Seeds of D. drummondii DA462 and D. octopetala DA460 were purchased from the seed producer Jelitto (Schwarmstedt, Germany). The Nymphenburg Botanical Garden of Munich supplied seeds for D. drummondii BGM. Ecotypes Albe. (origin Clearwater County, Alberta, Canada, collected in 2000); and Alas. (origin Alaska, United States, collected in 2002) were found in and supplied by the KEW Millennium Seed Bank (Royal Botanic Gardens, Kew, London, United Kingdom). D. octopetala ecotype E548 was harvested in the Italian Alps (approximate GPS coordinates: 46°24′36.7″N 11°37′48.2″E) by Anna Heuberger.

Primers were designed based on the first draft genome of D. drummondii (Griesmann et al., 2018). They are as follows: GADPH forward 5′-CCCCAGTACGAATGCTCCCATGTTTG-3′, GADPH reverse 5′-TTAGCCAAAGGAGCAAGACAGTTGGTGG-3′; EF1-a forward 5′-TGGGTTTGAGGGTGACAACATGA-3′; EF1-a reverse 5′-GTACACATCCTGAAGTGGGAGACGGAGG-3′; 26S rRNA forward 5′-TACTGCAGGTCGGCAATCGG-3′, 26S rRNA reverse 5′-TCATCGCGCTTGGTTGAAAA-3′. ITS primers were designed based on Cheng et al. (2016): ITS forward 5′-CCTTATCAYTTAGAGGAAGGAG-3′, ITS reverse 5′-RGTTTCTTTTCCTCCGCTTA-3′.

Based on advice from seed producers and on results from Nichols (1934) who observed that without prior refrigeration, germination of several alpine species was considerably reduced, we assumed that seeds of Dryas species might require cold stratification prior to germination and therefore stored them at 4°C. Dryas seeds were surface sterilized by immersion in 30% H₂O₂ (10 min for D. octopetala; 15 min for D. drummondii) and washed three times with sterile H₂O. These experiments were performed with four biological replicates per species, with at least 100 seeds per replicate.

Sterilized seeds were transferred on 1% agar-water plates and incubated in the dark at 22°C for 12 and 8 days for D. octopetala and D. drummondii, respectively. Several sources of agar were tested such as Bacto^TM agar (Becton Dickinson and company) and agar Kalys HP 696 (Kalys SA, Bernin, France). The germination assays were set up in the dark because this reportedly increased germination rates Bliss (1958). After germination, seedlings of Dryas spp. were grown on plates, in a hydroponic system or in pots.

Growth on plates was performed on ¼ Hoagland’s pH 5.8 (using the protocol for N-free medium; Hoagland and Arnon, 1950) and adding 1 KNO₃ to a final concentration of 1 mM) with 0.4% of Gelrite (Duchefa, Haarlem, Netherlands), at 22°C, 55% of humidity with 16 h-light/8 h-dark cycles. After 1 week, plantlets were transferred either into pots or Weck jars (containing production substrate A210; Stender AG, Germany) or into a hydroponic system.

The hydroponic system consisted of a standard 1 mL pipette tip box in two parts: the bottom part contained 250 mL of growth medium (¼ Hoagland, 1 mM KNO₃, pH 5.8) and the tip holder with 24 holes in which the plantlets were inserted (Figure 2). To avoid seedlings or young plantlets falling into the medium compartment, the holes were covered with adhesive tape and plantlets were introduced through thin slits cut into the tape. The growth medium was changed twice per week. The hydroponic system was kept in a growth cabinet at 22°C, 55% humidity with 16 h-light/8 h-dark cycles for a maximum of 4 months.

Plants in pots were transferred to the greenhouse (day temperature 21–24°C, night temperature 18–21°C, with additional lighting from 6:00 to 10:00 h and from 15:00 to 22:00 h). The pots were filled either with sand:vermiculite (2:1) or with propagating substrate (A210 Stender AG, Germany). Note that temperature and light conditions were applied as available in our plant growth facilities and not experimentally optimized for Dryas.

