Noteworthy features of native orchid species used in this study. (A) The evergreen foliage rosettes of Goodyera tesselata in Huntingdon County, PA, Sept. 2016. (B) A flowering Platanthera blephariglottis in Luzerne County, PA, July 2017. (C) Sorba rods inoculated with Ceratobasidium sp. immediately before planting with G. tesselata seedlings. (D) An example of containerized P. blephariglottis seedlings used in this study.
Data for 2018 measurements of Goodyera tesselata in four different substrates with (+F) and without (–F) mycorrhizal fungus inoculation. (A) Leaves. (B) Survival. (C) Performance [mass (g) × survival].
Relationships between two measures of plant size in Goodyera tesselata and the abundance of the added mycorrhizal fungi in the substrate. (A) Number of leaves produced. (B) Rosette diameter. (C) Survival in +fungus pots. The relationship was similar among substrates, so the four substrates are shown combined, with substrate color coded (black = B, light green = substrate NZ, dark green = NZ/10W, blue = NZ/50W).
Data for 2018 measurements of Platanthera blephariglottis in four substrates with (+F) and without (–F) mycorrhizal fungus inoculation. (A) Leaves. (B) Survival. (C). Performance [mass (g) × survival].
Relationship between 2018 performance [plant mass (g) × survival] of Platanthera blephariglottis and abundance of inoculated fungus in four different substrates, indicated by color (black = B, light green = substrate NZ, dark green = NZ/10W, blue = NZ/50W): (A) B; (B) NZ; (C) NZ/10W; (D) NZ/50W.
Relationship between 2019 performance [plant mass (g) × survival] of Platanthera blephariglottis and abundance of inoculated fungus in four substrates: (A) B; (B) NZ; (C) NZ/10W; (D) NZ/50W.
The abundance of target fungus DNA for (A) Goodyera tesselata and (B) Platanthera blephariglottis obtained from each of four substrates with (+F) or without (–F) added target fungi.
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U.S. native temperate terrestrial orchids are of horticultural and conservation interest but are considered difficult to propagate from seed due to complex ecological requirements and a variable need for a mycorrhizal fungus. Although there has been significant research on germinating seeds and in vitro seedling development on a variety of temperate terrestrial orchid taxa from around the world, few studies have combined germination experiments with appropriate mycorrhizal fungi that support establishment and continued growth for purposes of ex situ collections development, conservation, or restoration. We conducted experiments with two species of asymbiotically propagated U.S. native orchids [Goodyera tesselata Lodd. and Platanthera blephariglottis (Willd.) Lindl.] to determine the effect of four substrates [Bog garden mix (peat:all-purpose sand) (B), New Zealand sphagnum (NZ), NZ sphagnum + 10% powdered tulip tree wood (NZ/W10), and NZ sphagnum + 50% powdered tulip tree wood (NZ/W50)] and whether inoculating with an appropriate mycorrhizal fungus grown on cellulose sorba rods would support orchid growth and survival in containers and subsequently in outdoor conditions. Morphological measurements and survival data were used in conjunction with real-time quantitative polymerase chain reaction to assess fungal abundance in containers and the impact of fungal presence on growth and survival characteristics. The addition of appropriate mycorrhizal fungi increased the growth and survival of both species across three (NZ, NZ/W10, and NZ/W50) of four substrates. The addition of a mycorrhizal fungus was not a universal solution to improving growth, but the addition resulted in increased abundance of the fungus and better plant performance. This novel experiment suggests that although addition of appropriate mycorrhizal fungi to orchids may increase performance, environmental and horticultural parameters also play an important role in successful orchid cultivation.
More than 220 temperate orchid species occur in North America north of Mexico and ≈150 are terrestrial, but the requirements for horticultural production and cultivation are known for few species (Luer, 1975; Swarts and Dixon, 2017). Horticultural requirements are well known for a few taxa, including Calopogon tuberosus and several species in the genus Cypripedium (Kauth et al., 2006, 2008, Steele, 1996, 2007; Whitlow, 1996). Information is incomplete or lacking for most of the other taxa of potential horticultural significance in genera such as Goodyera, Isotria, Platanthera, and Spiranthes. In addition to horticultural value, some terrestrial orchids are of federal or state conservation concern, and two genera have species of global conservation concern (Krupnick et al., 2013).
Guidance on growing some native orchids is available (Mathis, 2005; Seaton and Ramsey, 2005; Tullock, 2005), but none of those publications provide information on propagating native orchids in vitro in greenhouse, nursery, or garden conditions that include information on inoculation with an orchid mycorrhizal fungus as part of the propagation process. Research has demonstrated the importance of orchid mycorrhizal fungi (Rasmussen and Rasmussen, 2009), and successful conservation of native orchids likely requires the inclusion of appropriate fungi in the propagation process (Swarts and Dixon, 2017). Difficulties in growing native orchids from seed, for example, may stem from lack of appropriate mycorrhizal fungi to support seedlings in the establishment phase. Efficient, reproducible protocols that combine seed germination, seedling production, and practices for successful acclimatization with appropriate orchid mycorrhizal fungi are almost universally lacking for native orchids in support of horticultural, conservation, and restoration initiatives.
