1. Introduction
Currently, there are about 860 species of carnivorous plant species that evolved from within at least 10 different linages [1,2]. Carnivorous plants generally reside in mesic to wet nutrient-poor habitats [3], and many species (about 10%) are currently known from a single location [1]. Carnivorous plants are believed to catch prey to supplement their nitrogen, phosphate, potassium, and other nutritional requirements [4]. Interestingly, studies have shown that some carnivorous plants produce digestive enzymes [5,6], while other species appear to rely on microbes living within their traps to produce the necessary enzymes required for prey digestion [7].
In carnivorous plants, there are five types of traps (adhesive, eel, pitcher, snap, and suction), with eel traps being the least understood [8,9]. Eel traps are Y-shaped, underground hollow leaves that act as roots and assist with nutrient uptake. Eel traps have hairs inside that force prey to move in one direction toward the vesicle (stomach), where digestion occurs [10]. Interestingly, the carnivorous plant family Lentibulariaceae contains genera with different trapping mechanisms: Genlisea species have eel traps, Pinguicula spp. have adhesive traps, and Utricularia spp. have suction traps [11]. Because species that form eel traps are the least understood, we undertook the present study to examine the microbial community of G. hispidula.
The genus Genlisea is estimated to be comprised of around thirty species [10] with a distribution from tropical Africa, including Madagascar, to Central and South America [12]. Genlisea species form a small, above-ground photosynthetic rosette of leaves and have below-ground achlorophyllous rhizophyll leaves. Rhizophylls are modified leaves that act as roots (Figure 1) and form eel traps and vesicles in this genus, which act as the digestive structure [13]. Genlisea eel traps function by indiscriminately allowing small microorganisms to enter slits along the hollow spiral-shaped leaves where pointed hairs prevent the prey from leaving. This forces the prey to move up the hollow spiral-shaped leaves and into the vesicle where the anoxic conditions cause death [14] and is the location where digestion occurs [15]. Currently, both reactive oxygen species and acid phosphatases are known to be produced by Genlisea for digestion [13,16,17]. In addition to host-derived enzymes/reactive oxygen species, there is also the presence of lipases, phosphatases, and proteases that appear to originate from the leaf trap microbial community [17].
Researchers have recognized the importance of carnivorous plant trap-associated microbes and have conducted microbiome studies. For example, Caravieri et al. [18] investigated the trap bacterial communities of G. filiformis and U. hydrocarpa, and Koopman and Carstens [19] investigated the microbial communities in Sarracenia alata traps. Chan et al. [20] investigated the bacterial community within a Nepenthes sp. digestive fluid, while Cao et al. [17] examined the host–microbiome interactions of two Genlisea spp. using a meta-transcriptomic approach. Overall, it appears that Genlisea spp. have a wide range of prey due to the passive nature of prey capture [10], which includes algae, collembola, crustaceans, cyanobacteria, mites, protozoa, nematodes, and protozoa [21].
Most of these studies focused on the bacterial community; however, it is known that fungi play key roles in digestion for many insect species [22]. Therefore, examining trap-associated fungal species may provide a greater understanding of the role fungi may play in host digestion. Although Cao et al. [17] performed a meta-transcriptomic approach to understanding the host–microbiome of two Genlisea spp., Romão et al. [23] concluded that culture-dependent techniques are complementary to these approaches as many cultured microbes are not found within culture-independent datasets. This suggests that further studies of the culturable microbiome are needed to fully elucidate the composition of the host–microbiome of G. hispidula. In particular, the mycobiome of G. hispidula has not been studied in detail, and the culture-dependent analysis of the microbiome (bacteria) of G. hispidula has not been previously completed, leading to a knowledge gap in our understanding of G. hispidula-associated bacteria and fungi.