For clonal propagation, young and soft shoots of Dryas spp. were cut after the third internode (2–5 cm) above the woody part of the shoot. These explants were directly transferred into moist production substrate A210 (Stender AG, Germany), then kept under plastic cover in the greenhouse. High humidity was maintained under the cover by spraying with water every 2 days for 2 weeks; thereafter, spraying was stopped and cuttings were kept in moist soil under the cover until new leaves had developed and the covers were removed. Three series of ca. 20 cuttings per species were cultivated in the greenhouse during different seasons.

We established a protocol for hairy root transformation in Dryas spp. by adapting Lotus protocols. The Agrobacterium rhizogenes strain AR1193 (Stougaard et al., 1987) was used because it had been shown to be very efficient for some plant species such as pea (Clemow et al., 2011) and because it was one of the strains available in our lab previously successfully tested for Lotus japonicus hairy root transformation.

Agrobacterium rhizogenes AR1193 bacteria carrying a Golden Gate LIIIβ F A-B (Binder et al., 2014) plasmid containing the mCherry gene under control of the Ubiquitin promoter (AtUbi10pro) as transformation marker (Pimprikar et al., 2016), were grown in liquid culture (LB medium with 50 μg mL^-1 each of rifampicin, carbenicillin and kanamycin) at 28°C overnight. Bacteria were collected via a centrifugation step (15 min at 4.369 ×g) and resuspended in water to obtain the wanted OD₆₀₀ (0.01; 0.1; 1; 7.2). Cut hypocotyls of 10–12 days old axenically grown Dryas spp. seedlings were dipped in the bacteria suspension and placed on ¼ Hoagland (1 mM KNO₃, pH 5.8), 0.4% Gelrite (Duchefa, Haarlem, The Netherlands) plates. The plates were kept for 4 days in the dark at 22°C, then under a 16 h-light/8 h-dark cycle with 55% humidity. To prevent overgrowth of bacteria and dehydration, the plants were transferred onto new plates every week. Four to six weeks after transformation, roots were screened using a Leica MZ16 FA stereomicroscope (Leica Microsystems GmbH, Wetzlar, Germany) using the N3 filter from Leica (BP 546/12;600/40).

The CsCl gradient DNA extraction method was performed according to Ribeiro et al. (1995). Six to ten grams of leaves (mix of young and old from the same plant) were ground by hand using pistil and mortar with 4 g of PolyclarAT in liquid nitrogen. For the other extraction methods, the two youngest leaves of a shoot with the apical meristem were used as starting material. After being shock frozen in liquid nitrogen, they were ground with a Retsch Mill MM400 (Fa. Retsch, Haan, Germany) two times at 30 Hz for 30 s in 2 mL Eppendorf tubes containing 2 mm diameter stainless steel beads each. The “classical CTAB” extraction method is described in Doyle and Doyle (1987), whereas the “PVP/NaCl” extraction method was developed by Khanuja et al. (1999) based on the classical CTAB method.

PCRs were performed on 1 μl of DNA (20–700 ng of DNA, usually ca. 100 ng) using GoTaq^® DNA polymerase (Promega, Germany), SYBR Green buffer and 0.2 μM of each primer. Amplifications were carried for 5 min at 95°C, followed by 35 cycles (30 s at 95°C, 30 s at 60°C, and 40 s at 72°C), and a final extension for 1 min at 72°C. Electrophoresis of a 4 μL of PCR reaction was performed on a 3% agarose gel for 100 min at 130 V. DNA was visualized with UVP UV solo touch from Analytik Jena© (Jena, Germany) after incubation of the gel for 10 min in an Ethidium bromide bath at 2 ng mL^-1.

