Regular issues
Special Issues

Southeastern Naturalist
    SENA Home
    Range and Scope
    Board of Editors
    Editorial Workflow
    Publication Charges

Other EH Journals
    Northeastern Naturalist
    Caribbean Naturalist
    Urban Naturalist
    Eastern Paleontologist
    Eastern Biologist
    Journal of the North Atlantic

EH Natural History Home

Restoration of the Endangered Ruth’s Golden Aster (Pityopsis ruthii)
Phillip A. Wadl, Arnold M. Saxton, Geoff Call, and Adam J. Dattilo

Southeastern Naturalist, Volume 17, Issue 1 (2018): 19–31

Full-text pdf (Accessible only to subscribers.To subscribe click here.)


Site by Bennett Web & Design Co.
Southeastern Naturalist 19 P.A. Wadl, A.M. Saxton, G. Call, and A.J. Dattilo 22001188 SOUTHEASTERN NATURALIST Vo1l7.( 117):,1 N9–o3. 11 Restoration of the Endangered Ruth’s Golden Aster (Pityopsis ruthii) Phillip A. Wadl1,*, Arnold M. Saxton2, Geoff Call3, and Adam J. Dattilo4 Abstract - Pityopsis ruthii, Ruth’s Golden Aster, is an endangered herbaceous perennial that is endemic to small sections of the Hiwassee and Ocoee Rivers in the southeastern US. Our objective was to test the effect of bonded fiber matrix (BFM) on establishment and fecundity of Ruth’s Golden Aster in order to develop a robust restoration protocol. We augmented existing populations with plants grown from achenes collected at each restoration location. We monitored plantings through 3 growing seasons by measuring stem number, stem height, leaf number, flowering incidence, and number of flower heads per plant in the spring and fall of each season. We assessed survival at 1 month post-planting. We randomly assigned plants at each location to a treatment (BFM vs. no BFM) for analysis as a randomized complete-block design. Germination rate of filled seeds, number of acclimated seedlings, and percent of seedlings planted after 14 days of acclimatization differed significantly across sites. Survival was significantly higher at 1 month, fall year 1, spring/fall year 2, and spring year 3 for the plants mulched with BFM compared to the control. However, there were no significant differences between treatment for stem number, stem height, leaf number, flowering incidence, or final 3-year survival. The methods developed herein represent a major step towards meeting the recovery-plan objective of developing the ability to establish Ruth’s Golden Aster on suitable habitat. Herein, we provide a framework for augmentation or restoration of critical populations threatened by extirpation. Introduction Pityopsis ruthii (Small) Small (Ruth’s Golden Aster) is a diminutive (10–30 cm tall), fall-flowering member of the Asteraceae that occurs only in cracks of phyllite boulders (Semple 2006). This species can persist in shaded situations, but flowering, seed set, and establishment of juvenile plants is most successful in open habitats where plants receive full sun for a significant portion of the day (Moore et al. 2016, White 1977). Ruth’s Golden Aster is tolerant of prolonged drought (Moore et al. 2016) and inundating high-flow events (A.J. Dattilo, unpubl. data), but the species is a very poor competitor when it becomes established in areas of deeper soils that build up on the boulder complex or outside of the exposed boulder complexes where healthy populations occur (Cruzan and Beaty 1998, USFWS 2012). In the southeastern US, where sites with sufficient soil depth generally succeed to forest in lieu of anthropogenic disturbance, periodic high river-flow and cyclical 1US Department of Agriculture, Agricultural Research Service, US Vegetable Laboratory, 2700 Savannah Highway, Charleston, SC 29414. 2Department of Animal Science, University of Tennessee, 2506 River Drive, 232 Brehm Animal Science Building, Knoxville, TN 37996. 3US Fish and Wildlife Service, 446 Neal Street, Cookeville, TN 38501-4027. 4Biological Compliance, Tennessee Valley Authority, West Tower 11B-K, 400 West Summit Hill Drive, Knoxville, TN 37902. *Corresponding author - Manuscript Editor: Richard Baird Southeastern Naturalist P.A. Wadl, A.M. Saxton, G. Call, and A.J. Dattilo 2018 Vol. 17, No. 1 20 drought maintain the niche occupied by Ruth’s Golden Aster. The confluence of appropriate geology and disturbance regime is very rare on the landscape, making the species inherently vulnerable to extinction caused by natural and anthropogenic forces capable of significantly altering its habitat. Plants grow in small crevices of massive rock outcrops situated between the open river channel and the adjacent forest slopes. It is a narrowly distributed, herbaceous perennial plant that occurs only along small reaches of the Hiwassee (~4 km) and Ocoee (~2.5 km) Rivers in Polk County, TN. All occurrences of Ruth’s Golden Aster occur downstream of Tennessee Valley Authority (TVA) dams