When identifying bacterial and fungal taxa using only molecular data, species delimitation generally falls into two categories: DNA sequence-based similarity and tree-based models [24] of which both initially rely on a gene sequence. For bacteria, the small ribosomal subunit RNA (16S rDNA) gene is the most important taxonomic marker [25], while the internal transcribed spacer (ITS) region was defined as the fungal barcode for molecular identification [26]. Molecular sequence similarity thresholds, at the species level, for both bacteria and fungi have changed over time, with the species-level identification of bacteria currently around 98.7% using the full-length 16S rDNA gene (ca. 1500 bp) [27]; for fungi, ITS sequence similarity thresholds of 99.61% are suitable for filamentous fungi [28], while ITS sequence similarity of 98.31% is suitable for Ascomycota yeast species and 98.61% is suitable for Basidiomycota yeast species [29]. Tree-based model approaches for species delimitation include the General Mixed Yule Coalescent Model (GMYC) [30] and the Bayesian Poisson Tree Processes model (bPTP) [31].
The objectives of the study were to isolate bacteria and fungi from all major parts of G. hispidula (leaves, rhizophyll, and vesicles) to determine (1) species presence, (2) diversity, and (3) the presence of novel taxa. In addition, we review what is known about the microbial contribution to host digestion. We hypothesized that there would be culturable bacterial and fungal species within/on each major plant part, and that the above-ground bacterial and fungal species would be different compared to the below-ground species. Lastly, we anticipated that novel microbial species would be present. This study is important because host conservation efforts may require the conservation of host-associated microbes, and lastly, carnivorous plant structures are unique micro-habitats that have been relatively unexplored for novel taxa.
2. Materials and Methods
2.1. Genlisea Hispidula Plants, Growing Conditions, and Parts Examined
Four G. hispidula plants (true to name and propagated via division in-house and approximately 6–9 months of age; -personal communications 14 December 2023 and 18 January 2024) were obtained from Carnivorous Plant Nursery (Smithsburge, MD, USA). All plants were maintained in a greenhouse (20–31 °C and 16–70% RH) in open trays containing sphagnum peat moss (Premier). The trays were maintained in a continuously wet state with reverse osmosis water. Plant tissues used in this experiment included the above-ground leaves, the rhizophylls, and the vesicles (Figure 1A,B). Flowers were not examined as part of this experiment as they were not present.
Figure 1Genlisea hispidula general body plan showing sampled tissues. (A) drawing indicating sampling areas; (B) image of young G. hispidula with photosynthetic leaves and young developing rhizophyll. Leaves are photosynthetic; rhizophylls are achlorophyllous leaves that act as roots and comprise the eel traps. Microorganisms enter slits along the corkscrew branches. From there, hairs only permit prey movement upward, where they are digested in the vesicles.
[Figure omitted. See PDF]
2.2. Tissue Preparation, Medium Inoculation, and Isolation
From each individual plant (4 total), 5 pieces of plant tissue were antiseptically removed from each tissue type (above) using sterile forceps and a sterilized dissecting needle. Three pieces of each tissue type were surface sterilized in 1 mL NaOCl solution (3% available Cl) for one min and rinsed twice in 1 mL autoclaved distilled water to remove residual NaOCl. The other two pieces were rinsed in 1 mL autoclaved distilled water for one min to remove debris. Samples were transferred to 1.5 mL microcentrifuge tubes and grounded into fine pieces with sterilized plastic pestles (Sigma Aldrich, St. Louis, MO, USA), and 1 mL of each tissue homogenate was serially diluted (1:2, 1:4, 1:8, 1:16, and 1:32) in sterile water. Next, the inoculant (1 mL) was lawn plated onto 90 mm Petri Plates containing potato dextrose agar (PDA, Difco, Franklin Lakes, NJ, USA, pH 5.6) after mixing each dilution thoroughly to suspend the ground tissue. To encourage the growth of both bacteria and fungi, the PDA medium was not acidified, nor were any antibiotics added. Plates were sealed with parafilm and incubated in a room that fluctuated between 18 and 23 °C, in 24 h darkness. Inoculated Petri plates were monitored daily for bacterial and fungal colony growth, and colonies with unique morphology were transferred to PDA plates and stored under the same conditions. All axenic cultures were placed onto PDA slants (4 °C) and 20–40% glycerol (sterile water and glycerol (−80 °C) for long-term storage.