Leaves, seedlings and root systems were shock frozen in liquid nitrogen. RNA of ground material (with the same procedure previously described for DNA extraction) was extracted using the Spectrum^TM Plant Total RNA Kit (Sigma-Aldrich, CA, United States) without adaptation in the protocol except for older and thicker leaves for extraction of which Polyclar AT was added. The RNA was treated with DNAse (Invitrogen^TM TURBO DNA-free Kit; Carlsbad, CA, United States) and tested for purity and integrity with a Bioanalyzer from Agilent (Agilent Technologies, Palo Alto, CA, United States). Twelve independently isolated RNA samples were analyzed.

Fungal contamination of seeds was often observed, whether they were collected in the field or obtained from a professional seed producer. In our study, white fungal hyphae growing out of the seeds led to seedling death at an early stage. Dryas seeds were quite sensitive to different surface sterilization procedures: any traces of ethanol would completely inhibit germination, while the thinness of the seed coat rendered the use of sulphuric acid for scarification risky. Furthermore, the contaminating fungi were quite resistant to NaOCl. However, after stratification of Dryas seeds at 4°C (Figure 1A), most efficient sterilization and highest germination rates were observed using hydrogen peroxide. Indeed approximately 100% of D. octopetala seeds and between 99 and 100% of D. drummondii seeds were free of contaminants after the procedure. Four replicates, with at least 100 seeds per replicate, were observed every 2 days. For D. drummondii, the maximal germination rate (85%) was obtained 8 days post-sterilization, whereas the maximum germination rate of D. octopetala, ca. 40%, was only reached 12 days post-sterilization (Figure 1C).

The time difference of 4 days to reach maximum germination observed between D. drummondii and Dryas octopetala was consistently observed at least within the samples tested: seeds of all D. drummondii seed sources examined germinated within 8 days and D. octopetala seeds from all sources available within 12 days. This phenotype was consistent not only for seeds produced in the greenhouse or botanical garden during the same month, but also for commercial seeds, the age of which was unknown. However, given that D. octopetala has a very wide distribution, it is possible that the two seed sources examined do not encompass the entire variability of the species.

We examined different growth conditions and systems including Gelrite and agar plates, hydroponic systems and classical pots. These distinct growth systems combine diverse advantages for research such as axenic culture, conditions for root system observations and for inoculation with the microsymbiont. Frankia strains able to nodulate D. drummondii have not yet been successfully cultured (Normand et al., 2017), necessitating infection with crushed nodules. As these nodules carry a rich fungal and bacterial microbiome on the surface, inoculation of Dryas with these nodules while maintaining a gnotobiotic system is challenging. On the other hand, plants grown in pots do not represent the most suitable system for root analyses. The process of cleaning the soil from the roots stresses the plant and furthermore, the harvesting of root systems entails the risk of breaking thin and fragile lateral roots and root hairs. To circumvent this drawback, plants can be grown in Petri dishes. For Dryas species this system was suited for early stages of development; for experiments exceeding 4–5 weeks, root and shoot growth required more space. Furthermore, shielding of plates never totally protected the roots from light, and a long exposure of roots to light tends to interfere with the analysis of root responses to any treatment. Exposure of roots to direct light modifies their transcriptome (Hemm et al., 2004) and often leads to stress responses, which can perturb the analyses and cause misleading effects. Hydroponic systems offer the possibility to observe the roots in a non-invasive way while also shielding them from light. They can be used with or without an inert substrate that mimics physical soil contact. The fact that Dryas species can grow in well-aerated soil but can also tolerate flood periods (West et al., 1993), suggested the use of hydroponics as a method of choice.

Once germinated and grown on 1% agar plates with classical plant media like B5 or MS (Murashige and Skoog, 1962; Gamborg et al., 1968; Duchefa, Haarlem, Netherlands) or Fåhræus medium (Fåhraeus, 1957) with 1 mM KNO₃, Dryas species seedlings turned reddish, likely due to μlanin production, a response typically interpreted as stress- or defense-related. This anthocyanin production was less pronounced when the seedlings were grown on 0.4% Gelrite with ¼ strength Hoagland medium containing 1 mM KNO₃ (Figure 1B). Moreover, after 2 weeks on plates, Dryas spp. plantlets grown on ¼ strength Hoagland medium showed darker green cotyledons and further developed root systems than on B5 medium. Thus, among the tested media for Dryas seedlings on gel-forming media, the best results were obtained with 0.4% Gelrite containing ¼ strength Hoagland solution.

