Abstract
Coastalsand frostweed (Crocanthemum arenicola (Chapm.) Barnhart [Cistaceae]) is a back dune plant of the north central Gulf of Mexico endemic to coastal Florida, Alabama, and Mississippi. We initiated seed and cutting propagation experiments to test the effects of scarification, photoperiod, and temperature on germination of C. arenicola. In addition, we examined the effects of cutting maturity (vegetative or reproductive stems), auxin (IBA) concentration, and time of rooting on the percentage of cuttings with roots and the quality of rooted cuttings of C. arenicola. We conducted 3 single-factor experiments in which seeds were subjected to scarification treatments, photoperiod treatments, or a gradient of temperature treatments, and germination (radicle emergence) was monitored over time (2–4 wk). A two-factor greenhouse cutting experiment was conducted comparing cutting maturity and auxin concentration on root number and root length of C. arenicola 3 and 7 wk after sticking. We assessed cuttings for root class at 7 wk. Scarification by sandpaper abrasion (50–200 s) increased germination compared to a non-scarified control (≥90% compared to 11% germination). Photoperiod had no effect on germination, with similar germination in the light and dark. Higher germination occurred under cooler temperatures than warmer temperatures. Rooting at 3 wk (51%) was lower compared to rooting at 7 wk (76%). More roots were present on vegetative cuttings compared to reproductive cuttings (4.2 compared to 2.7 roots per cutting). Root length and root class did not differ with the application of auxin.
- coastal restoration
- germination
- physical dormancy
- seed scarification
- photoperiod
- cutting propagation
- Cistaceae
CONVERSIONS
(°C × 1.8) + 32 = °F
1 g = 0.04 oz
1 ml = 0.03 oz
1 mm = 0.04 in
1 cm = 0.40 in
Crocanthemum spp. (Cistaceae) are perennial herbs to subshrubs native to dry, sandy soils of the Americas (eFloras 2008; Guzmán and Vargas 2009; Sorrie 2011). Species within this genus support many insect species (Bottimer 1969; White and others 2016; Tschinkel and Domínguez 2017) through complex plant–insect interactions (Hillier and others 2018). Coastalsand frostweed (Crocanthemum arenicola (Chapm.) Barnhart [Cistaceae]) is an endemic with a range limited to 12 coastal counties of the Florida panhandle, Mississippi, and Alabama (USDA NRCS 2020; Wunderlin and others 2020). It occurs in ecosystems with exceptionally dry, infertile soils, such as coastal back dunes (Wunderlin and Hansen 2011). This species is an important dune colonizer post-disturbance. After Hurricane Katrina, C. arenicola occurred at the highest frequency of all plant species present on back dunes of Horn Island, Mississippi (Lucas and Carter 2013). It is a strong candidate for coastal restoration and coastal vegetation projects within its native range. Likewise, it has ornamental value with prolific flower production and a prostrate growth from (Figure 1). It also has potential for use as a ground cover in low-input landscapes, pollinator gardens, and in coastal landscapes. Crocanthemum arenicola has 2 types of flowers, showy insect-pollinated (chasmagamous) yellow flowers and non-showy selfing (cleistogamous) flowers (eFloras 2008). As such, the seeds for this plant can be the result of either self-pollination or outcrossing. Crocanthemum species have been moved from Helianthemum (Cistaceae) (Sorrie 2011) but share ancestry with members of Helianthemum (Guzman and Vargas 2009) that are commonly used in low-input landscaping.
Propagation information for C. arenicola is lacking; however, other Crocanthemum and the closely related Helianthemum species are known to have physical dormancy that may be alleviated by scarification (Thanos and others 1992; Pérez-García and González-Benito 2006; Luna and others 2007; Yeşilyurt and others 2017). Seeds of Crocanthemum scoparium (Nuttall) Millspaugh, a native of California found in coastal habitats, had physical dormancy alleviated by exposure to 120 °C dry heat for 5 min (Keeley and others 1985). Physical dormancy was alleviated for Helianthemum apennium (L.) Miller (Cistaceae) when exposed to 80 °C dry heat for 10 min, whereas dormancy in Helianthemun hirtum (L.) Miller and Helianthemum rotundifolium Dunal was not alleviated by 10 min of 80, 100, or 120 °C dry heat treatments (Luna and others 2007). Hot water baths (95–100 °C) followed by cooling in water at room temperature and chemical scarification (95–97% concentrated sulfuric acid for 15–30 min) alleviated dormancy upon incubation at 25/15 °C with a 16-h photoperiod for Helianthemum polygonoides Peinado, Martínez-Parras, Alcaraz & Espuelas. Physical dormancy of Helianthemum squamatum (L.) Pers. was alleviated only by mechanical scarification with sandpaper (Pérez-García and others 1995).