in sections of river that are intensively managed for flood control and electricity generation. The species is listed as endangered under the Endangered Species Act of 1973 and is considered critically imperiled (G-1; NatureServe 2015). Although modification of natural river flows has been implicated in population decline of this species (Bowers 1972; Thomson and Schwartz 2006; USFWS 1992, 2012; White 1977; Wofford and Smith 1980), there are no quantitative studies that address how river flows interact with Ruth’s Golden Aster. Regardless of the net effect of river management, the species has a very narrow range in a habitat that is subjected to extreme conditions. These combined factors make Ruth’s Golden Aster vulnerable to isolated events that have the potential to extirpate or severely degrade 1 or more populations. The ability to successfully establish plants in unoccupied suitable habitat is a viable option for reducing extinction risk (Pavlik 1996) and is a stated objective necessary for the recovery of the species (USFWS 1992). Falk et al. (1996) defined 3 types of restorations for endangered plants: reintroductions, enhancement or reinforcement, and introductions. Reintroductions occur at sites where a species was extirpated, introductions occur at previously unoccupied sites, and enhancement or reinforcement (augmentation) occurs at sites with extant populations. Although Ruth’s Golden Aster is well adapted to the harsh environment, the growing conditions pose challenges for managers seeking to restore or enhance populations of the plant. Initial restoration attempts with Ruth’s Golden Aster were not successful; only 1% of the plants survived past the first season (Cruzan and Beaty 1998). The reasons for the failure are not entirely clear, but the investigators recognized the potential for drought stress and soil disturbance to impact plants. In an attempt to address drought stress, managers augmented some restoration plots containing both seeds and transplanted rosettes with a moisture-retaining soil amendment. Over time, the amended medium swelled significantly, became dislodged from the planting crevices, and was washed way along with the propagules. Alternate treatments using sphagnum moss to retain moisture and netting to anchor plants and seeds were equally ineffective (Cruzan and Beaty 1998). Subsequent efforts by Wadl et al. (2014) to replant Ruth’s Golden Aster in suitable habitat employed bonded fiber matrix (BFM), a composite of polymers and wood fiber, used to stabilize soil and vegetation on disturbed sites. When wetted, BFM forms a thick slurry that adheres to itself and the surface to which it is applied. Resource managers assumed that the ability to glue Ruth’s Golden Aster plants in place after installation in narrow cracks would confer some resistance to scouring river flows Southeastern Naturalist 21 P.A. Wadl, A.M. Saxton, G. Call, and A.J. Dattilo 2018 Vol. 17, No. 1 and moisture loss on the drought- and flood-prone sites. Initial, small-scale efforts utilizing BFM in replanting proved successful; survival of Ruth’s Golden Asters transplanted into suitable habitat at a single location on the Ocoee River was 73% after 1 growing season. Despite the high survival rate, the methodology of the pilot study limited the applicability of the data to larger restoration efforts. All individuals transplanted to the study site were derived through clonal propagation of a single individual of unknown origin growing on the University of Tennessee, Knoxville, TN, campus. Without replication using multiple genotypes and sites, it is not possible to make meaningful inferences about the efficacy of the methods across the range of Ruth’s Golden Aster. In addition, larger-scale efforts involving more individual plants necessitate using plants derived from seed to avoid genetic swamping within populations. Given the variability in germination rates and seedling vigor exhibited by the species, it is possible the clones produced from stem cuttings would outperform plants grown from seed (Farmer 1977, White 1977). Our long-term objective is to develop methods to restore Ruth’s Golden Aster into unoccupied suitable habitat, as outlined in the recovery criteria for the species (USFWS 1992, 2012). Pavlik (1996) described short-term restoration success as the point at which a new population can carry on its basic life-history processes of establishment, reproduction, and dispersal, such that the probability of complete extinction by random or chaotic forces is low. Our objective in this study was to build on the work of Wadl et al. (2014) by testing the effect of BFM on abundance (establishment, fecundity, full life cycle can be completed) of seed-grown Ruth’s Golden Aster augmented along the Hiwassee and Ocoee Rivers. Materials