In vitro plant regeneration and Agrobacterium tumefaciens–mediated transformation of Datura stramonium (Solanaceae)

Premise of the Study Datura stramonium is a pharmacologically and evolutionarily important plant species in the family Solanaceae. Stable transformation methodology of this species would be advantageous for future genetic studies. Methods In vitro plant regeneration and Agrobacterium tumefaciens–mediated transformation techniques were developed for D. stramonium based on methods reported for tomato. A binary vector containing pAtUBQ10::erGFP was used for transformation. Results We recovered primary transformants harboring the green fluorescent protein (GFP) transgene that resulted in expression of fluorescence in all tissues analyzed. Transformants were allowed to self‐pollinate, and two of five progeny contained the GFP transgene and displayed fluorescence identical to the primary transformants. Discussion We have demonstrated the first stable transformation in the genus Datura. This is a key first step to study the genetic basis of traits in this evolutionarily interesting species.


Transformation and co-cultivation
Agrobacterium tumefaciens GV3101 containing a pAtUBQ10::erGFP binary vector ( Fig. 1) was kindly provided by Dr. Jaimie Van Norman (Van Norman et al., 2014) and grown in 25 mL of liquid Luria-Bertani medium supplemented with gentamicin and spectinomycin to an optical density measured at a wavelength of 600 nm (OD 600 ) of 0.6 (approximately 48 h). The culture was pelleted by centrifugation at 4000 rpm for 10 min and resuspended in 25 mL of liquid 2% MS0 medium (4.3 g/L MS salts, 100 mg/L myo-inositol, 0.4 mg/L thiamine, 0.5 mg/L pyridoxine, 0.5 mg/L nicotinic acid, 2 mg/L glycine, 20 g/L sucrose [pH 5.6]).
Cotyledon segments were incubated in the Agrobacterium suspension for approximately 5 min, then placed adaxial side down on a new plate of KCMS medium for co-cultivation in the dark for 48 h.

Shoot regeneration
After co-cultivation, 70 cotyledon segments were moved to 2ZBT medium containing 4.3 g/L MS salts with Nitsch Vitamins (Caisson Labs, Smithfield, Utah, USA), 100 mg/L myo-inositol, 20 g/L sucrose, 2 mg/L zeatin, 300 mg/L timentin, 9 mg/L phosphinothricin, 5.2 g/L Agargel (pH 6.0). Filter-sterilized zeatin, timentin, and phosphinothricin were added after autoclaving once the medium reached ~55°C. The cotyledon segments were incubated under the same light conditions used for seed germination. Over the next two weeks, the cotyledon segments were transferred to new 2ZBT plates three times.
During this period, 23 cotyledon segments became necrotic and were discarded. The 47 surviving segments displayed callus growth and were transferred to 16-ounce polypropylene deli containers (Fabri-Kal, Kalamazoo, Michigan, USA) containing 1ZBT medium (identical to 2ZBT except for the addition of 1 mg/L of zeatin instead of 2 mg/L). Six weeks after co-cultivation, the calli began to produce leaves. Over the next several weeks, the calli produced approximately 24 shoots.

Rooting and greenhouse transfer
The survival of these plants for six weeks on antibiotic-containing media indicated that they were antibiotic resistant and therefore transformed. To speed rooting, shoots that were 1-2-cm tall were excised and placed on non-selective rooting medium (4.3 g/L MS salts with Nitsch vitamins, 30 g/L sucrose, 1 mg/L indole-3-aceticacid [IAA], 8 g/L Difco Bacto agar [pH 6.0]). After one week, bacterial contamination was evident in the containers and we therefore selected nine robust shoots for direct rooting in soil in order to avoid the loss of explants to the bacterial contamination. These shoots were excised, the cut stems dipped in Rootone (Bayer CropScience, Research Triangle Park, North Carolina, USA), and placed directly in soil under a plastic dome to maintain humidity until root growth was evident.
For four weeks, the nine primary transformants did not elongate or display vigorous leaf growth as they developed roots. Three primary transformants survived this acclimatization period, while the other six were lost likely due to stressful conditions in the growth room. Approximately 4.5 months after co-cultivation with Agrobacterium, the three remaining primary transformants were transferred into a greenhouse, where one further plant was lost to pest damage. The surviving primary transformants (T 0 -1 and T 0 -2) had vigorous growth and produced typical-sized leaves, fruits, and seeds after their transfer to the greenhouse. These two surviving T 0 plants were selected for further phenotyping to confirm GFP fluorescence and the presence of the transgene.