Previous studies on germination across Cistaceae described optimal germination temperatures at 15 to 20 °C in dark conditions (Thanos and others 1992). Seeds of C. scoparium germinated at 23 °C (Keeley and others 1985). Germination requirements for the sister group of Helianthemum are better studied. Higher germination (33–46%) for H. squamatum was noted at temperatures between 20 and 30 °C when compared to 5 to 15 °C, which had 20% germination (Escudaro and others 1997). High (>70%), medium (~50%), and low (<10%) germination percentages were observed at 10 to 20, 25, and 30 to 35 °C, respectively, for Helianthemum vesicarium Boiss. (Gutterman and Agami 1987), while Helianthemum ventosum Boiss. had 60 to 70% germination between 15 and 35 °C and approximately 20% germination at 10 °C.
Information regarding the asexual propagation of Crocanthemum species is lacking. Stem propagation within a micropropagation system has been achieved with and without auxin for species of Helianthemum (Morte and Honrubia 1992; López and others 2006; Hamza and others 2013), however, indicating propagation with stem cuttings may be possible for species in Crocanthemum. In the following experiments, we tested the effects of scarification, photoperiod, and temperature on germination and the effects of auxin and cutting maturity on rooting performance of stem cuttings of C. arenicola.
METHODS
Stock Plants and Seed Collection
We collected seeds and cuttings from stock plants throughout fall and winter 2017. Stock plants were derived from spring-collected softwood stem cuttings from wild plants growing in back dunes on Santa Rosa Island, Florida (30.297933, −87.438964). Plants were then grown in containers filled with a 2:1 mix of 0.6 cm screened pine bark:MetroMix 830 (Sun Gro Horticulture, Agawam, Massachusetts) and fertilized with 2.5 g of Osmocote 18-6-12 (Scotts-Sierra Horticultural Products, Marysville, Ohio). Plants were hand-watered and grown under natural day length in a climate-controlled polyhouse with no shade at the West Florida Research and Education Center at the University of Florida in Milton, Florida (Figure 2).
Germination Experiments
We collected fruits containing discrete seed units (Figure 3) from stock plants as fruits matured (turned brown) and either fell off by abscission or fell off when the plant was lightly shaken. Fruits were air-dried under laboratory conditions (approximately 25 °C and 70% RH) for at least 2 wk, placed in an airtight jar, and stored in the dark until seed extraction prior to experiment initiation. Seeds were extracted from fruits by lightly placing pressure on seed units until seeds disassembled (Figure 3). Preliminary work (Campbell-Martínez and others 2018) demonstrated the seeds have a testa that prevents water uptake (physical dormancy). We considered seeds to be imbibed when they became swollen and the testa became translucent, germinated when the radicle protruded through the testa, and diseased when hyphal growth was present on the seed (Figure 4).
Scarification Experiment
We conducted a single-factor experiment (scarification) with 6 levels (0 s of scarification without bleach sterilization and 0, 50, 100, 150, and 200 s of scarification). Scarification was accomplished by sandpaper abrasion in an electric seed scarifier lined with sandpaper (Gator Grit 80-grit aluminum oxide). Seeds for this experiment were graded, and seeds that appeared empty, deformed, or discolored were discarded. All but the control group seeds were surface sterilized with a 1-min 70% isopropyl alcohol bath, followed by a 10-min bath of 0.825% dilute bleach solution (10% bleach [8.25% sodium hypochlorite] and 90% water), and finally triple rinsed with distilled water. Seeds that were not surface sterilized (control) were used to determine if the sterilization process alone would break physical dormancy. We used 4 Petri dishes per treatment, each with 25 seeds, which were kept in the dark (double wrapped in aluminum foil) and maintained at room temperature (approximately 25 °C) in a laboratory. We recorded radicle emergence after 14 d.
Photoperiod Experiment
We conducted a single-factor experiment (photoperiod) with 2 levels of light exposure (0 or 12 h of light). Seeds for this experiment were not graded or sterilized prior to scarification for 50 s in an electric seed scarifier lined with 80-grit aluminum oxide sandpaper. Seeds were exposed to a 12-h photoperiod of cool-white fluorescent light or were kept in the dark. Five replicate Petri dishes containing 25 seeds were used for each treatment. An additional 2 Petri dishes with non-scarified seeds were exposed to the dark to confirm that seeds remained physically dormant throughout the experiment. We placed Petri dishes in growth chambers at 25 °C and evaluated for germination (1 mm radicle emergence) and disease (presence of contamination) tri-weekly for 28 d. Petri dishes wrapped in aluminum foil were unwrapped for no more than 5 min and exposed to ambient laboratory lighting.
