A sustainable protoplast isolation method with optional pause points
Existing methods for protoplast isolation from rice using germinated seedlings and suspension cultures are valuable and well described [19-25]. However, the protocol using germinated seedlings consumes seeds at a high rate and the suspension culture protocol requires the technical and labor-intensive know-how of maintaining a suspension culture. In contrast, the present protoplast isolation protocol using rice calli described here is highly effective and accessible because callus induction from mature seeds is straightforward and fast. We could generate about 5 g of calli 1 month after callus induction from 30 mature seeds. These tissues can be proliferated to about 30-40 g of calli after another month, and 150 to 200 g of calli are available 3 months after the initial callus induction. These tissues can be further proliferated with additional rounds of transfers and can be utilized at any stage on a large scale. Furthermore, the methods using germinated seedlings and suspension cultures require an uninterrupted and lengthy workflow from donor tissue digestion through transfection performed on the isolated protoplasts.
Here, techniques are outlined for a branched method with built-in optional pause points that allow for consistent and efficient procurement of healthy protoplasts that may be used gradually, over the course of several days, for downstream transient assays. The donor tissues for the isolation of protoplasts are calli induced from seeds and regularly sub-cultured on solid OsCIM2 callus induction media (Table 1). Calli were also induced from immature embryos in this manner, with comparable outcomes. In general, callus tissue propagated for more than six months could be used in this method. As a result, the donor tissue becomes available at an exponential rate once initiated, abrogating the obstacle of donor material availability for this procedure.
The workflow for the protoplast isolation protocol produced from this study is portrayed in Fig. 1. We gathered 5 g of compact pale-yellow rice callus tissue (Fig. 2A) and used a scalpel or metal spatula to bring all of the pieces to roughly the same size. Careful slicing and razor exchanges were not needed. An enzymatic cocktail of 1.5% cellulase R10 and 0.1% pectolyase or 0.75% macerozyme R10 resulted in a successful breakdown of rice callus tissue cell walls while maintaining healthy viable protoplasts (Figs. 2B, C). An additional step of vacuum infiltration of the digestion solution with the donor tissue, an approach utilized in other protocols [22,30], was unnecessary for our method, eliminating a common step, decreasing equipment load, and increasing simplicity. Rather, we could simply incubate the callus tissue with 15 mL digestion solution with gentle shaking at 70 rpm for 3 hours, less time than is required for seedling-derived cells. After digestion, protoplasts were isolated from spent tissue via filtration (Fig. 2D) and centrifugation through a 0.55 M sucrose cushion. A gentle overlay of the cell suspension onto the sucrose was found to be a crucial step for optimal yield. If the cells were handled crudely and dropped with a force that significantly broke the surface tension of the sucrose, the ultimate protoplast yield could be diminished. After centrifugation of the cell suspension through 0.55 M sucrose, healthy protoplasts separated from debris and accumulated to form a dense band of purified protoplasts at the W5 - sucrose interface (Fig. 2E), bringing the protocol to its first optional pause point.
Here, the method could be paused for one or more days. The band of protoplasts could be left undisturbed at the interface for processing at a later time or handled immediately. The protoplasts produced from the utilization of this pause point are termed “S protoplasts.”
To isolate protoplasts from the sucrose cushion, the entirety of the cloudy phase band containing the protoplasts was gently pipetted out. This was followed by final washing and centrifugation steps to maximize the purity of the protoplasts and eliminate cellular debris. Finally, protoplasts were resuspended in MMG solution to a concentration of 2.5 x 106 protoplasts/mL. Protoplasts could be transfected immediately, or the second optional pause point could be employed—storing the protoplasts in MMG solution, termed “M protoplasts,” until transfection at a later time.
Protoplasts isolated from the band on the same day as digestion of the donor tissue cell walls were referred to as “Sucrose Cushion Day 0/ MMG Day 0” (S/M0) protoplasts. Those isolated from the interface one, two, three, or 7 days after digestion were designated S1, S2, S3, or S7 protoplasts, respectively. S/M0 protoplasts, stored in MMG solution and utilized in experiments over the following one, two, three, or 7 days after digestion were labeled “MMG Day 1” (M1), M2, M3, or M7 protoplasts.