and Methods Plant materials and transplanting We collected achenes (seeds) from open-pollinated Ruth’s Golden Aster plants at 5 Hiwassee River locations and 1 Ocoee River location in fall 2012 (Fig. 1) while plants were blooming and dispersing seeds. We pooled and placed seeds randomly collected from multiple individuals at each location directly into paper coin envelopes. Immediately following field collection, we transferred the seed into paper bags and dried them at ambient temperature (~23–24 °C) for 24 h (Farmer 1977). Filled seeds (embryo within fruit) are visibly larger in diameter compared to thinner, unfilled seeds (lacking an embryo); thus, we discarded the unfilled seeds and used only germinated, filled seeds according to Wadl et al. (2014). The general protocol was as follows. We removed the pappus, disinfected by immersing in 70% ethanol for 1 min, and briefly passed all seeds through a flame. Seeds were then placed into 50-mL conical tubes containing a 50% commercial bleach solution and shaken vigorously for 20 min. We decanted the bleach solution, rinsed the seeds 3 times with sterile water, placed individual seeds into a Bioworld Magenta® GA-7 plant culture box (Fisher Scientific, Waltham, MA) containing 50 mL of MS basal medium (Murashige and Skoog 1962), and incubated the boxes in the dark at 22–25 °C for up to 3 weeks or until germination. After 3 weeks, we calculated seed-germination percentages (seeds germinated/total number of seeds x 100). Southeastern Naturalist P.A. Wadl, A.M. Saxton, G. Call, and A.J. Dattilo 2018 Vol. 17, No. 1 22 We maintained the seedlings for 8 weeks in a growth room at 25 °C under a 16-h photoperiod provided by cool white fluorescent lamps. The lamps provided a photosynthetic photon flux of 125 μmol m-2 s-1 as measured by a Licor LI-250 light meter (LI-COR Inc., Lincoln, NE) held at the top of the culture vessels. After 8 weeks, we transferred plants to 72-cell flats containing ProMix soilless medium covered with a humidity dome for acclimatization in a greenhouse. We cut holes into the humidity dome after 24 h, and completely removed the dome after 3 days. The plants were acclimated to natural environmental conditions for 14 d at the University of Tennessee, Knoxville, TN, until outplanting. Planting and plant measurement We transplanted the ~3-month-old seedlings into suitable habitat at the seedcollection sites. At planting, we gently removed the ProMix soilless medium from the roots and backfilled the rock cracks with native soil from each planting location. After planting, we applied BFM to the treatment group and left the untreated control as bare soil (Fig. 2.). We used a metal spatula to apply the BFM as a thin layer of slurry (2–4 mm). All plantings were at least 1 m from the nearest native Ruth’s Golden Aster to avoid physical impacts to native plants and confusion about plant origin during the study period. We designed the study to test the effect of BFM on establishment and growth of Ruth’s Golden Aster. Competing vegetation at planting sites could have confounded results; thus, without disturbing Figure 1. Ruth’s Golden Aster restoration plantings at 5 sites on the Hiwassee River and 1 site on the Ocoee River in Polk County, TN. Hatched bars indicate the upstream and downstream extent of populations and delineate the entire range of the species. Southeastern Naturalist 23 P.A. Wadl, A.M. Saxton, G. Call, and A.J. Dattilo 2018 Vol. 17, No. 1 Figure 2. (A, C, E, G) Successful restoration of Ruth’s Golden Aster planted with soil, and (B, D, F, H) soil covered with bonded fiber matrix at (A, B) planting, (C, D) fall 2013, (E, F) spring 2014, and (G, H) fall 2014. Southeastern Naturalist P.A. Wadl, A.M. Saxton, G. Call, and A.J. Dattilo 2018 Vol. 17, No. 1 24 Ruth’s Golden Aster plants, we physically removed competing plants rooted in the same crack and within 50 cm of experimental plants. We marked planting sites so that we could reliably relocate plants and ensure accuracy of the data collected. To accomplish this, we used a battery-operated hammer drill to make 1 small hole in the rock at each of the 6 planting sites. The diameter of the hole accommodated a rebar stake that we used as a plot center. During sampling, we inserted rebar into the hole and fastened a meter tape to it. Using the meter tape, we recorded the distance and bearing to each plant (nearest cm) from the plot center. We used this information and other pertinent features in the landscape to generate a unique location for each plant. From this information, we recorded crack location for all plants and assigned plants in close proximity to each other and with similar habitat characteristics (boulder aspect, slope, crack orientation) into discrete habitat clusters. We recorded stem number, stem height, leaf