In the era of genome editing, the protoplast method has started playing a pivotal role in facilitating researchers in many dimensions, including efficient delivery of genetic material, quick assessment of editing strategies and design, generating transgene-free mutants, and high throughput screening.
Protoplast isolation and transient transfection systems have been established in several crops, facilitating functional validation of target genes (Yoo et al. 2007; Li et al. 2016; Lin et al. 2020). Previous protocols for protoplast isolation from green tissue and calli have been established (Yoo et al. 2007; Zhang et al. 2011; Poddar et al. 2020). Although isolated protoplasts from callus are suitable for experimental studies, they require time, and callus culture is susceptible to contamination. Here, we have standardized a rapid, and highly efficient protoplast isolation procedure from etiolated rice seedlings. This protocol is adaptable to other plants as evidenced from our demonstration in dicot species, Arabidopsis, and Chickpeas. There is a direct relationship between protoplast viability and transfection efficiency. Hence, an isolation protocol that yield a high percentage of viable protoplast is always desirable for downstream analysis. Adding a centrifugation step with 0.55 M sucrose greatly facilitated to accumulation of viable protoplasts in the interface while removing cellular debris (Figs. 2 and 3). Since DNA may transform nonviable protoplasts, which however would not render transgene expression. As a result, transformation efficiency is reduced. The use of the sucrose step drastically reduces the percentage of dead and broken protoplast in the isolated sample (Figs. 2 and 3). Sucrose overlaying step has been used successfully in several earlier studies for isolating protoplasts from different plants (Brandt et al. 2020; Poddar et al. 2020; Jeong et al. 2021). The sucrose step also showed significant improvement of transfection efficiency in rice protoplasts over the protocol where the step is not included (Fig. 5).
Performing protoplast isolation and transfection on the same day can be a time-consuming and demanding process. It also poses limitation on the number of transfections that could be handled. Also, it is not uncommon that midi-prep of some plasmids may fail in the first attempt. Preserving protoplast viability for a minimum of 24 hours would offer valuable flexibility to researchers, allowing them to take a break between the isolation and transfection steps. Our result showed that the isolated protoplasts can be stored in MMG solution for 24 hours at 25°C with minimal loss of viability and they can be very well transfected even after 48 hours (Fig. 6). An earlier study showed preserving viability of protoplast isolated from rice calli tissue (Poddar et al. 2020). Our study for first time showed the storability of isolated protoplasts from rice seedlings. Culturing rice embryos to generate calli tissue involves the risks of contamination and is a time-consuming process. In contrast, utilizing rice seedlings as the starting material for protoplast experiments offers numerous advantages. Our protocol with a pause step between protoplast isolation from seedling and transfection would allow handling larger number of transfections on the following day or conducting another round of midi-prep for failed constructs.
Protoplast-based transient expression systems have been employed for functional genomics studies (Lorenzo et al. 2019), and a transfection efficiency of at least 50% is considered reliable (Yoo et al. 2007). We have optimized various factors for PEG mediated protoplast transfection. In this study, we achieved highly efficient transfection (81%) in rice protoplast isolated from 10 days grown etiolated rice seedlings using 30 µg of plasmid DNA, and 20 minutes of incubation in a PEG solution. Notably, we have also tried with etiolated seedlings of both Arabidopsis and Chickpea but etiolation causes problems in plants viz. abnormal growth of both stems and leaves, weakens the cell walls, elongates the internodes with fewer leaves than normal condition. Therefore, it was difficult to obtain sufficient leaves when grown in dark. Here, we have chosen light grown 20–25 days old green leaves for protoplast isolation for both dicot plant species. In an earlier study, transfection efficiency was observed to be nearly 53–75% (Zhang et al. 2011). Similarly, up to 73.5% efficiency was achieved in protoplast isolated from rice calli (Poddar et al. 2020). We have observed superior transfection efficiency with smaller size of plasmids (Fig. 4D-H). Similar to the present investigation, maximum transfection efficiency (75%) was recorded with small sized plasmids, whereas larger binary plasmid obtained 45–66% efficiency (Zhang et al. 2011). Amount of plasmid DNA has a great impact on transfection efficiencies. In our study, we have tested the transfection efficiencies with different concentration of plasmid DNA, and observed highest number of transfected rice protoplasts (81%) with 30 µg of plasmid DNA. Comparatively lower (30–60% in rice) percentage of transfected protoplasts were recorded with 10–25µg of the plasmid DNA. In our study, we also observed that transfection efficiencies for etiolated rice protoplasts were 10–20% more compared to the green protoplasts.
Genome editing using CRISPR/Cas9 generates mutations at specific target sites. However, not all the CRISPR vectors and gRNAs are active and efficient. Given that plant regeneration using tissue culture method in monocot like rice and dicots is a time-consuming process, so, it becomes crucial to validate the efficiency of sgRNA before performing stable transformation. Our CRISPR-deletion strategy using two sgRNAs showed high applicability in indicating successful editing on agarose gel. This method allows for rapid validation of sgRNAs and Cas proteins, and different editing strategies. CRISPR-induced successful mutations were deciphered using PCR, gel electrophoresis, and Sanger sequencing from transfected protoplasts after 72 hrs of transfection. We have demonstrated successful genome editing with our protocol in all three target genes by detecting additional smaller PCR bands with the WT band (Fig. 7A, B, and C). The appearance of a smaller band is an indication that both the sgRNAs are capable of inducing DSB causing deletion of the intervening region. The result was confirmed by Sanger sequencing.
Genome editing using CRISPR-cas9 has been demonstrated successfully in several legumes (Wang et al. 2016; Meng et al. 2017; Ji et al. 2019; Liu et al. 2019). In spite of this accomplishment, the recalcitrant nature of in vitro genetic transformation and stable regeneration of chickpeas remain a serious bottleneck for the implementation of genome editing tools in this most important and nutrient-rich crop (Das Bhowmik et al. 2019). Protoplast transformation using CRISPR-Cas9 targeting drought-associated genes was reported recently (Badhan et al. 2021). Here, we have demonstrated the applicability of our isolation and transfection protocol in chickpeas and Arabidopsis. Similar to rice, the CRISPR-deletion approach yielded expected deletion in one Arabidopsis gene and two chickpea genes (Fig. 8C, D, and F). A handful of reports are available that show editing experiments in chickpeas (Gupta et al. 2023). This protocol will facilitate researchers to undertake genome editing in chickpeas. Thus, our work offers an effective strategy to easily and rapidly validate the CRISPR reagent sites, and it will endorse an application for genome editing in both monocots and dicots.
Standardisation of protoplast regeneration will help avoid potential problems such as chimerism which can be seen during transient transformation using Agrobacterium or particle bombardment (Reed and Bargmann 2021). Regenerated plants from protoplast are of single-cell origin and avoid such problems. Additionally, protoplast transformation efficiencies are higher compared to other transformation processes, increasing the likelihood of successful editing events in plants transformed with editing reagents. Our protocol should be compatible with other CRISPR systems, such as CRISPR-Cas12a and precise base editors and prime editors (Zhang et al. 2019; Molla et al. 2021). This method could also be used for RNP transfection for generating transgene free mutants (Zhang et al. 2021). Encouragingly, recent studies showed that RNP delivery of CRISPR-Cas12a could generate nearly 100% genome editing efficiency in protoplasts from different plant species (Zhang et al. 2022), which translated to high-efficiency genome editing in regenerated plants such as citrus (Su et al. 2023). We hope that the protocol described here will encourage others to adopt it for application in their favorite plant systems which will open many unknown potentials.