After germination on plates, Dryas species. plantlets were transferred to a hydroponic system (Figure 2A) with ¼ Hoagland solution. In this system Dryas species plants grew and developed without obvious stress symptoms like accumulation of anthocyanins; they formed well-developed primary and lateral roots, and the speed of shoot development resembled that of pot-grown plants (Figure 2B). Altogether, Dryas species plants adapted very well to the hydroponic system tested. The absence of gel and soil substrates offers the opportunity to perform non-invasive observations of roots.

Dryas is a perennial plant genus. D. drummondii has been found to flower in its fifth year (Lawrence et al., 1967), indicating a long generation time that renders crossing experiments difficult. However, nodulation, plant growth and flowering processes in Dryas sp. seem to be extremely dependent on the environment and on light quality and intensity (Kohls et al., 1994). Therefore, we attempted to reduce the generation time under greenhouse conditions.

In our study, flower and seed production did not occur when plants were grown at a distance of 2 m from standard high-pressure mercury vapor lamps (providing 90 μmol m^-2 s^-1 at the plant level) used in initial trials (fluorescent lamps were not tried for flowering as seedlings from both species were growing significantly more slowly under them than under mercury vapor lamps for the first 5 weeks after germination). However, when the plants were placed at a distance of 2 m under high-pressure sodium vapor lamps (150 μmol m^-2 s^-1) for 16 h per day, flowering was induced (Figure 3B–D). High-pressure sodium lamps provide light with a richer emission in yellow-orange and a red/far red ratio shifted to the far red compared to standard high-pressure mercury vapor lamps and fluorescent lamps. The flowers produced seeds (Figure 1A) within less than a year after germination, whether the plants originated from cuttings or from sexual propagation. This is a far shorter generation time than the 5 years required for D. drummondii according to Lawrence et al. (1967). In the field, Dryas spp. flower primordia are formed during the summer, i.e., far in advance of flowering, which occurs in the next year shortly after snowmelt, with most individuals flowering within a month (Lawrence et al., 1967). This behavior suggests that the development of floral primordia and blooming depends on photoperiod or vernalization (or both). It is surprising that light from high-pressure sodium lamps, characterized by a lower red/far red ratio, leads to induction of flowering in an arctic/alpine species like D. octopetala, and of a species that has been described as extremely shade-sensitive (D. drummondii; Cooper, 1931). We observed that plants grew much better outdoors than in the glasshouse, but did not identify the limiting parameters. At any rate, changes of light period and temperature might further speed up the induction of flowering and shorten the generation time.

Seed set under greenhouse conditions occurred at ca. 75% of all D. drummondii flowers and at ca. 65% of all D. octopetala flowers. In their natural habitat, Dryas species combine autogamy and allogamy; seed set is improved when insects are available for pollination (Kevan, 1975; Roslin et al., 2013; Tiusanen et al., 2016, 2019). In the greenhouse, while some insects were usually present, seeds would be formed by flowers covered with paper bags, indicating that all seed batches used gave rise to plants that could perform self-fertilization.

Manual pollination was performed in the greenhouse and in a botanical garden to obtain hybrids. While the large and open flowers of D. octopetala (Figure 3D) made manual pollination possible, D. drummondii flowers were never completely open during full bloom (Figure 3C). Therefore, crossings of female D. octopetala with male D. drummondii were attempted. However, these attempts were not successful. Flowers of species contain multiple stamens, and often, after manual pollination it turned out that not all of them had been removed.

Altogether, sexual reproduction of Dryas spp. was feasible in a laboratory context when sufficient light intensity of a suitable spectrum was provided.