Temperature Experiment
We conducted a single-factor experiment (temperature) with 8 treatment levels (18, 20, 22, 24, 26, 28, 30, and 32 °C constant temperatures). Temperatures were achieved by placing germination boxes containing seeds on top of a thermogradient table (Model 5010, Seed Processing Holland, Enkhuizen, Holland) exposed to the dark. Seeds were scarified for 50 s using an electric seed scarifier lined with 80-grit aluminum oxide sandpaper, then placed in germination boxes with germination paper placed on top of blotter paper saturated with 10 ml of 0.1% solution containing Plant Preservative Mixture (Apollo Scientific Limited, UK) and deionized distilled water. We added solution as needed throughout the experiment. We used 4 germination boxes, each with 50 seeds, for each temperature treatment, and germination boxes were the experimental unit. We recorded germination (radicle emergence) tri-weekly for 28 d.
Cutting Experiment
The experiment utilized a randomized complete block design with a 2 (wk after sticking) × 2 (cutting maturity) × 4 (auxin concentration) full factorial arrangement of treatments. Cuttings were harvested for evaluation 3 or 7 wk after sticking. Cutting maturity was either vegetative or reproductive softwood stem cuttings that were graded on length (all at least 10 cm) and assigned to 1 of 4 blocks. Cuttings were taken early morning on 16 February 2018 from stock plants. We quick dipped (1 s) the distal 1 cm in either 0, 1000, 2500, or 5000 ppm auxin solution consisting of distilled water and dissolved indole-3-butyric acid (K-IBA) (Hortus USA, New York, EPA Reg # 63310-22) and then inserted them into 72-cell propagation flats containing Sun Gro MetroMix 830 growing medium. Plants were then placed under intermittent mist, which operated from 6 am to 5 pm with 3 s of mist every 20 min until experiment termination.
We evaluated rooting by means of a destructive harvest 3 and 7 wk after sticking. The number of cuttings with roots was recorded at wk 3 and 7. For each rooted cutting, we measured the length of the longest root (mm) and counted the number of roots at wk 3. Root class was assessed at wk 7. Root class was defined using a scale based on a range of the proportion of the rootball media held cohesively after removal from pots (1 = 0–5%; 2 = 6–25%; 3 = 26–75%; 4 = 76–95%; 5 = 96–100%). Six, 6-cell packs were the experimental units for evaluating the proportion of cuttings with roots, and the experimental units for all other variables were alive, rooted cuttings within each block.
Statistical Analysis
We analyzed the scarification experiment and the cutting experiment using generalized linear mixed models. We accomplished this using PROC GLIMMIX in SAS 9.4 (SAS Institute 2013). A Kenward-Rogers approximation was used for computing the denominator degrees of freedom for the fixed-effects tests. Block and experiment runs were treated as random effects. Temperature and photoperiod experiments were analyzed using survival analysis techniques described in Campbell-Martínez and others (2019), as germination was tracked over time rather than at a single data collection date, and diseased seeds were removed (right-censored) throughout experiments. In short, we used Cox regression models and Kaplan-Meier estimations with seed germination as the event. Cox regression analyses were conducted using SAS version 9.4; Kaplan-Meier curves of survivor functions were generated for each covariate using R version 3.4.3. The Kaplan-Meier curves were inverted when graphed to show probability of germination (radicle emergence) over time. For all analyses, P ≤0.05, a probability value of less than or equal to 0.05, was considered significant.
RESULTS
Analysis of data from the scarification experiment demonstrated that the only significant main effect was scarification (F5,18 = 357.93, P <0.0001). Germination percentages for scarified seeds (90–96%) were significantly higher than for non-scarified seeds (9–11%) regardless of time of abrasion in the seed scarifier (Figure 5). Additionally, sterilization did not scarify seeds, with 9% germination for surface-sterilized seeds and 11% germination for seeds not surface sterilized (Figure 5).
Analysis of data from the photoperiod experiment demonstrated that photoperiod had no effect (P >0.05) on germination probability across time (Table 1). Germination began within a week and continued through to the end of the experiment (Figure 6). At the end of the experiment, 63–64% of non-diseased seed had germinated for seeds in the light and in the dark. Non-scarified seeds remained dormant with minimal germination (14%) throughout the experiment.