2.3. Molecular Identification
DNA was isolated by placing a small portion of each actively growing isolate (less than 1 mm (bb size) into a 1.5 mL microcentrifuge tube containing 100 μL of 1X TE buffer and then heated for 2 min in an 1100-watt microwave. Then, each tube was centrifuged at 21,130 rcf for 2 min to pelletize the cells. The supernatant containing the DNA was transferred to a new 1.5 mL microcentrifuge tube and stored at −20 °C until needed. For bacteria, the 16S rDNA universal primer pair 27F (5′-AGAGTTTGATCCTGGCTCAG-3′) and 1392R (5′-GGTTACCTTGTTACGACTT-3′) were used [32]. For fungi, the internal transcribed spacer region (ITS1-5.8S-ITS2) was amplified using ITS1F (5′-CTTGGTCATTTAGAGGAAGTAA-3′) and ITS4 (5′-TCCTCCGCTTATTGATATGC-3′) primers [33,34]. All PCRs were completed in a total reaction volume of 25 μL (12.5 μL MyTag Mix (Bioline, Memphis, TN, USA), 1 μL of each 20 μM primer, 8.5 μL DNA-free water, 2 μL DNA) on an Eppendorf Mastercycler Pro S thermocycler. The thermocycle settings for bacterial 16S rDNA amplification were an initial denaturation at 94 °C for 6 min followed by 35 cycles (94 °C for 15 s, 49 °C for 15 s, and 72 °C for 90 s) and a final extension at 72 °C for 5 min. The thermocycle settings for fungal ITS amplification were an initial denaturation at 94 °C for 6 min followed by 30 cycles (94 °C for 30 s, 55 °C for 30 s, and 72 °C for 60 s) and a final extension at 72 °C for 10 min. Gel electrophoresis (1% TAE agarose gel) and SYBR GelRed were used to verify the presence of a PCR product. Both PCR purification and Sanger sequencing were completed by the Purdue Genomics Center for sequencing. The base calls were reviewed and edited using Sequencer 5.4.6. All high-quality bacterial and fungal sequences were deposited in GenBank (bacteria accession numbers (OR799523-OR799556) and fungal accession numbers (OR779983–OR780009).
2.4. Unique Species Determination and Visualization
Unique species delineation was completed using the Bayesian implementation of the Poisson tree processes (bPTP) model using the online server (
3. Results
3.1. Bacteria
In total, 76 bacterial isolates were cultured, consisting of 48 unique isolates based on bPTP analysis (Table S1, Figure 2). Nine of these unique bPTP species (18.75% (9/48)) are new records for the association of G. hispidula (Table S1). Of the unique isolates based on bPTP analysis, six were identified at the species level (Table S1). All remaining unique isolates based on bPTP analysis could not be identified at the species level, with unknown species being grouped into 11 genera (Acidocella, Bacillus, Burkholderia, Herminiimonas, Jatrophihabitans, Methylobacterium, Microbacterium, Novosphingobium, Paenibacillus, Paraburkholderia, and Rhodopseudomonas), other unidentified species being grouped into 4 bacterial families, additional unidentified species being placed into 2 orders, and based on 16S rDNA gene PhyML reconstruction, 1 unknown bacterium was placed into the phylum Pseudomonadota (Table 2; Figure 3). Acidocella facilis was the most frequent bacterial species isolated from all tissues (Table S1). The genus Burkholderia had the most bPTP species (Table 2; Figure 3) and was the dominant bacterial genus across all tissue types. Only bacterial isolates from the genera Acidocella and Burkholderia were isolated from vesicle tissue, in addition to unidentified bacteria that were only placed into the family Alcaligenaceae (Table 2). Only one Burkholderia sp. and the unidentified Alcaligenaceae bacterium were isolated from sterilized vesicle tissues (Table S1, Figure 3).