The key to successful cultivation of most native orchids is determining the requirements of associated mycorrhizal fungi at one or more key life history stages (Swarts and Dixon, 2017). In nature, all orchids associate with one or more orchid mycorrhizal fungi that serve as a source of nutrition during all life history stages (Gebauer et al., 2016; Jacquemyn et al., 2021; McCormick et al., 2012; McCormick and Jacquemyn, 2014; Miura et al., 2018; Rasmussen, 1995; Rasmussen and Rasmussen, 2009; Swarts and Dixon, 2017). Some orchid taxa have been so successful at interacting with fungi that they have replaced their ability to photosynthesize and obtain all nutrition they need from fungi at every stage in their life cycle. Orchids that have evolved this strategy are termed mycoheterotrophic (Jacquemyn et al., 2021; Merckx, 2013; Miura et al., 2018). The relationship between orchids and orchid mycorrhizal fungi varies among taxa and ranges from general to highly specific, and species with highly specific fungal relationships are generally more difficult to cultivate (e.g., fully mycoheterotropic orchids) (Rasmussen, 1995). Although it is not commonplace practice among orchid horticulturists, culturing and using the appropriate orchid mycorrhizal fungus to aid in cultivation could help with the transition from in vitro to ex vitro conditions and increase the likelihood of successful long-term cultivation.
Two methods of in vitro orchid seed germination, symbiotic and asymbiotic, impact the possible association of seeds and seedlings with mycorrhizal fungi in in vitro and ex vitro environments. Asymbiotic germination does not include an orchid mycorrhizal fungus, and symbiotic germination occurs with seeds that are being germinated in the presence of a mycorrhizal fungus (Rasmussen, 1995; Swarts and Dixon, 2017). To date, commercial production of temperate terrestrial orchids relies on asymbiotic germination using either physiologically mature or immature seeds, followed by seedling development on nutrient-rich media in absence of a mycorrhizal fungus. Asymbiotic protocols have been most successfully developed for species of Cypripedium (Kauth et al., 2008; Klavina et al., 2009; Zeng et al., 2013). Although few in vitro production studies have included orchid mycorrhizal fungi, there is evidence that cultivated plants can associate with mycorrhizal fungi when appropriate cultivation conditions are met, and the fungus is present in the substrate. High mortality in the in vitro–ex vitro acclimatization phase of asymbiotically propagated native orchids is still commonplace, a possible result of the absence of an appropriate mycorrhizal fungus, inappropriate substrate and environmental conditions, or both.
Symbiotic germination uses a known orchid mycorrhizal fungus to support in vitro seed germination (Rasmussen, 1995). The advantage of this method is that protocorms and seedlings are already paired with a mycorrhizal fungus at the time of acclimatization and may be better equipped to survive transplanting. However, mycorrhizal fungi are typically harvested from adult plants and there is some evidence that different species fungi associate with seedlings in the same species (McCormick et al., 2021). Furthermore, some orchid mycorrhizal fungi are also obligately ectomycorrhizal with trees and cannot be cultivated with the same ease as saprotrophic fungi (Rock-Blake et al., 2017; Whigham et al., 2021). Also, multiple symbiotic seed germination studies of temperate terrestrial orchids report successful germination, but reports of continued development to the stage of leaf-bearing seedling large enough to acclimatize to ex vitro conditions are rare (Sharma et al., 2003; Zettler and McInnis, 1992) Although both methods have their advantages, challenges to improving the rate of success cultivating seedlings resulting from either germination type.
Numerous container substrates have been formulated for the successful cultivation of temperate terrestrial orchids that consist of a variety of organic and inorganic components (Jørgensen and Andersen, 1998; Swarts and Dixon, 2017). These are based on industry-standard components that are readily available and physical properties of the medium including pH, ratio of water to air pore space, rate of decomposition, and electrical conductivity. Composted bark and wood fiber are common components of commercial substrates with specific characteristics that promote growth and establishment of terrestrial orchids (Bustam et al., 2016; Whigham et al., 2002). Decomposing wood of various types has been shown to have a stimulatory effect on seed germination and establishment, but relatively few types of processed or decomposed wood have been tested for their efficacy in supporting orchid growth in container culture (Rasmussen and Whigham, 1993). Powdered wood of Liriodendron tulipifera, for example, has been shown to be effective in supporting symbiotic germination of some native orchid species and growing fungus in sterile petri dishes, but to our knowledge it has never been used as an additive to container substrates to support both orchids and mycorrhizal fungus growth.