number, flowering incidence, and number of flower heads per plant. In 2013, we made measurements at the time of planting (March), 1 month post-planting (April), and in October. In 2014 and 2015, we measured plants in April and October. To test fecundity, we collected mature seeds from an individual flower head from 2 plants in fall 2015, planted them, and assessed germination rate. Experimental design and statistical analyses For the population augmentation, we randomly assigned plants at each location to a treatment (BFM or no BFM) and the experimental design was a randomized complete block. We compared treatment and period differences for each response variable using a mixed model ANOVA (SAS v9.4, Cary, NC). Site and habitat-cluster effects were used as blocks, and we modeled repeated measures across periods with an autoregressive covariance. We employed Fisher’s LSD to separate least squares means at the 5% significance level. Normality and equal variance requirements were met for all variables, including survival (normal approximation to the binomial). We ran secondary analyses of variance with habitat cluster and specific crack location as fixed effects to assess environmental effects. We estimated Pearson correlations within and across periods to assist interpretation. Results Germination rate of filled seeds, number of acclimated seedlings, and total number of seedlings planted after 14 d of acclimatization was significant. Germination rate varied from 48% (O-01) to77% (H-02), number of acclimated seedlings varied from 51 (H-03) to 67 (H-01), and total number of seedlings planted varied from 18 (H-05) to 62 (H-01). There were no significant treatment differences for mean plant height (P = 0.2238), mean leaf number (P = 0.0742), mean flower number (P = 0.3611), stem number (P = 0.1544), final percent 3-year survival (P = 0.1006), or final percent 3-year survival if plant was alive at 1 month after planting out (P = 0.9505). Treatment was highly significant (P = 0.005) for percent survival at 1 month post-planting and moderately significant (P ≤ 0.02) for all dates monitored except the fall in year 3 (Fig. 3). Survival at 1 month was 32.2% across treatments, Southeastern Naturalist 25 P.A. Wadl, A.M. Saxton, G. Call, and A.J. Dattilo 2018 Vol. 17, No. 1 40.8% for plants mulched with BFM, and 23.6% for plants without BFM. By the end of the first season, survival was 24.4% for BFM plantings compared to 10.6% for the non-mulched plantings. For the remainder of the monitoring periods, survival remained relatively stable, and was 17.8% for the BFM planting and 10.5% for plantings without BFM after 3 seasons. Survival was variable across sites; plants flowered at H-2 and O-1 in the first season. Flowering occurred for all sites and treatments within the second season, except for non-mulched plants at H-1, and by the third season all sites and treatments with surviving individuals produced flowers (Table 1, Fig. 3). Germination in fecundity tests was 87.5% for a plant at H-1 (Soil + BFM treatment) and 0% for a plant at H-2 (soil treatment). The effect of crack location within site was not significant for survival at 1 month or final 3-year survival for any site. However, habitat cluster was significant for survival at 1 month at H-2 (P = 0.0225), H-3 (P = 0.0345), O-1 (P = 0.0119), and final survival at O-1 (P = 0.0081). Discussion Restoration of rare plant populations has become an increasingly common tool used for conservation of rare plant species, but there is uncertainty surrounding the long-term efficacy of the practice (Albrecht and Maschinski 2012; Dalrymple et al. 2012; Godefroid et al. 2011; Guerrant 2012, 2013). In a review of 249 restorations from 172 taxa from published literature and a survey of botanic gardens, universities, and conservation organizations, Godefroid et al. (2011), characterized Figure 3. Percent survival of Ruth’s Golden Aster restoration plantings at 5 sites on the Hiwassee River and 1 site on the Ocoee River in Polk County, TN. Survival was monitored at 6 different periods for each treatment group and periods labelled with an asterisk (*) differed between treatments at the 5% level. Southeastern Naturalist P.A. Wadl, A.M. Saxton, G. Call, and A.J. Dattilo 2018 Vol. 17, No. 1 26 Table 1. Total number of Ruth’s Golden Aster planted into suitable habitat on the Hiwassee and Ocoee Rivers (Polk County, TN) and percent survival and flowering percent of alive plants for 3 seasons. First year (2013) Second year (2014) Third year (2015) Number 1-month Fall Flowering Spring Fall Flowering Spring Fall Flowering Site Treatment of plants survival (%) survival (%) (%) survival (%) survival (%) (%) survival (%) survival (%) (%) H-1 Soil + BFM 31 2 (6.5) 2 (3.2) 0 (0.0) 2 (3.2) 2 (3.2) 2 (100.0) 2 (3.2) 2 (3.2) 2 (100.0A) Soil 31 2 (6.5) 2 (3.2) 0 (0.0) 2 (3.2) 2 (3.2) 0 (0.0) 2 (3.2) 2 (3.2) 2 (100.0) H-2 Soil + BFM 27 18 (66.7) 11 (40.7) 5 (45.4) 11 (40.7) 11 (40.7) 9 (81.8) 11 (40.7) 9 (33.3) 8 (88.9) Soil 27 11 (40.7) 6 (22.2) 1 (16.7) 5 (18.5) 5 (18.5) 5 (100.0) 5 (18.5) 5 (18.5) 4 (80.0A) H-3 Soil + BFM 18 10 (55.6) 8 (44.4) 0 (0.0) 8 (44.4) 8 (44.4) 5 (62.5) 8 (44.4) 6 (33.3) 5 (83.3) Soil 18 4 (22.2) 2 (11.1) 0 (0.0) 2 (11.1) 2 (11.1) 1 (50.0) 2 (11.1) 2 (11.1) 1 (50.0) H-4 Soil + BFM 18 3 (16.7) 1 (5.6) 0 (0.0) 1 (5.6) 1 (5.6) 1 (100.0) 1 (5.6) 1 (5.6) 1 (100.0) Soil 18 2 (11.1) 1 (5.6) 0 (0.0) 1 (5.6) 1 (5.6) 1 (100.0) 1 (5.6) 1 (5.6) 1 (100.0) H-5 Soil + BFM 9 7 (77.8) 4 (44.4) 0 (0.0) 3 (33.3) 3 (33.3) 2 (66.7) 3 (33.3) 2 (22.2) 2 (100.0) Soil 9 4 (44.4) 0 (0.0) 0 (0.0) 0 (0.0) 0 (0.0) 0 (0.0) 0 (0.0) 0 (0.0) 0 (0.0) O-1 Soil + BFM 18 6 (33.3) 3 (16.7) 0 (0.0) 2 (11.1) 2 (11.1) 2 (100.0) 2 (11.1) 2 (11.1) 1 (50.0) Soil 18 5 (27.8) 4 (22.2) 2 (50.0) 4 (22.2) 4 (22.2) 4 (100.0) 4 (22.2) 4 (22.2) 4 (100.0) AMature achenes (seeds) were harvested and germination was 87.5% for an individual from H-1 and 0% for an individual from H-2. Southeastern Naturalist 27 P.A. Wadl, A.M. Saxton, G. Call, and A.J. Dattilo 2018 Vol. 17, No. 1 mean rates of survival (52%), flowering (19%), and fruiting (16%) as quite low. Though categorizing the reported rates themselves as low maybe somewhat arbitrary, Godefroid et al. (2011) also observed a continuing decline in these metrics, indicating a continuing downward trajectory in the success of the restored plants. The combination of the survival, flowering, and fruiting rates, and the negative trend led the author to conclude that “most plant restorations will not be successful over the long-term.” Survival data for Ruth’s Golden Aster over the 3-year study period declined most significantly in the first month post-planting. After this initial decline, sites stabilized, and many locations lost few plants thereafter. Although stronger initial survival would be more encouraging, downward trends of survival in the early stages of restoration are not unusual and may not be indicative of long-term failure (Albrecht et al. 2011, Guerrant 2013). This finding does indicate the need or continuing monitoring past the 3-year study period to elucidate the ultimate success of the Ruth’s Golden Aster restoration (Godefroid et al. 2011). In contrast to the negative trend in flowering incidence observed in many restorations, Ruth’s Golden Aster generally exhibited increased flowering rates throughout the 3-year study; the mean flowering rate was 59.9% for BFM plantings and 47% for bare soil plantings. High flowering rates, combined with production of viable seed from restored Ruth’s Golden Aster, suggest the possibility of new recruitment into the study sites even during the relatively short study period. However, because of the small sample size, we cannot draw conclusions about recruitment without further testing. Although recruitment of new individuals derived from restored plants may be one of the most relevant metrics of success (Pavlik 1996), measuring this at the study sites is difficult because the nearby available habitat is largely occupied by naturally occurring Ruth’s Golden Aster of all age classes. More narrowly, in the context of direct conservation of Ruth’s Golden Aster, we contend that our study was successful for multiple reasons. First, we developed a methodology for restoration of Ruth’s Golden Aster in which the survival rate was higher than the 1% survival rate reported by Cruzan and Beaty (1998). Also, we found that survival was significantly higher at 1 month, fall year 1, spring/fall year 2, and spring year 3 for the plants mulched with BFM compared to the control. Survival rate was not significantly different for the treatment at the final fall year 3 measurement; however, the effect of BFM may still be biologically meaningful when considered in the context of future larger-scale restoration efforts for Ruth’s Golden Aster. The longevity of individual Ruth’s Golden Aster plants is unknown, and it is plausible that increased survival into year 2, when restored plants are flowering and presumably setting seed, may be sufficient justification for utilizing this low cost and easily implemented treatment. The decreasing positive effect of BFM is likely attributable to the ephemeral nature of the amendment, which degrades and disperses over time. BFM remained largely intact through the first year , but breakdown of BFM began to accelerate in year 2, and the substance was often absent or severely compromised by the end of the growing season. The positive effect of BFM on Ruth’s Golden Aster survival seems to persist as long the material remains affixed to the substrate around individual plants. Southeastern