3.2. Fungi
In total, 59 fungal isolates were obtained, which consisted of 29 putative unique isolates based on bPTP analysis (Table S1, Figure 2). Twenty-four of these unique bPTP species (82.75% (24/29)) are new records for the association of G. hispidula (Table S1). Of the 29 unique bPTP isolates, 5 could be identified to species, additional isolates were identified to 3 genera, and one isolate was identified only to order (Table 3). Based on ITS PhyML reconstruction, the sister taxa to the Eurotiales isolate was a Penicillium species (Figure 4). Trichomonascus vanleenenianus was the most frequently isolated species, while the genus Saitozyma had the greatest number of unique species, followed by the genus Penicillium (Table 3). Saitozyma was the only Basidiomycota genus to be isolated. Only Saitozyma spp. and Trichomonascus vanleenenianus were isolated from the vesicle tissue (Table S1, Figure 5). One Saitozyma sp. was solely isolated from sterilized vesicle tissue, while two additional Saitozyma spp. were isolated from both non-sterilized and sterilized vesicle tissues (Table S1, Figure 4).
3.3. Diversity and Novel Species
More species of culturable bacteria were isolated than fungi (Table S1). The leaf tissue of G. hispidula contained greater bacterial and greater fungal species-richness, followed by the rhizophyll, while the vesicle tissues had the least (Table S1). Most bacterial and fungal species were tissue-specific (Figure 3 and Figure 4). Only one bacterial bPTP species (A. facilis) and two fungal species (T. vanleenenianus and S. podozolica) were isolated across more than one tissue type (Figure 3 and Figure 4). Most bacterial isolates could not be identified to species, with 37.5% (18/48) of bPTP isolates being assigned to Family rank or higher (Table 2). In addition, one leaf-associated bacterium could only be assigned at the phylum level with its closest sequence similarity match in NCBI (99% query coverage, 93.8% identity) to AB111107 Proteobacterium LS-1. Most fungal isolates identified to higher ranks cannot be placed to species using the ITS region.
4. Discussion
Our culture-based approach has provided a surprising number of novel G. hispidula-associated bacteria and fungi that were not previously known to be associated with the host. Based on 16S rDNA gene sequence similarity there is a potential for 18 new bacterial species (8 Alcaligenaceae spp., 1 Burkholderiales sp., 1 Micrococcales sp., 5 Nitrobacteraceae spp., 1 Rhodospirillaceae sp., 1 Sphingomonadaceae sp., and 1 unidentified bacterial species), and based on ITS sequence similarity, there is a potential for 1 new fungal species within the order Eurotiales. We also demonstrated that a culture-based approach identified additional species which were not identified in the previous metagenomic study conducted by Coa et al. [17] although we note that there was some overlap in identified species based on sequence similarity. Although culture-based approaches are more laborious, they are extremely important for providing living culture for taxonomy (type specimens), for industrial purposes, and for understanding their role in host health via experimentation.
We note that our G. hispidula plants were not collected from their native habitat, and research has shown that greenhouse grown plants may have different microbial communities as compared to those grown in the wild. Sexton et al. [44] showed that the bacterial communities (based on the number of isolated genera) of Sarracenia leucophylla and S. purpurea rhizomes were more diverse when grown in the greenhouse as compared to the wild. They concluded that this difference between greenhouse and wild microbial communities was likely due to the different environmental conditions that influence the bacterial species present within each environment [44]. In addition to environmental conditions, other researchers have shown that plant microbial communities fluctuate at different plant growth stages [45], via the prey they capture [46], and seasonally [47]. Therefore, future culture-dependent and culture-independent sampling from wild populations, different growth stages, and temporal sampling would be advantageous to determine the broad extent of the G. hispidula microbiome.
Lastly, it has been suggested that standard surface sterilization techniques may need to be altered for different plant species to be effective at removing the epiphytic community [48]. Therefore, we do not know for certain if our surface sterilized isolates are endophytes or if they were epiphytic survivors of the surface sterilization process. Therefore, further refinement of the sterilization process, designed specifically for G. hispidula, should be conducted in the future.