Recent research has demonstrated that the quantity of orchid mycorrhizal fungus in substrates may be as important as the presence or absence of the fungus in supporting orchid growth and survival (McCormick et al., 2018, 2021; Rock-Blake et al., 2017; Whigham et al., 2021). To date, however, the importance of fungal abundance has never been used in horticulturally based research on native orchids. In this first-of-its-kind experiment, we quantified the effect of four substrates with and without an appropriate orchid mycorrhizal fungus on the growth and survival of two native orchids (Goodyera tesselata and Platanthera blephariglottis) in containers and subsequently in outdoor garden conditions. This is the first study in a series of applied conservation horticulture experiments that we hope to develop into protocols for large-scale propagation and conservation of native orchids that result in establishment of genetically diverse, sustainable ex situ conservation collections in public gardens throughout the United States and Canada and supports goals of the North American Orchid Conservation Center.
Goodyera tesselata Lodd. is distributed in the northeastern and northern Great Lakes regions of the United States where it grows in coniferous and mixed deciduous forest (Fig. 1A). Plants typically grow in a thick layer of forest duff in dry-mesic forest under a canopy of pines (Pinus spp.) or hemlock (Tsuga canadensis). Platanthera blephariglottis (Willd.) Lindl. is also a species of the northeastern and northern Great Lakes regions but grows in live sphagnum moss of open bogs that are permanently saturated (Fig. 1B) and has a distribution that extends south to Florida and west to Texas. Numerous other orchid species as well as Ericaceous plants are sympatric in the same habitats where both species occur.
Citation: HortScience 57, 5; 10.21273/HORTSCI16509-22
Mature seed capsules of P. blephariglottis were collected from six individuals growing at the Valmont Bog in Luzerne County, PA, on 19 Oct. 2016. Capsules of Goodyera tesselata were collected from Huntingdon County, PA, on 21 Sept. 2016. Seeds were air-dried and, after 4 weeks, were removed from dried capsules, cleaned of debris, and stored in continual darkness at 3 °C and 30% relative humidity in glass vials until used.
Seeds were surface sterilized and scarified for 20 min in a 9:1 solution of deionized distilled (dd) water to 5.25% NaOCl (Clorox bleach, The Clorox Co., Oakland, CA) and two drops of Tween 20. Solutions containing seeds were agitated during the 20-min period and vessels containing seeds were transferred to a sterile working space in a laminar flow hood, removed from the sterilization solution, and rinsed three times in sterile dd water. Two commercially available asymbiotic orchid seed germination media, M551 and P723 (Phytotechnology Laboratories, Shawnee Mission, KS) were prepared and the pH adjusted to 5.8 with KOH before autoclaving at 103.4 kPa for 20 min at 122 °C. The media was poured into 6-cm-diameter Petri dishes (≈12.5 mL medium/plate; Thomas Scientific, Swedesboro, NJ) in a sterile hood, cooled, then stored for use in germination experiments.
Approximately 150 surface sterilized seeds were inoculated onto each petri dish in a sterile hood using a sterile microspatula. Plates were sealed with parafilm M (Bemis Co., Neenah, WI) and incubated under 0/24 h L/D photoperiod at 25 ± 3 °C. Eight replicate plates were used for each germination medium. Plates were observed every 2 weeks after sowing, and germination was recorded on a 0 to 4 scale based on stage of germination as follows. Stage 0 was an ungerminated seed with a swollen embryo, stage 1 was a seed with a swollen embryo that had ruptured the testa, stage 2 was a fully developed protocorm, stage 3 was a protocorm with a visible shoot primordium, and stage 4 was a protocorm with developing shoot and root primordia seedling stage. After reaching stage 4, single seedlings were transferred to individual 25 × 150-mm test tubes (DWK Life Sciences, Millville, NJ) with 13 mL of T839 orchid medium (Phytotechnology Laboratories). Developing seedlings were placed in a growth chamber for 52 weeks with a 16-h photoperiod set at 21 ± 3 °C. Seedlings were then removed from test tubes and the agar-based medium gently brushed from the roots. Seedlings were placed in plastic bags with dampened New Zealand sphagnum moss (a common substrate for cultivating orchids; Sphagnum cristatum harvested from the Southwestern coast of the South Island of New Zealand and air-dried; Besgrow Co., Christchurch, NZ) and placed in a cooler at 3 ± 2 °C for 120 d for vernalization to alleviate shoot dormancy before planting the experiment.
