Protoplast isolation and transformation
Many experimental approaches, such as biochemical, protein-protein or protein-DNA interaction analyses do not require fully-grown plants and protoplasts are suitable experimental systems. We thus developed a protocol for rapid protoplast isolation and transformation (online resource 3, step-by-step protocol). Protoplasts were isolated from the leaf and hypocotyl tissue of 10 days old in vitro grown plants using macerozyme and cellulose R10 to digest cell walls. The protoplast isolation yielded between 0.45 and 0.85 x 106 protoplasts / g plant tissue. These protoplasts were round, only occasionally was the cell wall still attached, and they included many chloroplasts (Fig. 1 A and B).
Next, we transformed the freshly isolated protoplasts with a nuclear localized, ubiquitously expressed GFP protein as fluorescent marker driven by CaMV 35S promoter (p35:GFP:NLS). Employing a PEG-mediated transformation protocol, the transformation efficiency was between 13.5 % and 24.8 % (Fig. 1C). The transformation efficiency is dependent on the incubation time of the enzyme solution, such that an incubation time of 1.5 h yielded 11.8% transformed protoplasts and 4 h 24.8 % transformed protoplasts (Fig. 1D). Incompletely digested cell walls may thus inhibit the PEG-mediated transformation. Nuclear localized GFP signal was detected using fluorescent microscopy 64 h post transformation (Fig. E-G). Protoplasts were visually unchanged (Fig. 1E) when compared to freshly isolated protoplasts (Fig. 1A) suggesting that the transformation and subsequent incubation time had no obvious negative effect on protoplast viability. The protoplasts showed a very low level of auto fluorescence (Fig. 1F). Suppl. Figure 2 provides a flowchart summarizing the experimental steps for protoplast transformation and all other approaches described here (Online Resource 1, Fig. 1).
Plant regeneration from explants
Establishing E. californica as a genetic model plant requires the possibility for stable transformation and thus we developed a reliable, tissue-culture based method for plant regeneration as a first step towards a transformation protocol (Online Resource 1, Fig. 1).
We first analyzed the type of explant, and type and concentration of auxin best suitable for callus regeneration of E. califonica (Table 1). Using 2.4-D as auxin source resultsed in all used concentration in callus regeneration, with hypocotyl being more amenable to callus regeneration. A concentration of 1 mg/l 2.4-D was sufficient for regeneration of callus tissue from every explant. After four weeks, almost all hypocotyl explants regenerated callus (97.8 %) that were subsequently induced to develop somatic embryos (Online Resource 2, Tables 1 and 2). After seven weeks, 44% of the calli produced somatic embryos and after this time, only marginally more calli generated somatic embryos (Online Resource 2, Table 2). The somatic embryos were removed from the calli and matured for three weeks (Online Resource 2, Table 3) on a specific maturation medium including maltose instead of sucrose and subsequently, they were kept in the light until they turned green and commenced organogenesis. From 58% of the somatic embryos, plantlets could be regenerated, and 60% of these plantlets developed into fully grown plants in soil (Online Resource 2, Tables 3 to 5). The regenerated plantlets were transferred to soil after four to six weeks (Online Resource 2, Table 6). The surviving plants formed morphologies similar to plants grown from seeds and were fully fertile. Generally, the regeneration of E. californica plants was carried out in at least five repititions.
Taken together, we established an efficient E. californica in vitro plant regeneration protocol, from hypocotyl explants to mature plants via callus induction and somatic embryogenesis. Figure 2 shows an overview of the time scale of E. californica regeneration.
Optimization of the plant regeneration protocol
To obtain an efficient and reproducible protocol for E. californica transformation, several modifications to the transformation protocol reported earlier (Park and Facchini 2000b) were tested. For example, the addition of 2.0 mg/l NAA and 0.1 mg/l BAP to the callus induction medium was replaced by adding only 1 mg/l 2,4 D, which raised the number of explants producing calli to nearly 100 % (Table 1). Further, we analyzed if cotyledons or hypocotyl produced callus material more reliably and found that with the optimal concentration of 1 mg/l 2,4-D, all hypocotyl explants produced callus while only max. 80% of the cotyledon explants produced calli (with an optimal 2,4-D concentration of 1,5 mg/l, Table 1). Next we analysed if unripe seeds (21 days after polination) have the potential to regenerate callus. We compared 1 mg/l 2.4-D with 2 mg/l NAA and 0.1 mg/l BAP and our results show that unripe seeds have a lower callus regeneration rate than cotelydons and hypocotyl, independend of the auxin source (Table 1).We also analyzed if reducing cell culture inhibitory secondary metabolites by adding 0,5 g/l activated charcoal to the cell culture medium improves callus formation (Möller et al. 2006; Thomas 2008). However, the callus regeneration from hypocotyl dropped from 100–33% when charcoal was added (Table 1), suggesting that charcoal may block phytohormone activity. Additionally, we analysed if somatic embryos can dedifferentiate to callus by adding 1 mg/l 2.4-D or 2 mg/l NAA, 0.1 mg/l BAP and activated charcoal. 2.4-D leads to new callus regeneration in more than 90%. Interestingly, the combination of NAA, BAP and activated charcoal leads to the regeneration of new somatic embryos within six weeks in nearly 100% of all samples (Table 1). Finally, a somatic embryo maturation phase was introduced to the protocol to improve greening and proper morphogenesis of the embryos. We analyzed several combinations of maltose, PEG and abscisic acid (ABA), and found that exchanging sucrose as carbon source with 3% maltose alone performed best, with around 84% of the embryos started photosynthesis in contrast to a combination of 3% maltose with 2.5 mg/l ABA which killed around 97% of the calli (Online Resource 2, Table 3). In addition, the regenerated plants grew better in plain standard soil; around 80% survived the transfer from in vitro culture to soil when compared to vermiculite as substrate, which only around 54% of the plants survived (Online Resource 2, Table 6).