4.1. Contribution of Bacteria and Fungi to Aid Host Digestion and Further Thoughts
Genlisea hispidula grows in wet environments where the substrate is anoxic [49]. In addition, the measured oxygen level within the central vesicle cavity was also suggested to be anoxic [14]. This suggests reduced oxygen levels below ground and in below-grown structures. Consequently, it was proposed that commensal microorganisms that inhabit the central cavity would need to tolerate anoxic conditions [14]. This was supported when a meta-transcriptome analysis indicated that the bacterial community within Genlisea vesicles was enriched for facultatively anaerobic bacteria [17]. Interestingly, the same authors also reported that aerobic bacterial species were dominant within the eel trap. Although these authors found diverse bacterial community, the authors ultimately concluded that bacterial species mainly contributed to the nitrogen cycle and not host digestion [17]. They concluded that non-host degradative enzymes were derived from the larger microbes, particularly Alveolata and amoeboid protists, green algae species, and metazoan species [17]. Although this metagenomic study provides insights into the potential contribution of the microbial community to host digestion, this study (as well as ours) used commercially obtained plants which were then subsequently grown under greenhouse conditions. Under greenhouse conditions, Sexton et al. [44] found elevated diazotrophic bacteria (nitrogen-fixing bacteria) numbers in S. oreophila rhizomes as compared to S. oreophila rhizomes in their natural habitat, suggesting that bacterial contribution to the nitrogen cycle found by Coa et al. [17] may not be as strong in their natural habitat. Ultimately, additional studies on G. hispidula in their natural habitat are needed to better understand the extent to which the microbial community contributes to host digestion.
4.2. Differences among G. hispidula Tissue Types
We hypothesized that there would be culturable bacterial and fungal species within/on each major plant part and that the above-ground bacterial and fungal species would be different as compared to the below-ground species. As it related to bacteria, we isolated a surprising number of novel culturable bacteria based on full-length 16S rDNA gene sequence similarity, which has the potential for species-level determination [50]. Interestingly, three bacterial species, A. facilis, a Burkholderia sp., and an Alcaligenaceae sp., were found within the vesicle, which was consistent with results from a previous metagenomic study [17]. In addition, Coa et al. [17] found the genera Acidocella and Burkholderia within G. hispidula eel traps. We isolated one Burkholderia sp. and one unknown Alcaligenaceae sp. from surface sterilized vesicle tissues, which could represent vesicle-associated bacteria that may be providing an important function, but further functional and association studies would be needed, and to the extent this association exists in natural populations would need to also be determined.
Interestingly, we did not see an above-ground or below-ground bacterial species overlap, with only a few species being present in all tissue locations (Figure 3). A greater diversity of bacteria on leaves is likely due to leaves having distinct epiphytic and endophytic bacterial communities [51]. We also isolated these bacteria in an aerobic environment, which is consistent with oxygen levels in leaf tissue. In contrast, as previously mentioned, both underground tissues are in a hypoxic/anoxic environment (rhizophyll) or hypoxic/anoxic state (vesicle), which would limit our ability to isolate some of these species which will not grow or grow very slowly using only aerobic isolation techniques. Consequentially, future culture-dependent isolation techniques should utilize multiple culture medias and isolation strategies to maximize the number of species isolated. As it relates to fungi, the same two patterns were evident, with leaf tissue having the most diversity of culturable fungal species as compared to the rhizophyll and vesicle tissues, and little above-ground or below-ground species overlap. This is also likely due to the hypoxic and anaerobic conditions found within the below-ground tissues as compared to the above-ground tissue. In this study, only two fungal species, S. podozolica and T. vanleenenianus, were found within the vesicles. In contrast to bacteria, only a few fungal genera have been reported from G. hispidula, and there appears to be no sole study on the contribution of fungi to G. hispidula host digestion. The previous meta-transcriptome analysis [17] identified only three genera: Glarea, Rhizophydium, and Symbiotaphrina from eel traps. However, based on blastn analysis, it appears that Cladosporium spp. and Toxocladosporium spp. are found within the datasets of Choa et al. [17]. Both Glarea and Rhizophydium species have been isolated from aquatic habitats [52,53], and Symbiotaphrina are free-living and known symbionts with beetles [54]. In this study, the two fungal species that were isolated from the host’s vesicle had a yeast morphology. The asexual state of Symbiotaphrina species is also yeast-like. This suggests that a yeast morphology may be advantageous for fungal survival within the vesicle.