Lateral roots of both taxa were collected in Autumn 2017 from the same locations, and populations where seeds were collected. Fungal pelotons from the roots were isolated and cultured on E-medium at the Smithsonian Environmental Research Center (SERC) (Caldwell et al., 1991). Fungi growing on the E-medium were transferred to sorba rods saturated with liquid E-medium in 200-mL deli tubs. Sorba rods are a cellulose-based product used for environmental and research applications that support the growth of orchid mycorrhizae but are no longer manufactured. Readily available cotton balls are a suitable substitute. An equal number of sorba rods that served as controls were placed in the culture media that was not inoculated with the orchid mycorrhizal fungi. The containers with the sorba rods were grown in the laboratory until they were taken to Longwood Gardens and used in the first phase of the experiment.
Container substrate ingredients were assembled at Longwood Gardens using a combination of commercially available and specially prepared materials. The peat used was the dark brown, fibrous material resulting from the decomposition of sphagnum moss that is harvested in Canada and readily available in the United States in compressed bales (Premier Horticulture Inc., Quakertown, PA). Peat was soaked with dH2O before mixing and the pH was unadjusted. All-purpose sand is washed, graded coarse sand used for a wide variety of horticultural purposes (Quikrete, Atlanta, GA). New Zealand sphagnum was mentioned earlier and differs from Canadian peat in that the living sphagnum moss is harvested and dried, resulting in a more fibrous, coarser substrate. Ground tuliptree (Liriodendron tulipifera) wood was custom made by harvesting saplings with 2- to 8-cm stem diameters from natural areas of Longwood Gardens. Leaves, stems, and bark were immediately removed from freshly harvested saplings and the remaining trunks air-dried at 27 °C for 6 weeks. The dried material was the ground to powder-like consistency using a Thomas Wiley mini cutting mill (Thomas Scientific Inc.).
Four substrates were prepared using the previously mentioned ingredients: (B) Bog garden mix (1:1 peat:all-purpose sand); (NZ) New Zealand sphagnum; (NZ/10W) NZ Sphagnum + 10% by volume of ground tulip tree (L. tulipifera) wood; (NZ/50W) NZ sphagnum + 50% by volume of ground tulip tree wood. All media were prepared at Longwood Gardens and autoclaved at 103.4 kPa for 20 min at 122 °C before use. For P. blephariglottis, we prepared 100, 8.50 × 9.75 cm square plastic containers (Dillen Products, Inc., Middlefield, OH). Each container received a single asymbiotically grown orchid seedling and a sorba rod. Half of the containers received sorba rods that were colonized by the fungus that had been isolated from a root of P. blephariglottis (identified as an unnamed species of Tulasnella), and half were uncolonized controls. For G. tesselata, we prepared fifty 5.75 × 9.75-cm square plastic containers of each of the same four substrates (Dillen Products, Inc.). Each pot received one asymbiotically grown seedling and a sorba rod. Half of the sorba rods were colonized by fungi from a root of an adult G. tesselata (identified as an unnamed species of Ceratobasidium) and the other half were not inoculated (Fig. 1C). Fungal samples used in this study are stored long-term at SERC as part of a comprehensive, living orchid fungal bank developed to facilitate native orchid research and conservation efforts.
Containers for both species were randomly placed in trays on tables in the greenhouse at Longwood Gardens (Kennett Square, PA) on 21 May 2018 under 50% shadecloth with an average light intensity of 125 µmol·m−2·s−1 as measured at 1300 hr once a week for the duration of the study. Each tray included only containers that were inoculated or only containers that were uninoculated and included only a single orchid species to prevent accidental colonization of uninoculated containers (Fig. 1D). Container locations within a tray and tray locations on the bench were rearranged once per week during the growing season. While in active growth, containers were hand-watered with clear water every 3 to 5 d in conjunction with readings from a soil moisture tensiometer. All substrates required watering on the same schedule. Irrigation water was supplied from the water treatment and storage system at Longwood Gardens. The water quality is comparable to municipal water supplies but has lower levels of dissolved chlorine, fluorine, calcium carbonate, and dissolved salts because it is managed to support horticultural operations. Seedlings were not fertilized at any point during this study because there is evidence that fertilizers may be harmful to or change the orchid–fungus relationship, resulting in a potentially confounding factor for this type of research (Rasmussen, 1995). All plants were measured twice per year: the first at anthesis, or when the new growth had matured, and the second after the plants went dormant.
The number of leaves, length and width of longest leaf, and presence or absence of inflorescence were measured for each plant. When plants were removed from the containers and planted in an outdoor bed (described subsequently) and again at the end of the experiment, the number of roots, length, and width (diameter) of each root, and reproductive status of each plant were recorded. Individual root measurements on each plant were combined to determine total root length and mean root width. Plants were grown in the greenhouse for 5 months and then out-planted into a specially designed in-ground planting bed filled with substrate B. Plants were only given supplemental water at the time of planting.