Analysis of data from the temperature experiment demonstrated temperature influenced (P ≤0.05) germination with hazard ratios indicating similar probabilities of germination at 18, 20, and 22 °C (Table 2; Figure 7). Germination was observed within a week and it continued through to the end of the experiment, and we observed a general decrease in germination as temperatures increased with almost no germination at 30 and 32 °C (Figure 7). Highest germination (approximately 60%) occurred at 18, 20, and 22 °C compared to higher temperatures (<50% germination) across time.
Analysis of data from the cutting experiment demonstrated that cutting maturity, auxin concentration, and interactions among main effects did not affect (P >0.05) the percentage of cuttings with roots (Table 3). Timing of evaluation, however, did affect (P ≤0.05) the proportion of cuttings with roots: 51 ± 4% of cuttings had roots 3 wk after sticking and 76 ± 3% of cuttings had roots 7 wk after sticking. Root number per cutting did not differ with auxin application when evaluated 3 wk after sticking (Table 3), but there were more roots on vegetative cuttings compared to reproductive cuttings (4.2 ± 0.49 compared to 2.7 ± 0.21 roots per cutting). Root length (20.4 ± 1.7 mm) was not affected by cutting maturity, auxin, or their interaction (Table 3). Root number and root length could not be measured 7 wk after sticking, hence root class was evaluated at wk 7. Root class did not differ (P >0.05) among auxin treatments (root class = 3.4 ± 0.13), indicating all rooted cuttings held a similar proportion of the propagation media when removed from the propagation cells at 7 wk (Table 3). A root class value of 3 indicates 26–75% of the propagation media was held by the cuttings.
DISCUSSION
Like seeds of other species of Cistaceae, seeds of C. arenicola have physical dormancy due to a water-impermeable seedcoat (Thanos and others 1992). However, a few seeds of C. arenicola germinate readily without pretreatment (i.e., are not dormant). Similarly, a portion of seeds of C. scoparium have also been reported to germinate without pretreatment (Keeley and others 1985). Like the closely related Helianthemum, seed dormancy of C. arenicola can be alleviated by abrasion with sandpaper (Pérez-García and others 1995; Pérez-García and González-Benito 2006). Likewise, Helianthemum germinate readily with exposure to light (Escudero and others 1997) or in the dark (Yeşilyurt and others 2017), as was the case for C. arenicola. There was not a scarifying effect from bleach sterilization on seeds, indicating our sterilization process did not break dormancy.
Crocanthemum arenicola prefers cooler (18–22 °C) rather than warmer temperatures (24–32 °C) for seed germination. Similar responses have been documented across multiple species of Helianthemum, with high germination percentages (59–68%) reported at temperatures between 15 and 20 °C (Gutterman and Agami 1987; Thanos and others 1992; Martin and others 1995; Pérez-García and others 1995; Robles and Castro 2002; Pérez-García and González-Benito 2006). However, tolerance of warmer temperatures (25–35 °C) has been reported for Helianthemum species (Gutterman and Agami 1987; Escudero and others 1997), which was not evident for C. arenicola. Once imbibition occurs, germination quickly ensues for C. arenicola and many species of Helianthemum (Escudero and others 1997; Pérez-García and González-Benito 2008).
Stem cuttings of C. arenicola produced roots with or without the application of auxin. A similar response to auxin was also noted for micropropagated stem cuttings of Helianthemum (Morte and Honrubia 1992; López and others 2006; Hamza and others 2013). While the proportion of cuttings with roots was generally unaffected by cutting maturity, there were more roots per cutting for vegetative cuttings than for reproductive cuttings.
CONCLUSION
Seeds of C. arenicola are orthodox (tolerate dry storage) and should be collected from plants when fruits turn brown. Once collected, scarified seeds should be sown during periods of cool weather in light or in dark. To germinate seeds of C. arenicola, we recommend at least 50 s of abrasion in an electric seed scarifier lined with 80-grit sandpaper prior to sowing, with or without light exposure.
Vegetative propagation of C. arenicola with softwood stem cuttings may also be accomplished without the use of supplemental auxins. Auxin did not appreciably improve rooting performance in any measured response. The proportion of cuttings with roots increased when cuttings remained under mist for 7 wk compared to 3 wk. When necessary, reproductive stems can be propagated with only minor reductions in rooting performance.
ACKNOWLEDGMENTS
This work was supported, in part, by the USDA National Institute of Food and Agriculture McIntire Stennis project FLA-JAY-005222, McIntire Stennis project FLA-WFC-005653, US Fish and Wildlife Service, and the Florida Wildflower Foundation. A special thanks to Ashley Moore, Gina Mangold, and Logan Patterson for assistance with experiment management and data collection. Assistance with final review of the manuscript provided by Gina Mangold.
Footnotes
Photos by Gabriel Campbell-Martínez