some potential issues in the methods which will affect the focus of the paper as it currently is written. In general, I suggest that the authors be careful with the word “succession” in this study. Instead, I suggest that the authors use “seasonal” dynamics or something along those lines.

**********

7. PLOS authors have the option to publish the peer review history of their article (what does this mean?). If published, this will include your full peer review and any attached files.

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Reviewer #1: No

Reviewer #2: Yes: Erica Holdridge

Reviewer #3: No

PLoS One. 2022 Jul 27;17(7):e0270913. doi: 10.1371/journal.pone.0270913.r004

Author response to Decision Letter 1

15 Jun 2022

We thank the reviewers for their helpful comments, which we have addressed in detail below.

****

Reviewer #3:

Thank you very much for the revisions to the manuscript. I commend the authors for the work they did. Overall, I find the study interesting and timely. Eukaryotic diversity is understudied in general, and especially in pitcher plant systems. I still, however, have concerns about the methods in Study 2. My concerns do not mean that there is a problem with the data, but rather in the question that is being addressed (Succession). My reasons for this are detailed below:

REPLY: As a result of the comments of this reviewer, we have further clarified the language in our manuscript including reframing successional patterns as seasonal change to the community over time.

1) Standardization of pitcher leaf age:

REPLY: We thank the reviewer for these comments. We have reviewed field photos and believe it is possible that pitchers from the previous season were sampled, given how early in the season it was, and the size of some of the pitchers. We now clarify that the pitchers we sample may be of different generations, and that seasonality refers to turn over of the microbial community from spring through summer. We have also added that variance among spring pitchers might be due to the age of individual pitchers (i.e. those that overwintered vs new growth).

Study 2 still informs us about the inquiline community that inhabits pitchers, but without the initial standardization, it acts more of a snapshot at each point in the season, and not succession.

REPLY: We believe that our improved language has clarified this issue.

Apologies if my phenology is incorrect. Addition of field site information (flowering time, new leaf production, precipitation) should be added to the manuscript.

REPLY: We agree that to understand how the age and phenology of the pitcher plant impacts the inquiline community (e.g. Armitage 2017) requires repeated sampling of pitchers with standardized ages. However, our current study is interested in how the microbial community changes during the spring growing season, irrespective of pitcher age (e.g. Buosi et al. 2015). Pitcher age may be a factor in structuring these communities, but our current study demonstrates that seasonality tends to be a more important driver, given that communities in leaves of different ages converge by the end of the sampling window (Figure 4), as opposed to communities being structured by pitcher age. Nevertheless, to avoid giving a false impression we have removed the term succession when discussing our results and simply use the term seasonal changes. We have also clarified the age of pitchers and sampling strategy as well as site information in the Methods and Discussion.

2) Dipteran larvae in the inquiline community:

REPLY: We appreciate this insight and have looked at the Ellison & Gotelli dataset. The hf193-04: food web structure data show that for Hawley Bog (as well as Molly Bog) the density for Wyeomyia peaks in August and September (with some detections in July, and very few in June). We have added this information to the discussion.

REPLY: The volume of fluid removed does represent a disturbance to the system, and we have added additional text addressing this issue in lines 388-398 of the Discussion. The final sampling window shows that communities in the pitcher plants on Day 53 were quite similar to the background communities (Figure 4, yellow filled symbols are clustered with open fill bog water samples). If we were diluting diversity and changing the community composition drastically, we would expect the communities to be quite unique and would expect samples from Day 53 to cluster on their own near no natural samples. We contend that the dynamic/stochastic nature of the pitcher system is such that the disturbance created by our sampling is not beyond the scope of what is possible in nature, and therefore our results are a useful addition in the study of microbial eukaryote communities. We hope that this study inspires others to track additional communities using other experimental methods.

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PLoS One. doi: 10.1371/journal.pone.0270913.r005

Decision Letter 2

Theodore Raymond Muth

20 Jun 2022

Illuminating Protist Diversity in Pitcher Plants and Bromeliad Tanks

PONE-D-21-40078R2

Dear Dr. Sleith,

We’re pleased to inform you that your manuscript has been judged scientifically suitable for publication and will be formally accepted for publication once it meets all outstanding technical requirements.

Within one week, you’ll receive an e-mail detailing the required amendments. When these have been addressed, you’ll receive a formal acceptance letter and your manuscript will be scheduled for publication.

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Theodore Raymond Muth

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PLOS ONE

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Reviewers' comments:

PLoS One. doi: 10.1371/journal.pone.0270913.r006

Acceptance letter

Theodore Raymond Muth

30 Jun 2022

PONE-D-21-40078R2

Illuminating Protist Diversity in Pitcher Plants and Bromeliad Tanks

Dear Dr. Sleith:

I'm pleased to inform you that your manuscript has been deemed suitable for publication in PLOS ONE. Congratulations! Your manuscript is now with our production department.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

(TIF)

Click here for additional data file.^{(741.9KB, tif)}

S2 Fig. Observed richness (number of OTUs) by pitcher opening diameter and pitcher volume shows no relationship between richness and pitcher morphology.

(TIF)

Click here for additional data file.^{(533.6KB, tif)}

S3 Fig. The PCoA from Fig 2 with 95% data ellipses representing the multivariate t-distribution (solid) and multivariate normal distribution (dashed).

(TIF)

Click here for additional data file.^{(1MB, tif)}

S1 File. Sample SRA and collection detail information.

(CSV)

Click here for additional data file.^{(11.4KB, csv)}

S2 File. OTU table with phylogenetic taxonomic assignment and read counts per OTU.

(CSV)

Click here for additional data file.^{(31.3KB, csv)}

Attachment

Submitted filename: Response to Reviewers.docx

Click here for additional data file.^{(27.5KB, docx)}

Attachment

Submitted filename: Response to Reviewers_LAK_RSS.docx

Click here for additional data file.^{(19.5KB, docx)}

Data Availability Statement

Raw reads associated with this study are available under NCBI BioProject ID: PRJNA682436.

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PERMALINK

Illuminating protist diversity in pitcher plants and bromeliad tanks

Robin S Sleith

Laura A Katz

Roles

Abstract

Introduction

Materials and methods

Sample collection—Study 1

Table 1. Study and sampling information.

Sample collection—Study 2

Sample processing—Studies 1 and 2

Data analysis

Results

Amplicon analysis of phytotelmata (Studies 1 and 2)

Fig 1. The 135 focal OTUs from across plant hosts and geography illustrate generalist OTUs (i.e. not restricted by plant host or geography) and specialist OTUs (i.e. restricted by plant host or geography).

Study 1: The influence of plant host on OTU richness and community structure

Fig 2.

Study 2: The influence of season on OTU richness and community structure

Fig 3. Network analysis demonstrates that most OTUs in a rosette of 3 pitchers are shared with the bog water below the rosette.

Fig 4.

Fig 5.

Discussion

Richness

Study 1: The influence of plant host on community structure

Study 2: The influence of season on community structure

Conclusions

Supporting information

Acknowledgments

Data Availability

Funding Statement

References

Decision Letter 0

Theodore Raymond Muth

Roles

Author response to Decision Letter 0

Decision Letter 1

Theodore Raymond Muth

Roles

Author response to Decision Letter 1

Decision Letter 2

Theodore Raymond Muth

Roles

Acceptance letter

Theodore Raymond Muth

Roles

Associated Data

Supplementary Materials

Data Availability Statement

ACTIONS

PERMALINK

RESOURCES

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