Five months and 1 year after the greenhouse experiments began, four small (≈0.5 g) subsamples of substrate were removed from each container, with one subsample from each side of the container combined and transported to SERC for storage at –80 °C and analysis as described subsequently.
The frozen substrate samples were freeze dried at –60 °C in a vacuum freeze drier. Dried substrates were ground into a powder using a mortar and pestle, and each 0.25-g subsample was placed into wells of a 96-well plate and DNA was extracted using DNEasy Ultra-Soil DNA extraction kits (Qiagen Sciences, Germantown, MD). DNA was extracted per the kit protocol, and each resulting extract was quantified using a Nanodrop spectrophotometer. To quantify the DNA of each inoculated fungus, DNA isolated from each substrate sample was analyzed using quantitative real-time polymerase chain reaction (qPCR). Each qPCR reaction included 40 ng of DNA + sterile distilled water to total 10 µL, 1.25 µL of each primer, 0.1 µL bovine serum albumin (BSA), 12.5 µL IQ-SybrGreen Master Mix, and 0.05 µL Rox internal standard. Each sample was run in triplicate and no-template controls and quantitative standards, consisting of serial dilutions of 0.1, 0.01, 0.001, 0.0001, and 0.00001 ng/µL of DNA from a pure culture of each target fungus, were included on each plate. PCR for the samples of P. blephariglottis was conducted with Tulasnella-specific primers ITS5/ITS-4Tul and qPCR for samples from G. tesselata was conducted with Ceratobasidium-specific primers CeTh1/CeTh4. PCR conditions were 96 °C for 6 min, followed by 41 cycles of 96 °C for 30 s, annealing temperature (54 °C for Tulasnella, 60 °C for Ceratobasidium) for 15 s, and 72 °C for 30 s, followed by a melting curve. Melting curves were checked for the presence of a single peak, indicating that only the target fungus was amplified. Five samples of each that had abundant target fungi were subjected to Sanger sequencing to verify that the amplified DNA matched our target fungi.
For both orchids, analysis of variance (ANOVA), implemented in Systat 11 for Windows (Systat Software, Inc.), was used to determine whether there were any differences between the treatments in fungal abundance. In this analysis, multiple variables were used as dependent variables, and P values were corrected for multiple comparisons using Bonferroni corrections. For G. tesselata, the dependent variables were the number of leaves and rosette diameter in Aug. 2018. For P. blephariglottis, the dependent variables were number of leaves in 2018 and 2019 and plant mass, total root length, and mean root width in 2019. However, evaluation of propagation success is not solely an evaluation of growth or survival, but rather a combination of the two. To assess propagation success for both species, we created a statistic, which we called Performance, that weighted growth by mean survival for that treatment. This statistic was only calculated for 2018 for G. tesselata because survival by the end of the experiment was zero.
In both analyses, substrate type and the presence or absence of the fungus (+F/–F) were included as main effects. The initial analyses indicated that the fungus did not always successfully establish in the pots, making the fungus treatment effect less clear than the effect of fungus abundance, as estimated by fungus DNA. For both species, we conducted a second set of analyses with the same dependent variables (number of leaves, whorl diameter) within each fungus treatment to examine the effect of fungus abundance. These analyses were set up as ANOVAs with substrate as the main independent variable and fungal DNA quantity as a covariate. The relative abundance of each of the two fungi in the different substrates was compared using an ANOVA with species and substrate as main effects and using the significance of the interaction between species and substrate as indicative of fungal growth differences. In all analyses, the quantity of DNA (ng DNA/g dry substrate) was log transformed to improve normality of the data and reduce the influence of high-value outliers. Because the amount of DNA that was extracted differed between the four substrate types, the comparisons between fungus quantity and orchid performance were conducted separately for each substrate. Variable mass/volume relationships and variable functional mass, as well as variable substrate density, surface available for colonization, and DNA extraction efficiency, are known issues with trying to compare fungus quantity among different substrates, so the extent to which different fungus DNA concentrations among substrates reflect biologically meaningful comparisons is unclear. For this reason, it is more meaningful to assess the effect of fungus abundance within each substrate, rather than among them. Significance values were corrected to account for multiple comparisons. Other measures of growth (number of roots, root length, 2019 metrics) were not available for G. tesselata because all plants died before measurement. Analogous to analyses of growth metrics, survival was analyzed using logistic regressions, first of substrate and fungus (+F/–F) and then of target fungus DNA quantity [ln(ng DNA/g dry substrate)] within each fungus treatment (+F/–F).