Protoplast viability over time
To ensure the utility of this branched method for protoplast isolation, Evans blue staining was used to quantify viable protoplasts in all isolations from the sucrose cushion (S protoplasts) as well as protoplasts stored in MMG solution over time (M protoplasts) (Fig. 2F). Healthy intact protoplasts derived from this method are colorless, spherical, and resistant to staining. The viability assay indicated that S/M0 isolates yielded the greatest number of live protoplasts, with a gradual decrease with the increasing age of the protoplast-containing sucrose cushion. S/M0 isolates contained approximately 2.5 times the number of protoplasts as S7 isolates. However, it is notable that the order of magnitude for the protoplast count remained unchanged between S/M0 and S7. We show that from 5 grams of rice callus donor tissue, this method yields, on average, 9.8×106 live protoplasts if isolated on day 0 (S/M0) and 3.9×106 live protoplasts if isolated on day 7 (S7) (Fig. 3A). This translates to approximately 20 transfection reactions with S/M0 protoplasts, and 8 transfection reactions with S7 protoplasts. To compare, 100-120 finely sliced rice seedlings are required to obtain approximately the same number of protoplasts [22] as an S/M0 isolation using 5 g of calli by the current method (Fig. 3A), and which cannot be stored for future use.
Viability of M protoplasts was also tracked over time. Though viability decreases appreciably from day 2, the concentration of viable M1 protoplasts is comparable to S1 protoplasts and only mildly reduced from S/M0 protoplasts, making them an acceptable option for use in assays (Fig. 3A). It was also noted that protoplasts held in MMG solution for 7 days appeared approximately 1.7X larger (Fig. 3B), and S7 protoplasts were observed to be enlarged to a similar or greater extent. This may be attributed to cell growth or osmotic swelling. However, the larger protoplasts displayed a characteristically healthy spherical shape, unstained by Evans blue, suggesting that osmotic stress was not occurring.
Transfection efficiency over time
Both quantity and quality of protoplasts are critical factors for downstream experiments. In existing methods, transfection is performed only on freshly isolated protoplasts. Here, the transfection efficiency of both S and M protoplasts of different ages were assayed via PEG-mediated transfection of pAct1IsGFP-1, a GFP overexpression plasmid [27].
First, S and M transfection pools were imaged for GFP expression 1 and 2 days after transfection (Fig. 4). Protoplasts aggregate over time and the 1 mL pools were not pipetted for homogeneity. Rather, a fluorescence stereomicroscope was used to manually scan the sample and gather representative images in areas with a moderate density of protoplasts. Strong GFP fluorescence was detected in both S and M cells 1 and 2 days after transfection.
Transfection efficiency was calculated 2 days after plasmid transfection as a percentage of GFP positive protoplasts over total live protoplasts, as determined by fluorescence microscopy and Evans Blue staining on a hemocytometer (Fig. 5). For S/M0 protoplasts, transfection was highly efficient, producing 73.5% GFP expressing protoplasts. S1 transfection efficiency was comparable, at 69.5%. Taken together with the results from the previously described viability assay (Fig. 3A), the data suggest that S1 protoplasts are comparable in value to freshly isolated S/M0 protoplasts. This finding facilitates novel flexibility in research, allowing assays to be performed 24 hours after initiation of the protoplast isolation method with little to no loss of efficacy and data. Though transfection efficiency declines over time for both S and M protoplasts, it does not fall below 15% within 7 days (Fig. 5). Moreover, it is conceivable that certain assays, for example protein localization, do not require optimal transfection efficiency or viability. The data provided here allow for the informed design and versatile scheduling of protoplast experiments with a quantified summary of expected losses of viability and transfection efficiency over time.
Gene editing via Cas9 ribonucleoprotein transfection
Given the flourishing field of genome editing, it was critical to ensure that this method was suitable for such studies. To demonstrate this, S/M0 as well as S1 and M1 protoplasts were transfected with in vitro assembled Cas9-gRNA ribonucleoproteins targeting a single locus (Fig. 6). For comparison with protoplasts isolated via a previously published method [22], protoplasts derived from rice seedlings were also transfected. Editing at the Cas9 cleavage site was identified and quantified through NGS. Editing rates for S/M0 protoplasts and seedling-derived protoplasts were similar (Fig. 6), indicating that the present protocol can be used confidently in genome editing studies. It is notable that PEG-mediated transfection of plasmid DNA encoding CRISPR/Cas9 components has also been utilized for similar purposes in other protoplast systems [31, 32] and would be an effective alternative to ribonucleoproteins in the present method (Fig. 6).