Most of the currently available methods for protein subcellular localization can be readily applied to the majority of model plant species such as Arabidopsis and tobacco, which could, in principle, lend themselves as heterologous recipients for localization studies of proteins from non-model plant species [18, 19]. However, the relatively high failure rates reported in studies adopting such heterologous approaches (e.g., for Arabidopsis proteins expressed in tobacco) indicate that homologous expression systems are preferable whenever possible [10, 20]. The lack of established regeneration protocols for the large majority of non-model plant species hinders the application of whole-plant stable transformation methods for protein subcellular localization studies, which rely mainly on transient transformation approaches. The latter methods, however, are limited with respect to the level of detail to which localization can be assessed, as subcellular fractionation or sectioning and immunodetection methods cannot be applied, thus seriously preventing analyses of functional characterization of genes in non-model species. Therefore, adaptation of methods for the stable transformation of cell cultures that are currently applied mainly for functional studies [21, 22] to protein subcellullar localization analyses would provide two main advantages for cases dealing with non-model species: (1) the faithful representation of native localization patterns that are typical of homologous systems, and (2) the versatility, robustness, and virtually unlimited amount of material available for characterizing stable transformation methods. As a case study for the development of such an approach, we chose to use P. tenuiflora, a non-model halophite grass belonging to the Poaceae family that has been the subject of several previous studies owing to its ability to grow in soils with extremely high salinity and pH [23–26]. Given that P. tenuiflora is recalcitrant to regeneration, it can be conveniently used as a representative case for the proof of concept of the novel method.
Establishment of suspension cell lines of P. tenuiflora
To establish P. tenuiflora suspension-cultured cells, a callus was induced by dedifferentiation of mature P. tenuiflora seeds. In brief, among the calli with different morphologies induced on the seeds of P. tenuiflora, we chose a yellow, loose callus (Figure 2A) with a clear nucleus and without visible intercellular aggregations (Figure 2B) for subculture. Unlike the standard method used for filtering rice [21], dispersed cells were obtained by slight grinding of the callus. The obtained cell clusters were mixed with liquid culture medium and cultured for 15 d. The culture conditions were optimized to produce the suspension-cultured cell system of P. tenuiflora as described below.
Optimization of the liquid medium
We optimized the liquid medium in terms of concentration of sucrose (Figure 3A), amounts of 2,4-D (Figure 3B), pH (Figure 3C), and types of nitrogen nutrients (Figure 3D) added in a total volume of 50 mL of MS medium. After 15 d of cultivation, the dry weight of the harvested cells was measured. The dry weight of P. tenuiflora suspension-cultured cells was found to be highest in MSPECH medium (pH = 5.8) containing 30 g/L sucrose and 2 mg/L of 2,4-D (Figure 3), which was thus chosen as the optimum medium for the culture of P. tenuiflora suspension-cultured cells.
Growth performance of P. tenuiflora suspension-cultured cells
In order to understand the growth dynamics of P. tenuiflora suspension-cultured cells, we characterized their dry weight in terms of culturing time under the optimized conditions. Figure 4A shows the growth curve of the P. tenuiflora suspension-cultured cells at different growth stages. Suspension-cultured cells (10 mL) were inoculated into 50 mL of MS medium, and the dry weight of the suspension was determined to be approximately 40 mg. After 5 d of cultivation, the growth rate increased and stabilized at ~15 d of cultivation, after which the cells started to show signs of aging, thus prejudicing further tests. Furthermore, as shown in Figure 4B, the cells were well dispersed, and the small cell clusters presented inclusions and uniform appearance during the 5–15 d of cultivation. Our results showed that during the 5–15 d of cultivation, the cultured cells were stable and reached a high dry weight, which were suitable conditions to perform Agrobacterium transformation.
P. tenuiflorasuspension-cultured cells are a stable genetic transformation system
The Agrobacterium-mediated transformation was used to establish the genetic transformation system of the P. tenuiflora suspension-cultured cells. The pBI121-ER-GFP plasmid was used to transform the P. tenuiflora suspension-cultured cells. The transgenic cells were screened on a solid medium containing the appropriate marker. To ensure high transformation efficiency, the Agrobacterium concentration, AS effect, co-culture time during transformation, and working concentrations of Kan and Cef were optimized as described below.
Agrobacterium concentration
Considering that Agrobacterium concentration can affect the transformation efficiency, four different concentrations of Agrobacterium were tested: OD600 = 0.5, 1.0, 1.5, and 2.0. Although Kan-resistant cell clusters grew at Agrobacterium concentrations of an OD600 of 0.5, 1.0, and 1.5, the growth of the cell clusters was negatively affected at higher concentration (OD600 = 2.0). Therefore, we suggest that Agrobacterium concentration for genetic transformation should be kept as low as possible (an Agrobacterium concentration corresponding to OD600 = 0.3 was suitable for genetic transformation; data not shown). In addition, the length of co-culture during the transformation was also important for obtaining high transformation efficiency, as the bacteria were observed to be completely removed from the suspension cells after 3 d of co-culture (data not shown). Therefore, 3 d of co-culture was chosen as the optimum culturing time in order to obtain high transformation efficiency. Pandey et al. reported that optimal β-glucuronidase expression was observed in cumin embryos co-cultivated with an Agrobacterium cell suspension at an OD600 of 0.6 for 72 h [27], while the optimal transformation efficiency of suspension-cultured Glycyrrhiza inflata Batalin cells was achieved using an Agrobacterium suspension of an OD600 of 0.4 over 24 h of co-cultivation [22]. These results suggested that co-cultivating the lowest concentration of Agrobacterium possible with plant suspension-cultured cells for approximately 3 d could provide optimal conditions for achieving highly efficient transformation.