For G. tesselata, neither the number of leaves or rosette diameter differed significantly among substrates or whether a fungus was added to the containers (Table 1). Survival was significantly higher in the NZ (Z = 2.358, P = 0.02) and NZ/W10 (Z = 1.954, P = 0.05) substrates than in other media (Fig. 2). When the analysis was based on fungal abundance the +F treatment was significantly related to the number of leaves (P = 0.006), the diameter of the leaf whorl (P = 0.03), and survival after 5 months (P < 0.001) (Fig. 2).
Citation: HortScience 57, 5; 10.21273/HORTSCI16509-22
For P. blephariglottis, there were nonsignificant differences in number of leaves in 2018 (P ≤ 0.1) or 2019 (P ≤ 0.1), plant mass in 2018 (P ≤ 0.07) or 2019 (P ≤ 0.09) with respect to substrate, and the presence or absence of a fungus, or the interaction between the two (Figs. 3A and 4A). Survival in 2018 (Fig. 3B) and 2019 (Fig. 4B) was significantly different among substrates (P ≤ 0.01) but not with whether fungi were added (2018: P = 0.36; 2019: P = 0.15).
Citation: HortScience 57, 5; 10.21273/HORTSCI16509-22
Citation: HortScience 57, 5; 10.21273/HORTSCI16509-22
When survival was combined with growth for P. blephariglottis, performance in substrates NZ, NZ/W10, and NZ/W50 was higher when fungi were added, compared with when no fungi were added in both 2018 (Fig. 3C) and 2019 (Fig. 4C). In substrate B the opposite was true. The addition of fungi resulted in the lowest orchid survival when plants were placed in substrate B. Survival and growth were highest in the containers with NZ/W50.
When considering only the containers to which fungi had been added, the abundance of fungus was positively related to the performance of P. blephariglottis plants in substrates NZ and NZ/W50 or NZ, NZ/W10, and NZ/W50, but not significant in substrate B. The significance differed somewhat between 2018 (Fig. 5) and 2019 (Fig. 6).
Citation: HortScience 57, 5; 10.21273/HORTSCI16509-22
Citation: HortScience 57, 5; 10.21273/HORTSCI16509-22
Differences in the amount of target fungus DNA that was recovered were highest for the P. blephariglottis fungus compared with the G. tesselata fungus among substrates (G. tesselata, P = 0.161; P. blephariglottis: P < 0.001) and as an interaction between substrate and fungus addition (G. tesselata: P = 0.161, P. blephariglottis: P < 0.001). However, differences were clear for both between containers to which fungi were added and those to which uninoculated sorba rods were added (both P < 0.001) (Fig. 7A and B). The abundance of both fungi was least in the substrate B.
Citation: HortScience 57, 5; 10.21273/HORTSCI16509-22
This experiment was designed to determine whether the addition of appropriate mycorrhizal fungi could augment survival and growth of containerized, greenhouse-grown individuals of two native terrestrial orchid species in four substrates. The addition of a species-specific mycorrhizal fungus increased the growth and survival of both species across three of four substrates. The response of the orchids to the addition of the orchid mycorrhizal fungus depended on fungal abundance in the containers to which fungi were added. Generally, orchid performance increased with an increase in fungal abundance. However, the study also demonstrated that fungus addition is not a universal solution to improving growth and survival of native terrestrial orchids. In substrate B, addition of mycorrhizal fungi resulted in decreased growth and survival of both orchids. In the other three substrates, growth and survival increased with increasing fungal abundance. This result demonstrates that horticultural parameters (i.e., substrate type) also play a role in successful cultivation of orchids and fungi. Substrate B was clearly not a suitable substrate for the orchid mycorrhizal fungus of either species in containers, most likely because the fungi were either not able to obtain resources from the peat to support orchid growth and survival or obtained too many resources and outgrew the orchids.
For P. blephariglottis, growth and survival differences with or without orchid mycorrhizal fungi during growth in the greenhouse were continued and, to some degree, magnified after out-planting and another year of growth in outdoor beds filled with substrate B (P. Zale, unpublished data). This suggests that either growth differences that developed in the greenhouse contributed to additional later growth or that beneficial mycorrhizal associations established in the greenhouse were maintained and continued to be beneficial for out-planting success and that media unsuitable for container cultivation of native orchid can be useful for different planting environments. It is likely that early growth advantages, mycorrhizal establishment, and improved planting environment contributed to later and continued success in both fungus (F+/F–) treatments. In containers, it is possible that seedling and fungus performance was affected by large inorganic content of the substrate B. This is likely at least partly attributable to the large amount of substrate B that is inorganic (i.e., sand) and does not provide any resources that support fungal growth. It may also suggest that this substrate enabled the fungus to outgrow the orchid or perhaps in some way prevented successful mycorrhizal association, while increasing competition for nutrients. Continued growth of the plants indicate that substrate B is ideal for long-term outdoor cultivaton of P. blephariglottis, reinforcing the importance of trialing substrates in different environments and suggesting that planting seedlings directly into specially constructed outdoor beds of substrate B, or another substrate, may be preferable to a greenhouse container acclimatization phase. In terms of long-term performance, the opposite was true for G. tesselata. No plants of G. tesselata survived to be measured in 2019, likely due to excessively warm temperatures in the greenhouse, indicating that environmental parameters, such a root zone temperature, play a major role in successful acclimatization, and like P. blephariglottis, experimentations with directly acclimatizing plants to outdoor conditions needs further experimentation.