T 1 plants
Before the transfer to the greenhouse, the 4-5-cm-tall T 0 plants began to flower despite, at this stage, having very few leaves (usually fewer than three). Flowers from two plants self-pollinated and set fruit. The T 1 seeds collected from fruits before the transfer to the greenhouse were small compared to wild-type (~1 mm vs. 4 mm for wild-type), and, upon dissection, most were determined to be empty seed coats. Five viable T 1 seeds were produced by the primary transformants and pooled. The seed coats of these were removed, and the seeds were surface sterilized, germinated on 1/2 MS0 medium, and transferred to soil. These T 1 plants grew normally compared to wild-type plants, and displayed typical flowering time and seed set.

DNA extraction and PCR conditions
Young leaf tissue (~3 cm 2 ) from T 0 , T 1 , and wild-type plants was harvested in 2-mL collection tubes and snap frozen in liquid nitrogen. Genomic DNA was extracted according to King et al. (2014). See https://doi.org/10.17504/protocols.io.sgpebvn for a step-bystep protocol. Primers to amplify 982 bp of the the GFP coding sequence were designed and checked for dimerization and deleterious secondary structure using the IDT OligoAnalyzer 3.1 (Integrated DNA Technologies, Skokie, Illinois, USA). The primer sequences were forward 5′-CTGTCAGTGGAGAGGGTGAAGG-3′ and reverse 5′-TAAAGTTGCTCGAGGTACCCGG-3′. Approximately 50 ng of genomic DNA from each plant was used to amplify a region of the GFP coding sequence using EconoTaq Plus Green 2x Master Mix (Lucigen, Middleton, Wisconsin, USA). Cycling conditions were an initial denaturation at 94°C for 2 min; followed by 25 cycles of 94°C for 20 s, 56°C for 20 s, and 72°C for 60 s; and a final extension step at 72°C for 5 min. PCR amplification of approximately 650 bp of ACTIN was used as a positive control. Primers for ACTIN were forward 5′-GAT-GGATCCTCCAATCCAGACACTGTA-3′ and reverse 5′-GTATTG-TGTTGGACTCTGGTGATGGTGT-3′. Cycling conditions consisted of an initial denaturation at 95°C for 3 min; followed by 20 cycles of 95°C for 30 s, 55°C for 30 s, and 72°C for 30 s; and a final extension step at 72°C for 10 min. These amplicons were visualized on a 2% agarose gel stained with GelRed (Biotium, Fremont, California, USA).

GFP visualization
Vegetative and reproductive organs of wild-type plants, primary transformants (T 0 ), and T 1 progeny were imaged on a Leica M165 FC stereoscope (Leica Microsystems CMS GmbH, Wetzlar, Germany) using white light or, for GFP, using an 40-nm-bandwidth excitation filter centered at 470 nm with a 50-nm-bandwidth barrier filter centered at 525 nm to block chlorophyll fluorescence. All white light images were taken with an exposure time of 75-100 milliseconds, and all images for GFP fluorescence were taken with a 3-s exposure time using a Leica D450 C digital microscope camera (Leica Microsystems).

GFP transgene amplification
Two primary transformants showed strong amplification for the expected 982-bp PCR product (Fig. 2). Of the five T 1 progeny assayed, three (T 1 -1, T 1 -2, and T 1 -3) failed to show amplification for the GFP PCR product; however, two others (T 1 -4 and T 1 -5) did produce a band of the expected size (Fig. 2). Genomic DNA from one wild-type plant, two primary transformants, and all T 1 plants was amplified for the presence of ACTIN as a control for DNA quality, and all showed the expected band (Fig. 2).