In this study, we recovered 29 fungal species, one of which appears to be exclusively associated with the vesicles (Table S1, Figure 3). Saitozyma podozolica has been frequently isolated from aquatic environments [36,55]. In this study, S. podozolica was isolated from all three tissues examined. Enzymatically, S. podozolica is known to utilize hexose and pentose sugars, including xylose [56]. In addition, one Saitozyma sp. was solely isolated from sterilized vesicle tissues. This isolate may represent a vesicle-associated species; determining the extent of its association with G. hispidula is also of future interest. Trichomonascus vanleenenia was also isolated from all three tissues of G. hispidula. This fungus was previously isolated in 2018 from soil samples in the Netherlands and was shown to ferment glucose and did not grow on nitrate and nitrite sole nitrogen sources [57]. Increasing our understanding of T. vanleenenia is also of interest due to its relatively recent description as a new species in 2018.
Ultimately, additional research will be required to determine the extent of bacterial and fungal associations with G. hispidula’s overall health. Many new species of bacteria were isolated from G. hispidula, suggesting that these understudied plants are an important source for obtaining new bacterial taxa. Continued research should be completed as obtaining novel bacterial and fungal taxa from unexplored micro-habitats offers a promising venue for plant conservation, plant disease control, and drug discovery.
Conceptualization, M.C.A.; methodology, M.C.A.; formal analysis, D.B.R.; resources, M.C.A.; data curation, M.C.A.; writing—original draft preparation, D.B.R.; writing—review and editing, D.B.R. and M.C.A.; supervision, M.C.A.; project administration, M.C.A.; funding acquisition, M.C.A. All authors have read and agreed to the published version of the manuscript.
Not applicable.
The original contributions presented in the study are included in the article and
This study was initiated as an undergraduate research project by Kirk Rumple, to whom we are indebted. We thank Matt Fujita for providing plants for our illustrations.
The authors declare no conflicts of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript; or in the decision to publish the results.
Footnotes
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Enlarge this image.Figure 2. Cultural bacteria and fungi isolated from Genlisea hispidula. Bacteria: (A) Acidocella facillis, (B) Burkholderia sp., (C) Burkholderia sp., (D) Novosphingobium sp., (E) Microbacterium ginsengisoli, and (F) Rhodopseudomonas sp. Fungi: (G) Cladosporium sphaerospermum culture morphology, (H) C. sphaerospermum conidiophore (Cp), conidiogenous cell (Cc) and conidia (C), (I) Aspergillus sydowii, (J,K) A. sydowii conidiophore (Cp), conidiogenous cell (Cc) and conidia (C). Scale bars: (A–F) 1000 μm and (H,J,K) 20 μm.
Enlarge this image.Figure 3. Cladogram of unique bacterial isolates from Genlisea hispidula based on bPTP analysis. Cladogram based on 16S rDNA sequences using primers 27F/1392R, g-block analysis, and PhyML reconstruction. Pie chart color reflects tissue type: red = leaf, gray = rhizophyll, blue = vesicle. Pie chart size represents the number of cultured isolates within each unique bPTP species delimitation. The inner circle represents the bacterial phylum: gray = Pseudomonadota, green = Actinomycetota, and purple = Bacillar (Firmicutes). The outer circle represents the sterilization technique: orange = non-surface sterilization, yellow = 3% bleach sterilization, and blue = isolates from both non-surface sterilization and 3% bleach sterilization.
Enlarge this image.Figure 4. Cladogram of unique fungal isolates from Genlisea hispidula based on bPTP analysis. Cladogram based on ITS rDNA sequences using primers ITS1F/ITS4, g-block analysis, and PhyML reconstruction. Pie chart color reflects tissue type: red = leaf, gray = rhizophyll, blue = vesicle. Pie chart size represents the number of cultured isolates within each unique bPTP species delimitation. The inner circle represents the fungal phylum: gray = Ascomycota, light blue = Basidiomycota. The outer circle represents the sterilization technique: orange = non-surface sterilization, yellow = 3% bleach sterilization, and blue = isolates from both non-surface sterilization and 3% bleach sterilization.