The growth of introduced fungi varied among containers. In some containers, fungi failed to grow and, if not accounted for, these differences in growth obscured the benefits of introducing fungi. Assessing the abundance of introduced fungi also allowed us to demonstrate that the Ceratobasidium fungus from G. tesselata and the Tulasnella fungus from P. blephariglottis differed in which media they grew best. In particular, the highest abundance of G. tesselata fungus DNA was measured in pots with the substrate NZ to which fungi were added, whereas DNA from P. blephariglottis fungi was most abundant when fungi were added to NZ/W50. Assuming that the major difference among the substrates was in fungus DNA recovery, it would have affected recovery of DNA from the two fungi similarly, so the differences in relative abundance of target fungus DNA (P = 0.002) among substrates likely reflect real differences in growth preferences of the two fungi. These two fungal genera differ in many ways but perhaps most strikingly in their ability to use lignin and nitrogen. Ceratobasidium fungi can decompose lignin, whereas Tulasnella have no laccase genes (Kohler et al., 2015; Nurifadilah et al., 2013). Ceratobasidium can also use nitrate and nitrite forms of nitrogen, whereas Tulasnella lack any nitrate or nitrite handling genes (Kohler et al., 2015; M. McCormick, unpublished). These genetic differences correspond to differences in ability to use different sources of carbon and nitrogen found by Nurifadilah et al. (2013).
In addition to genetic differences between fungi used for this study and substrate types, the method used to inoculate substrates may also influence the ability of the fungus to colonize the substrate and pair with the asymbiotically propagated orchids. Other methods of fungal inoculation mentioned in the scientific literature include a fungal broth made from fungi cultivated on potato dextrose agar plates, leaf litter inoculated under controlled conditions, and millet seed inoculated with orchid fungus under controlled conditions (Higaki et al., 2017; Liu et al., 2020; Quay et al., 1995; Smith et al., 2007). In all studies, seed germination or an increase in plant dry mass were evident, indicating that numerous methods of inoculating orchid seeds or plants are viable. The variation in amount of fungus among pots within a treatment and across substrates could also be an artifact of inoculation type and suggests that, although sorba rods may be suitable to inoculate orchid substrates, other methods should also be tested to determine which is the most effective.
Fungal inoculation of asymbiotic orchid seedlings at the onset of the ex vitro cultivation phase may have species specific advantages compared with introducing mycorrhiza at the seed germination stage (symbiotic propagation) (Bustam et al., 2014; Castillo-Pérez et al., 2021; Kauth et al., 2008). Symbiotic orchid seedlings are grown in vitro on media that specifically support and regulate the growth of mycorrhizal fungi. Some evidence indicates that this may have a negative impact on the development of the orchids, resulting in smaller seedlings that may not be able to survive ex vitro acclimatization (Bae et al., 2015; Lauzer et al., 2007; Malmgren, 1996; Steele, 1996; Swarts and Dixon, 2017; Whitlow, 1996; Yannetti, 1996; Zettler and McInnis, 1992; Zettler et al., 2000). This could be because certain terrestrial orchids exhibit fungal specificity at different life history stages and that fungi isolated from the roots of mature plants may not be the correct fungus to promote germination and development of seedlings (Li et al., 2022; McCormick et al., 2021; McCormick and Jacquemyn, 2014; Rasmussen, 1995). Since asymbiotic seedlings are larger at the time of ex vitro planting, pairing them with fungi isolated from adult plants at this stage may increase the success of fungal pairing and increase performance. These factors support continued research to optimize ex vitro symbiotic cultivation experiments that explore varying techniques of fungal inoculation of orchid seedling.
The results of this experiment provide guidance for future research efforts and a framework for developing greenhouse and nursery acclimatization protocols that will benefit efforts to propagate and conserve native orchids. Many unanswered questions remain, but a few topics emerge from this project that require further attention and effort. First, the type of substrate is important, and the most suitable substrates for native orchids needs to be determined. The experiment also demonstrated the importance of the orchid mycorrhizal fungi. Only a few experiments have incorporated orchid mycorrhizal fungi into germination and growth studies of orchids. In some instances, orchid growth was greater when the orchid mycorrhizal fungus was present; however, multiple factors influence the successful propagation and production of native orchids (Alghamdi, 2019; Durán-López et al., 2019). The following question should be addressed in future studies of this type: 1) What are the most effective methods for inoculating orchid container substrates with appropriate fungi? 2) What is the role of germination type (asymbiotic vs. symbiotic) in successful orchid–fungus pairing? 3) Are the fungi used to inoculate container substrate pairing with orchid seedlings? and 4) What are the environmental parameters that promote optimal orchid and fungus growth and development?