Given that Dryas spp. are partially allogamous and experimental studies require homogenous plant material, the use of Dryas as a model genus requires an easy protocol for vegetative propagation. In the wild, clonal growth of Dryas species enables individuals to persist and grow in extreme environments where sexual proliferation is often unsuccessful (Wookey et al., 1995), and where individual clones of D. octopetala commonly live for more than 100 years (Kihlman, 1890; Crawford, 1989). Thus, clonal propagation of Dryas species was expected to be easy. For clonal propagation by cuttings, three series of ca. 20 cuttings per species were grown in a greenhouse at different times of the year. Two to three cm of Dryas stems containing one node were transferred into moist soil (Figure 3E) in a small growth container with a transparent plastic lid for conservation of high humidity levels. Under these conditions, 65–95% of the Dryas cuttings developed roots within 3 weeks in the absence of hormonal treatments (Figure 3F). Once the shoots had successfully rooted, the plants were transferred into single pots and grown under standard greenhouse conditions. This easy protocol for vegetative propagation of Dryas species by cuttings in the glasshouse represents an important tool for performing experiments on a high number of plants that have the same genotype, and it obviates the requirement for seeds.

For a model plant, a protocol for genetic modification is important in order to analyze the expression of marker gene promoter-reporter gene fusions, or to perform reverse genetics. Hairy root transformation mediated by Agrobacterium rhizogenes is the most commonly used technique to introduce chimeric constructs into plant roots. The fact that this method does not transform the shoot is no hindrance to the study of root symbioses; A. rhizogenes-mediated hairy root transformation is routinely used not only in the model legumes Lotus corniculatus var. japonicus (Díaz et al., 2005) and Medicago truncatula (Boisson-Dernier et al., 2001) but also for actinorhizal plants like Datisca glomerata (Markmann et al., 2008), Casuarina glauca (Diouf et al., 1995) and (Imanishi et al., 2011) and non-FaFaCuRo plants like tomato (Ron et al., 2014).

In order to develop a hairy root transformation protocol, we inoculated axenically grown Dryas spp. seedlings with A. rhizogenes at different cell densities. 5 weeks after transformation, the composite plants on plate were evaluated. For D. drummondii, a transformation efficiency of 55–70% was obtained under all conditions tested, while D. octopetala plants died more frequently in response to infection with A. rhizogenes. The use of higher bacterial densities had a negative effect on plant survival, while lower bacterial densities reduced transformation efficiency. Here, the best compromise between low mortality and transformation rate for D. octopetala was observed when the A. rhizogenes suspension was adjusted to an OD₆₀₀ of 1. However, the transformation efficiency was still low with only 30% (Figure 4A). The experiment was repeated three times using an A. rhizogenes suspension adjusted to an OD₆₀₀ of 1 on ca. 70 seedlings per species. In all cases, the results were the same: 55–70% transformation for D. drummondii, maximally 30% transformation for D. octopetala.

Previous studies have shown that hairy roots induced by different bacterial strains can vary in morphology and production of secondary metabolites (Thwe et al., 2016); it was also shown that plant defense reactions, phytohormone signaling and secondary metabolism could be affected by high expression levels of the agrobacterial rolB gene (Bulgakov et al., 2018). Thus, the difference in the reactions of two closely related species to the same A. rhizogenes strain is interesting. At any rate, since only one A. rhizogenes strain was used in this study, the use of other strains might leave room for further optimization of hairy root transformation of D. octopetala.

Up to 7 weeks after transfer to pots or to the hydroponic system, transgenic roots showed healthy growth and expressed the transformation marker mCherry driven by the ubiquitin promoter (Figure 4B). However, roots growing in particle substrates such as sand:vermiculite, developed sections with increased lignification, hence more autofluorescence and opacity. This led to the quenching of the mCherry signal as highlighted by the red arrows in Figure 4B. In contrast, when plants were grown in the hydroponic system, lignification was less pronounced. Thus, the hydroponic system is well suitable for the observation of fluorescent proteins in Dryas spp. hairy roots.