Enlarge this image.Figure 5. Fungi isolated from Genlisea hispidula vesicle tissue. (A) Trichomonascus vanleenenia culture morphology, (B) T. vanleenenia yeast-like morphology, (C) T. vanleenenia hyphal morphology, (D) Saitozyma podozolica culture morphology, (E) S. podozolica yeast cells. All scale bars = 20 μm.
Reported 16S rDNA and ITS sequence similarity threshold for bacteria and fungi.
| Taxonomic Level | Bacteria 1 | Fungi | ||
|---|---|---|---|---|
| Filamentous Fungi 2 | Ascomycota Yeasts 3 | Basidiomycota Yeasts 3 | ||
| Phylum | 75% | - | - | - |
| Class | 78.5% | 80.9% | - | - |
| Order | 85% | 81.2% | - | - |
| Family | 86.5% | 88.5% | - | - |
| Genus | 94.5% | 94.3% | - | - |
| Species | 98.7% | 99.6% | 98.31% | 98.61% |
1 Values represent 16S rDNA sequence similarity thresholds reported by Yarza et al. [
Higher taxonomy of the culturable bacterial diversity from different Genlisea hispidula tissues based on bPTP analysis of the full-length 16S rDNA region.
| Taxonomic Level | Isolate Identity | Unique bPTP Species (#) | Total Cultured Isolates (#) | Plant Tissue Type | ||
|---|---|---|---|---|---|---|
| Leaf | Rhizophyll | Vesicle | ||||
| Genus | Acidocella | 1 | 9 | Yes | Yes | Yes |
| Bacillus | 2 | 2 | Yes | Yes | ||
| Burkholderia | 8 | 13 | Yes | Yes | Yes | |
| Herminiimonas | 1 | 1 | Yes | |||
| Jatrophihabitans | 1 | 1 | Yes | |||
| Methylobacterium | 1 | 2 | Yes | |||
| Microbacterium | 1 | 4 | Yes | |||
| Novosphingobium | 7 | 7 | Yes | Yes | ||
| Paenibacillus | 1 | 1 | Yes | |||
| Paraburkholderia | 5 | 13 | Yes | Yes | ||
| Rhodopseudomonas | 2 | 5 | Yes | |||
| Family | Alcaligenaceae | 8 | 8 | Yes | Yes | Yes |
| Nitrobacteraceae | 5 | 5 | Yes | |||
| Rhodospirillaceae | 1 | 1 | Yes | |||
| Sphingomonadaceae | 1 | 1 | Yes | |||
| Order | Burkholderiales | 1 | 1 | Yes | ||
| Micrococcales | 1 | 1 | Yes | |||
| Phylum | Pseudomonadota sp. | 1 | 1 | Yes | ||
| Totals | 48 | 76 | ||||
Higher taxonomy of the culturable fungal diversity from different Genlisea hispidula tissues based on bPTP analysis of the ITS region.
| Taxonomic Level | Isolate Identity | Unique bPTP Species (#) | Total Cultured Isolates (#) | Plant Tissue Type | ||
|---|---|---|---|---|---|---|
| Leaf | Rhizophyll | Vesicle | ||||
| Genus | Aspergillus sp. | 1 | 2 | Yes | ||
| Cladosporium spp. | 2 | 2 | Yes | |||
| Penicillium spp. | 8 | 12 | Yes | Yes | ||
| Saitozyma spp. | 11 | 14 | Yes | Yes | Yes | |
| Talaromyces spp. | 2 | 3 | Yes | Yes | ||
| Toxicocladosporium spp. | 3 | 5 | Yes | |||
| Trichomonascus sp. | 1 | 20 | Yes | Yes | Yes | |
| Order | Eurotiales sp. | 1 | 1 | Yes | ||
| Totals | 29 | 59 | ||||
Supplementary Materials
The following supporting information can be downloaded at
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