Naturalist P.A. Wadl, A.M. Saxton, G. Call, and A.J. Dattilo 2018 Vol. 17, No. 1 28 Our study focused on augmenting existing populations of Ruth’s Golden Aster, but the methodology could be readily applied to areas of formerly suitable habitat along the Hiwassee River that have been lost to succession since completion of the Apalachia Dam in 1943. Previous efforts to clear trees and shrubs around areas of former Ruth’s Golden Aster habitat were successful in producing the open conditions that the species requires, but the rock outcrops were not recolonized from adjacent occurrences of the species (USFS 2014). Lack of recruitment onto the newly cleared sites could be the result of poor seed dispersal or germination, or because the crevices in bedrock were not sufficiently cleared of sediments, which allowed the establishment of aggressive early successional annual plants species that excluded Ruth’s Golden Aster. The outcome of subsequent restorations could potentially be improved if managers remove accumulated sediments from boulders, install propagated Ruth’s Golden Aster plants on the sites using the methods outlined in this paper, and control competing vegetation until Ruth’s Golden Aster plants become established. In a larger context, these riparian rock-outcrop habitats, which are subjected to environmental extremes including frequent drought and prolonged flooding, are similar to other habitats where land managers try to restore populations of rare plant species (Homoya and Abrell 2005, USFWS 2008, Wells 2012). BFM may prove useful in other locations and species where plantings could benefit from increased resilience to scouring flood-flows and greater moisture retention during dry periods. The survival percentages, flowering incidence, and fecundity of planted Ruth’s Golden Aster that we observed during this study are encouraging. However, several factors (some manageable and some governed by chance) may have influenced the results. First, the crevices restored with Ruth’s Golden Aster should be carefully selected to better resemble cracks in which the species naturally grows. Cruzan and Beaty (1998) attempted to quantify the effect of crevice width on growth for established Ruth’s Golden Aster and found that plants in 10–15-mm-wide crevices produced the most stems, though many plants inhabited cracks 3–5 mm wide. All planting locations for our study were located within larger occupied Ruth’s Golden Aster sites and were never more than a few meters from native plants. The crevices into which we placed our plantings fell into this width range, but some may have been too shallow to effectively support plants. Cruzan and Beaty (1998) noted that it is difficult to discern crevice depth, but with experience, it is relatively easy to know when a particular crack is too shallow. For instance, cracks that are less than 1 cm deep and unlikely to naturally accumulate soil over time are probably not appropriate for restoration even if BFM is used to secure plants (and soil) in place. The year 1 overall survival rate of 73% observed during the initial Ruth’s Golden Aster pilot study suggests that using larger individuals may increase transplanting success (Wadl et al. 2014). Although plants used in the Ruth’s Golden Aster pilot study had nearly identical values for average stem height (13.4 cm vs. 13.5 cm) and average number of leaves per stem (6.4 vs. 6.3) when compared to the current study, plants in the pilot study had over twice as many initial stems per plant (4.2 vs. 1.9). This result indicates that plants used in the pilot study were comparable in Southeastern Naturalist 29 P.A. Wadl, A.M. Saxton, G. Call, and A.J. Dattilo 2018 Vol. 17, No. 1 overall height, but had about twice as much above ground biomass and presumably greater below ground biomasss. The clonal origin and method of propagation (in vitro multiplication vs. in vitro seedlings) may have influenced survival, and the greater initial starting size may have also played an important role. Weather can strongly influence the results of any outplanting in an uncontrolled setting. On the whole, 2013 was a very wet year in the watershed encompassing the reintroduction sites; most areas received 110–150% of normal rainfall (NOAA 2017). Several large rain-events occurred during the growing season, necessitating 4 releases of water from Apalachia Dam to minimize potential flooding to human-populated areas, which likely inundated some percentage of plants and may have lowered survival by physically dislodging individuals from planting sites, particularly H-1 and H-4. The largest of these releases, in both magnitude and duration, lasted for 11 days in July 2013. During this release, peak flows exceeded 4700 cubic feet/sec (cfs) for 5 days (TVA, Knowville, TN, unpubl. data). Releases