Noteworthy features of native orchid species used in this study. (A) The evergreen foliage rosettes of Goodyera tesselata in Huntingdon County, PA, Sept. 2016. (B) A flowering Platanthera blephariglottis in Luzerne County, PA, July 2017. (C) Sorba rods inoculated with Ceratobasidium sp. immediately before planting with G. tesselata seedlings. (D) An example of containerized P. blephariglottis seedlings used in this study.
Data for 2018 measurements of Goodyera tesselata in four different substrates with (+F) and without (–F) mycorrhizal fungus inoculation. (A) Leaves. (B) Survival. (C) Performance [mass (g) × survival].
Relationships between two measures of plant size in Goodyera tesselata and the abundance of the added mycorrhizal fungi in the substrate. (A) Number of leaves produced. (B) Rosette diameter. (C) Survival in +fungus pots. The relationship was similar among substrates, so the four substrates are shown combined, with substrate color coded (black = B, light green = substrate NZ, dark green = NZ/10W, blue = NZ/50W).
Data for 2018 measurements of Platanthera blephariglottis in four substrates with (+F) and without (–F) mycorrhizal fungus inoculation. (A) Leaves. (B) Survival. (C). Performance [mass (g) × survival].
Relationship between 2018 performance [plant mass (g) × survival] of Platanthera blephariglottis and abundance of inoculated fungus in four different substrates, indicated by color (black = B, light green = substrate NZ, dark green = NZ/10W, blue = NZ/50W): (A) B; (B) NZ; (C) NZ/10W; (D) NZ/50W.
Relationship between 2019 performance [plant mass (g) × survival] of Platanthera blephariglottis and abundance of inoculated fungus in four substrates: (A) B; (B) NZ; (C) NZ/10W; (D) NZ/50W.
The abundance of target fungus DNA for (A) Goodyera tesselata and (B) Platanthera blephariglottis obtained from each of four substrates with (+F) or without (–F) added target fungi.
Contributor Notes
We thank Hye Woo Shin and Ashley Clayton for assistance in the greenhouse and laboratory. The Pennsylvania Department of Conservation of Natural Resources Bureau of Forestry and the North Branch Land Trust granted permission and access to seed collection sites.
P.J.Z. is Associate Director, Conservation, Plant Breeding, and Collections.
M.K.M. is a Research Scientist.
D.F.W. is a Senior Botanist.
P.J.Z. is the corresponding author. E-mail: pzale@longwoodgardens.org.
Noteworthy features of native orchid species used in this study. (A) The evergreen foliage rosettes of Goodyera tesselata in Huntingdon County, PA, Sept. 2016. (B) A flowering Platanthera blephariglottis in Luzerne County, PA, July 2017. (C) Sorba rods inoculated with Ceratobasidium sp. immediately before planting with G. tesselata seedlings. (D) An example of containerized P. blephariglottis seedlings used in this study.
Data for 2018 measurements of Goodyera tesselata in four different substrates with (+F) and without (–F) mycorrhizal fungus inoculation. (A) Leaves. (B) Survival. (C) Performance [mass (g) × survival].
Relationships between two measures of plant size in Goodyera tesselata and the abundance of the added mycorrhizal fungi in the substrate. (A) Number of leaves produced. (B) Rosette diameter. (C) Survival in +fungus pots. The relationship was similar among substrates, so the four substrates are shown combined, with substrate color coded (black = B, light green = substrate NZ, dark green = NZ/10W, blue = NZ/50W).
Data for 2018 measurements of Platanthera blephariglottis in four substrates with (+F) and without (–F) mycorrhizal fungus inoculation. (A) Leaves. (B) Survival. (C). Performance [mass (g) × survival].
Relationship between 2018 performance [plant mass (g) × survival] of Platanthera blephariglottis and abundance of inoculated fungus in four different substrates, indicated by color (black = B, light green = substrate NZ, dark green = NZ/10W, blue = NZ/50W): (A) B; (B) NZ; (C) NZ/10W; (D) NZ/50W.
Relationship between 2019 performance [plant mass (g) × survival] of Platanthera blephariglottis and abundance of inoculated fungus in four substrates: (A) B; (B) NZ; (C) NZ/10W; (D) NZ/50W.
The abundance of target fungus DNA for (A) Goodyera tesselata and (B) Platanthera blephariglottis obtained from each of four substrates with (+F) or without (–F) added target fungi.