Optimization of the concentrations of Kan, Cef, and AS
The working concentrations of Kan and Cef were examined. Figure 5A shows that the dry weight of the suspension-cultured cells significantly decreased with increasing concentrations of Kan, and that the working Kan concentration of ~75 mg/L was most appropriate for this study. Moreover, the dry weight of the suspension-cultured cells slightly decreased with increasing concentrations of Cef. However, since the growth inhibition at 200 mg/L Cef was only slightly reduced compared to that at 300 mg/L Cef (Figure 5B), the optimum concentrations of Kan and Cef were determined to be 75 mg/L and 200 mg/L, respectively.
AS is a commonly used agent affecting Agrobacterium-mediated plant genetic transformation. To test the AS-mediated effects on the Agrobacterium-mediated genetic transformation of P. tenuiflora suspension-cultured cells, 3-d-old AS-pretreated suspension cells were co-cultured with Agrobacterium at OD600 = 0.5 for 3 d; non-pretreated cells were used as the control. A slight increase in the number of cell clusters from the AS-treated suspension cells on the solid medium was observed compared to that in the control (data not shown), suggesting that AS pre-treatment could improve the efficiency of infection of P. tenuiflora suspension-cultured cells.
Subcellular localization of proteins
In order to carry out a proof of concept study for the use of stably transformed P. tenuiflora suspension-cultured cells in the analysis of the subcellular localization of proteins, the plasmid pBI121-ER-GFP, encoding an ER-GFP fusion protein was used for genetic transformation. First, the calli were inoculated into MSPECH culture medium containing 2 mg/L 2,4-D and 30 g/L sucrose (pH 5.8) for 15 d. After 3 d, AS-pretreated cells were co-cultured with Agrobacterium at OD600 ≈ 0.3 for 3 d, and the transgenic cell lines were selected on solid MSPECH medium containing 75 mg/L Kan and 200 mg/L Cef by observing them with a stereomicroscope (Olympus, SZX9, Japan). Green fluorescent cells were manually selected for subculture to obtain highly pure P. tenuiflora transgenic cell lines according to previously published protoplast isolation methods, with slight modifications for improvement [28–30]. After 5 d of cultivation, the GFP signals of the transgenic protoplasts were mainly concentrated on the plasma membrane and at the ER in association with the nuclear periphery, which is consistent with the subcellular localization pattern previously established for the PutAMT1;1 protein [17] (Figure 6A-B). Given the availability of stably transformed P. tenuiflora cell lines expressing the PutAMT1;1-GFP fusion protein, we plan to carry out further studies using subcellular fractionation and immunogold electron microscopy to determine the localization of this protein in its native cellular environment.
Taken together, the results presented herein indicate that the P. tenuiflora suspension-cultured cell system could be successfully established and employed for protein subcellular localization analysis. Although previous studies reported that this approach may not be suitable for subcellular localization and other fluorescence-based analyses [31, 32], we demonstrate that this optimized expression system based on P. tenuiflora suspension-cultured cells proved to be simple, reliable, and stable. Nevertheless, like all transformation methods relying on cells isolated from single organs without intervening regeneration of whole plants, this method can only be applied to examine the localization of proteins in cell compartments/structures that are present in the recipient cells. Therefore, the method is not suitable for studies requiring localization to the cell walls or plasmodesmata, for example, or to track protein movements in the context of organized tissues or organs. We note, however, that application of this method to other cell types deriving from different tissues such as the leaf, root, and inflorescence requires further validation to best ascertain the scope of applications of the method. In comparison to transient assays, our method is more labor-intensive and time-consuming, and thus does not lend itself readily to application in high-throughput studies [10]. However, the method does have an important advantage of providing homogeneous populations of transformed cells in virtually unlimited amounts for extended periods of time that can be used in all localization assays, which cannot otherwise be carried out on transiently transformed protoplasts owing to their complexity or particular technical requirements (e.g., cell fractionation, embedding and sectioning, immunolocalization [33, 34]). Furthermore, we expect that our stable transformation method could be suitable for dynamically documenting protein re-localization through the different phases of the cell cycle or in response to environmental cues, although this was not directly tested in the present study. Most importantly, the fact that this method does not rely on whole plant regeneration makes it applicable to any non-model plant species lacking suitable regeneration protocols, which is a substantial advantage. As the method is based on simple, species-specific optimization steps of protocols that were previously developed for functional analyses in a relatively distantly related monocot species (rice), we expect that, with minor modifications, the method could be applied to several other non-model species from the Poaceae family.