approaching 5000 cfs are uncommon along this part of the Hiwassee River, which normally operates at a base flow of 25 cfs, because management of the system for electricity generation and flood prevention necessitates that nearly all water is diverted around the section of river where Ruth’s Golden Aster occurs. Though it is difficult to quantify, follow-up field surveys indicated that tributary inputs into this cutoff reach during the event were much higher than normal because the local area received significant rainfall with resultant unusually elevated river-flows. While we did not quantify any losses that may have occurred due to the extreme flood events, to overcome effects of such environmental variation, restoration efforts should maximize numbers of propagules for initial planting and incorporate multiple plantings. This approach would increase the likelihood that suitable conditions for establishment would follow 1 or more planting events and that fit genotypes would become established as a result of the restoration effort (Reichard et al. 2012). Ex situ conservation and cultivation of plants is possible through both stem cuttings and tissue culture, providing robust methods for restoration studies. It is feasible to grow Ruth’s Golden Aster seedlings in vitro and transplant them into the natural habitat (Wadl et al. 2011, 2014). Tracking demography within populations and augmented plantings as well as further work on seed-dispersal mechanisms, breeding success, and determining optimal time for restoration plantings are still needed and would be useful in understanding and protecting this endangered species. Planting season has been demonstrated to affect survival of restoration plantings (Albrecht and McCue 2009, Guerrant and Kaye 2007). Another factor to consider, and which cold be addressed in future studies, is the importance of mycorrhizal associations on the successful establishment of this species. This study provides a reliable protocol for the successful restoration for the Ruth’s Golden Aster. Based on population-structure analyses, Hatmaker (2016) recommended that the species should be managed as 2–4 populations along the Hiwassee River and 2–3 along the Ocoee River. Thus, combination of the results of Hatmaker (2016) and our current study provide a foundation for an effective approach to utilize knowldedge of genetic diversity in future restoration or ex situ conservation efforts. Southeastern Naturalist P.A. Wadl, A.M. Saxton, G. Call, and A.J. Dattilo 2018 Vol. 17, No. 1 30 Acknowledgments This work was supported by the US Department of Agriculture (USDA/MOA number 58- 6404-1-637), and the Tennessee Valley Authority (TVA). Mention of trade names or commercial products in this article is solely for the purpose of providing specific information and does not imply recommendation or endorsement by the TVA or the USDA. The USDA is an equalopportunity provider and employer. The findings and conclusions in this article are those of the authors and do not necessarily represent the views of the US Fish and Wildlife Service. We thank Mark Pistrang (USDA, Forest Service), David Nestor (TVA), and Dr. Denita Hadziabdic (University of Tennessee) for assisting with data collection. Seeds were collected under Tennessee Valley Authority Permit # TE117405-2 and US Fish and Wildlife Service Permit # TE134817-1. P.A. Wadl and A.J. Dattilo contributed equally to this project. Literature Cited Albrecht, M.A., and J. Maschinski. 2012. Influence of founder-population size, propagule stages, and life history on the survival of reintroduced plant populations. Pp. 171–188, In J. Maschinski and K.E. Haskins (Eds.). Plant Reintroduction in a Changing Climate: Promises and Perils. Island Press, Washington, DC. 432 pp. Albrecht M.A., and K.A. McCue. 2009. Changes in demographic processes over long time- scales reveal the challenge of restoring an endangered plant. Restoration Ecology 18:235–-234. Albrecht, M.A., E.O. Guerrant, and J. Maschinski. 2011. A long-term view of rare plant reintroduction. Biological Conservation 144: 2557–2558. Bowers, F.D. 1972. The existence of Heterotheca ruthii (Compositae). Castanea 37:130–132. Cruzan, M., and P. Beaty. 1998. Population biology or Ruth’s Golden Aster (Pityopsis ruthii): Final report, ID-96-05937-6-00. Tennessee Department of Environment and Conservation, Division of Natural Heritage, Nashville, TN. Dalrymple, S.A., E. Banks, G.B Stewart, and A.S. Pullin. 2012. A meta-analysis of threatened plant reintroductions from across the globe. Pp. 31–50, In J. Maschinski and K.E. Haskins (Eds.). Plant Reintroduction in a Changing Climate: Promises and Perils. Island Press, Washington, DC. 432 pp. Falk, D.A., C.I. Millar, and M. Olwell. 