Agrobacterium -mediated transformation of E. californica
Based on the tissue culture and regeneration method described earlier, we developed an efficient transformation protocol using E. californica hypocotyl as explant and employing Agrobacterium tumefaciens based transformation to generate transgenic lines carrying the reporter construct p35S::GFP using hygromycin as selectable marker. A killing-curve experiment showed that a concentration of 40 mg/l hygromycin is sufficient to select transgenic plants (Online Resource 1, Fig. 2; Online Resource 2, Table 7). The transgenic calli produced somatic embryos, which were then regenerated into fully grown, fertile plants employing the plant regeneration protocol (online resource 1, Fig. 3). Figure 3 (A-P) shows mock-treated and 35S:GFP treated calli and somatic embryos in bright field and fluorescence microscopy. Only a small fraction of callus cells is transformed and has already started to divide (Fig. 3A to H). Those will later provide the transgenic calli shown in Fig. 3L and P, while the untransformed cells of the calli discontinue division and die eventually. Healthy calli under selection are then removed and incubated on somatic embryo induction medium. Wild type calli regenerating somatic embryos are shown in Fig. 3 I, J, M, and N). Figure 3K, L, O, and P show two fully fluorescent calli that are derived from transformation events. Those generate genetically identical somatic embryos that are also completely fluorescent, showing that the 35S promoter is active in callus tissue and somatic embryos. The presence of transgenic GFP protein in the transformed tissues was corroborated by Western blot analysis (Fig. 3Q). In total, 23 independently transformed transgenic calli were derived that, when cultured on somatic embryo induction medium, all continuously produced genetically identical somatic embryos. A strong GFP signal was detected in 65% of the regenerated somatic embryos (Fig. 3 and Online Resource 2, Table 8).
Unexpectedly, the mock-treated calli and somatic embryos also show a faint fluorescence (Fig. 3B, F, J, N), but this is much lower in intensity than the GFP fluorescence in Fig. 3 C, D, G, H, L, and P, and suggestive of fluorescing secondary metabolites. Interestingly, it was shown previously, that E. californica suspension cultures produce benzophenanthridine alkaloids with an excitation of 270–550 nm and emissions at 310–590 nm (Hisiger and Jolicoeur 2005). Those lie within the range of the GFP excitation and emission spectrum (max. 488 nm and 507 nm, respectively) and could explain the low intensity fluorescence observed in wild type calli and somatic embryos. p35S:GFP transformed plants show a morphology similar to that of wild type plants that underwent tissue culture based regeneration, except for the flowers, which show folded mature petals in the transgenics (Online Resource 1, Fig. 3).
Other E. californica transformation methods
There are other means to generate transgenic E. californica tissue, for example by using biolistic bombardment of callus tissue (Popelka et al. 2003; Viehweger et al. 2006; Angelova et al. 2010), but this requires an expensive biolistic particle delivery system. Using the regeneration protocol reported here, this method can be extended to generated transgenic lines. Another protocol published earlier reports an approach similar to ours, but with different media/phytohormone compositions and paromomycin as plant resistance markers (Park and Facchini 2000b). However, this method is less efficient in terms of callus generation from explants, for example, with our method, around 98 % of explant generate calli, with the previous protocol only 80 %. 30 % of the calli then generated somatic embryos within three to four weeks, while using the method reported here, nearly 45 % of calli developed somatic embryos within seven weeks, indicating that our protocol is more efficient than the previous one. Further, the somatic embryo induction medium we used induced continuous production of somatic embryos from transgenic calli, such that up to 37 transgenic calli could be subcultured from a single callus (Online Resource 2, Table 10). Unfortunately, we cannot compare our work with how the method by Park and Facchini (2000b) performs in other labs, as no further work based on this protocol was published.