The ability to clone Dryas spp. genes combined with the capacity to introduce chimeric constructs into root systems opens the possibility to study Dryas genetics in depth, allowing cross-species complementation, as well as transient expression, protein localization, and reverse genetics using CRISPR/Cas or RNAi methods.

For molecular biological studies, DNA and RNA have to be isolated with high purity, integrity and yield to be used for sequencing or reverse transcription, respectively. This was particularly challenging since the woody nature of Dryas species and the composition of the leaves adapted to harsh environmental conditions led to the presence of contaminants interfering with nucleic acid extraction protocols.

We tested different DNA extraction protocols on D. drummondii and D. octopetala, performing at least 30 extractions per method. DNA isolated from Dryas spp. with classical CTAB extraction protocols had an UV absorbance ratio at 260/280 of ca. 1.8, but the 260/230 ratio was always below 1.8, indicating polysaccharide contamination (Figure 5A,B). Several established DNA extraction methods were tested (Supplementary Table S1), but none of them led to a yield and purity sufficient for robust PCRs and de novo whole genome sequencing. However, a CTAB protocol adapted for recalcitrant plant material (Khanuja et al., 1999; “PVP/NaCl”) by addition of PVP, followed by a high salt lysis buffer and extraction with chloroform:isoamyl alcohol (24:1, v/v), resulted in good quality DNA suitable for PCRs with reproducible results (Figure 5B). Yet, the DNA yield and quality required for genome sequencing was so far only achieved using a modified Dellaporta et al. (1983) protocol followed by a CsCl gradient centrifugation as described by Ribeiro et al. (1995). The DNA extracted using this last method was used for de novo whole genome sequencing performed in collaboration with the Beijing Genomics Institute (BGI, China). The first version of the D. drummondii genome was used in a phylogenomic comparison study by Griesmann et al. (2018).

The Spectrum^TM Plant Total RNA Kit (Sigma-Aldrich) was used in order to extract RNA from different organs of Dryas spp. When Polyclar AT was added during the grinding step for recalcitrant samples (e.g., mature leaves and lignified roots), this method resulted in RNA of suitable integrity and purity for the performance of reverse transcription-quantitative PCR as indicated by the RNA integrity number (Figure 5C). RNA extracted from roots, leaves and seedlings following this method was used by the BGI in order to assist gene prediction for the D. drummondii genome (Griesmann et al., 2018). The transcripts were mapped to the protein-coding gene models, identified using the MAKER-P pipeline (version 2.31; Campbell et al., 2014), in order to obtain gene characteristics (size and number of exons/introns per gene, distribution of genes, features of splicing sites, etc.).

All method comparisons are summarized in Supplementary Table S1.

We tested the suitability of the DNA preparations resulting from different protocols as templates for PCR. Based on the published D. drummondii genome (Griesmann et al., 2018). Primers were designed based on the first version of the D. drummondii genome. The targets were regions in the internal transcribed spacer (ITS) of nuclear ribosomal DNA, 26S ribosomal RNA (26S rRNA), glyceraldehyde 3-phosphate dehydrogenase (GAPDH) and the elongation factor 1-alpha (EF1-a). Using D. octopetala gDNA as template, fragments were amplified and sequenced as well. The amplification confirms that the DNA preparations were of sufficient quality, while high sequence conservation in these regions highlights the similarity between D. drummondii and D. octopetala. Indeed, the size of amplicons of both species were similar and their sequences presented few single nucleotide polymorphisms (Figure 6).

With the basic but indispensable procedures and protocols for cultivation, vegetative and sexual propagation, hairy root transformation of and nucleic acid isolation from Dryas spp. described in this study, Dryas emerges as a new model genus to study important traits associated with survival in arctic and alpine conditions, including the formation of root symbioses with bacteria and ectomycorrhizal fungi.