1996. Defining and measuring success. Pp. XIII– XXII, In D.A. Falk, C.I. Millar, and M. Olwell (Eds.). Restoring Diversity: Strategies for Reintroduction of Endangered Plants. Island Press, Washington, DC. 505 pp. Farmer, R.E. 1977. Seed propagation of Heterothecea ruthii. Casteanea 42:146–149. Godefroid, S., C. Piazza, G. Rossi, S. Buord, A.D. Stevens, R. Aguraiuja, C. Cowell, C.W. Weekley, G. Vogg, J.M. Iriondo, I. Johnson, B. Dixon, D. Gordon, S. Magnanon, B. Valentin, K. Bjureke, R. Koopman, M. Vicens, M. Virevaire, and T. Vanderborght. 2011. How successful are plant species reintroductions? Biological Conservation 144:672–682. Guerrant, E.O. 2012. Characterizing two2 decades of rare plant reintroductions. Pp. 9–-29, In J. Maschinski and K.E. Haskins (Eds.). Plant Reintroduction in a Changing Climate: Promises and Perils. Island Press, Washington, D.C. 432 pp. Guerrant, E.O. 2013. The value and propriety of reintroduction as a conservation tool for rare plants. Botany 91:v–x. Guerrant, E.O., and T.N. Kaye. 2007. Reintroduction of rare and endangered plants: Common factors, questions, and common approaches. Australian Journal of Botany 55:362–370. Hatmaker, E.A. 2016. Population genetics and genomics within the genus Pityopsis. M.Sc. Thesis, University of Tennessee, Knoxville, TN. Southeastern Naturalist 31 P.A. Wadl, A.M. Saxton, G. Call, and A.J. Dattilo 2018 Vol. 17, No. 1 Homoya, M.A., and D.B. Abrell. 2005. A natural occurence of the federally endangered Short’s Goldenrod (Solidago shortii T. & G.) (Asteraceae) in Indiana: Its discovery, habitat, and associated flora. Castanea 70:255–262. Moore, P.A., P.A. Wadl, J.A. Skinner, R.N. Trigiano, E.C. Bernard, W.E. Klingeman, and A.J. Dattilo. 2016. Current knowledge, threats, and future efforts to sustain populations of Pityopsis ruthii (Asteraceae), an endangered southern Appalachian species. Journal of the Torrey Botanical Society 143:117–134. Murashige, T., and F. Skoog. 1962. A revised medium for rapid growth and bioassays with tobacco tissue cultures. Physiologia Plantarum 15:473–497. NatureServe. 2015. NatureServe Explorer: An online encyclopedia of life [web application]. Version 7.1. NatureServe, Arlington, Virginia. Available online at http://explorer. Accessed 7December 2016. National Oceanic and Atmospheric Administration (NOAA). 2017. National Weather Service, Advanced Hydrologic Prediction Service. Available online at gov/precip/. Accessed 2 February 2017 Pavlik, B. 1996. Defining and measuring success. Pp. 127–156, In D.A. Falk, C.I. Millar, and M. Olwell (Eds.). Restoring Diversity: Strategies for Reintroduction of Endangered Plants. Island Press, Washington, DC. 505 pp. Reichard, S., H. Liu, and C. Husby. 2012. Is managed relocation of rare plants another pathway for biological invasions? Pp. 243–261, In J. Maschinski and K.E. Haskins (Eds.). Plant Reintroduction in a Changing Climate: Promises and Perils. Island Press, Washington, DC. 432 pp. Semple, J.C. 2006. Pityopsis. Pp.223–228, In Flora of North America Editorial Committee (Eds.). Volume 20. Flora of North America North of Mexico. 20+ Vols. New York, NY. Thomson, D.M., and M.W. Schwartz. 2006. Using population-count data to assess the effects of changing river flow on an endangered riparian plant. Conservation Biology 20:1132–1142. United States Fish and Wildlife Service (USFWS). 1992. Ruth’s Golden Aster recovery plan. Atlanta, GA. USFWS. 2008. Jesup’s Milk-vetch (Astragalus robbinsii var. jesupii), 5-year review: Summary and evaluation. US Fish and Wildlife Service, Concord, NH. USFWS. 2012. Ruth’s Golden Aster (Pityopsis ruthii), 5-year review: summary and evaluation. US Fish and Wildlife Serv., Cookeville, TN. US Forest Service (USFS). 2014. Fiscal year 2014 monitoring and evaluation annual report for the revised land and resource-management plan, Cherokee National Forest. Available online at Accessed 2 February 2017. Wadl, P.A., A.J. Dattilo, L.M. Vito, and R.N. Trigiano. 2011. Shoot organogenesis and plant regeneration in Pityopsis ruthii. Plant Cell, Tissue, and Organ Culture 106:513–516. Wadl, P.A., T.A. Rinehart, A.J. Dattilo, M. Pistrang, L.M. Vito, R. Milstead, and R.N. Trigiano. 2014. Propagation for the conservation of Pityopsis ruthii, an endangered species from the southeastern United States. HortScience 49:194–200. Wells, E.F. 2012. Reintroduction of federally endangered Harperella (Harperella nodosum Rose) in flood-prone, artificial, and natural habitats. Castanea 77:146–157. White, A.J. 1977. An autoecological study of the endangered species, Heterothecea ruthii (Small) Harms. M.Sc. Thesis, University of Tennessee, Knoxville, TN. Wofford, B.E., and D.K. Smith. 1980. Status report on Heterothecea ruthii (Small) Harms. US Department of the Interior, Fish and Wildlife Service, Region 4. Department of Botany, University of Tennessee, Knoxville, TN.