Fluorescence
The abaxial leaf surface from two of the primary transformants was imaged for GFP fluorescence and both individuals showed consistent and uniform fluorescence across the leaf epidermis; however, fluorescence was greater in the vasculature than in the epidermal tissue (Fig. 3). Adaxial leaf tissue also displayed uniform fluorescence. Tissue from all four floral whorls, immature fruits, and stem cross sections were also imaged. Stamens and pistils showed very strong fluorescence, as did nectaries and pollen. Fluorescence was very weak but detectable in the sepals and petals (data not shown). No visual evidence of mosaicism was observed. Although the endoplasmic reticulum-localized GFP transgene (erGFP) reporter construct was designed in part for its even expression in Arabidopsis root tissues, it is expressed in all aerial tissues of the plant. Because we grew many of our plants in soil and not on agar plates, we chose the easier, aboveground tissue for screening and did not image belowground tissue for fluorescence.
The GFP transgene was not detected in three T 1 progeny (T 1 -1, T 1 -2, and T 1 -3), and these also failed to show fluorescence above background levels. However, the two T 1 plants that did show PCR amplification of the GFP transgene also showed fluorescence similar to the primary transformants. As observed in the primary transformants, GFP fluorescence was very strong in the stamens, pistil, pollen, and nectaries, and moderate fluorescence was consistently observed in the leaf tissue.
Wild-type plants did not show fluorescence in leaf, stem, and most reproductive tissues. Background fluorescence was elevated in anthers and stigmatic tissue, identical to that seen in the anthers and stigmas of non-transgenic T 1 plants.

DISCUSSION
Although GFP signal was clearly visible, the relatively low GFP fluorescence observed, especially in leaf tissues, could be due to a number of factors. The GFP transgene used in this study is endoplasmic reticulum-localized and driven by the Arabidopsis UBIQUITIN 10 (pAt4g05320) promoter. Because of the comparatively large vacuoles in many plant cells, the endoplasmic reticulum is often pressed against the cell membrane, making the GFP signal in a single cell dense; however, across a given tissue, the signal will potentially appear more diffuse. Additionally, it has FIGURE 2. PCR amplification of a 982-bp region of the erGFP transgene (top row) and a ~650-bp region of the ACTIN control (bottom row) in a wild-type plant (WT), two primary transformants (T 0 -1 and T 0 -2), five progeny of the primary transformants (T 1 -1 through T 1 -5), the vector used for transformation (Plasmid), and a negative control (NTC). All lanes with Datura DNA amplify for ACTIN, with the band falling between the 650-bp and 850-bp points on the ladder. Only the primary transformants, two progeny (T 1 -4 and T 1 -5), and the transformation vector amplify for the erGFP region, with a band falling between the 850-bp and 1000-bp points on the ladder. been reported that, when present in the oxidizing environment of the endoplasmic reticulum lumen, GFP folding can be disrupted and promote the formation of disulfide bonds between GFP molecules, potentially reducing fluorescent intensity (Jain et al., 2001;Aronson et al., 2011). We have successfully regenerated transgenic plants from callus tissue of D. stramonium and demonstrated stable inheritance of the GFP transgene. To our knowledge, this is the first report of stable transformation and transgene inheritance of any species in the genus Datura, and represents an important tool for genetic studies in this evolutionarily important genus. Availability of methodology for recovery of stable transgenic lines is a critical first step for Datura gene function studies through approaches such as overexpression and gene editing by CRISPR/Cas9 or other editing technology.

ACKNOWLEDGMENTS
This work was funded by a National Science Foundation grant to A.L. (IOS 1456109), a Department of Education Graduate Assistance in Areas of National Need (GAANN) grant to the University of California, Riverside, and by a Research Experience for Undergraduates Grant to the Center for Plant Cell Biology at UC Riverside.

AUTHOR CONTRIBUTIONS
A.C.R., A.L., and J.V.E. wrote the manuscript; A.C.R., K.B.E., and A.H. conducted the experiments; and A.L. and J.V.E. advised on experimental design.