Optimized and Reliable Protoplast Isolation for Transient Gene Expression Studies in the Gymnosperm Tree Species Pinus densiflora

Abstract

Efficient protoplast isolation and gene transfection remain significant challenges in gymnosperms, particularly in Pinus species, where stable transformation is highly limited. Conventional pine protoplast preparation methods have resulted in extremely low transfection efficiencies, hindering functional genomic studies. This study presents an optimized method for isolating high-yield, viable protoplasts from Pinus densiflora (Korean red pine), providing a robust system for transient gene expression assays. Splitting one-month-old cotyledons produced the highest mesophyll protoplast yield (5.0 × 10⁶ cells/g FW), which further increased to 1.2 × 10⁷ cells/g FW after optimizing the enzyme mixture (4.5% cellulase, 0.7% pectinase, 3% hemicellulase), maintaining viability above 86%. Developing xylem and whole-stem protoplasts were also successfully isolated by mitigating resin leakage and debris contamination, with a 17% sucrose gradient yielding 7.4 × 10⁴ cells/g FW at 81.9% viability. Overcoming prior inefficiencies, this protocol significantly enhances gene transfection efficiency, achieving 94.1% GFP transformation with 82.9% viability. Furthermore, transient activation assays confirmed strong activation of pine-derived reporters by native effectors, underscoring the assay’s suitability for studying gymnosperm-specific gene regulation. Given the limited stable transformation strategies available for Pinus species, this optimized protoplast transient gene expression system provides a practical and reliable platform for transient gene expression analysis, offering valuable opportunities for studying gene function and regulation in gymnosperms.

1. Introduction

Protoplasts, plant cells without cell walls, are totipotent and versatile, therefore being a powerful platform for genetic and molecular research [1,2]. The removal of the cell wall enables the efficient delivery of foreign molecules such as DNA, RNA, and proteins, facilitating both transient expression and stable transformation. Transient expressions allow for rapid and temporary gene expression over a short period (e.g., hours to days), while stable transformation involves the insertion of foreign genes into the plant genome for long-term expression and heritability [3].

Protoplast-based transformation methods typically employ polyethylene glycol (PEG)-mediated transfection, electroporation, or microinjection and have been widely used for molecular breeding, genome editing (e.g., CRISPR-Cas9, TALEN), gene silencing (e.g., dsRNA, miRNA, siRNA), transcriptional studies, transcriptomic analyses (e.g., single-cell RNA-seq), and intact plant regeneration via cell fusion and in vitro culture [4,5,6]. Recent advancements in automation have enabled high-throughput protoplast isolation and transformation using robotic systems, further enhancing efficiency and reproducibility [7].

Despite significant progress in protoplast isolation and transformation in angiosperms, the development of efficient protocols for gymnosperms remains a major challenge. Protoplasts from gymnosperm species, including Pinus and Picea, have been isolated from various sources such as immature seedlings, somatic embryos, cell suspensions, and developing xylem tissues [6,8,9,10]. However, these conventional gymnosperm protoplast isolation methods have been largely inefficient for gene transfection, particularly in Pinus species. The combination of thick epidermal layers, complex secondary metabolites, and the release of phenolic compounds and resins inhibits enzymatic digestion, necessitating extended digestion times that ultimately reduce protoplast yield and viability [11]. Moreover, previous studies using electroporation-based transient gene expression in pine protoplasts reported limited success, with poor reproducibility and a lack of quantitative efficiency data [12,13,14]. These limitations have severely restricted the application of protoplast systems for functional genomics in gymnosperms.

Angiosperms such as Arabidopsis thaliana [15], Nicotiana tabacum [16], and crops like Oryza sativa and Zea mays [17,18] have benefited from well-established, high-yield protoplast isolation protocols, primarily utilizing mesophyll tissues. However, gymnosperms present additional challenges due to their unique tissue structures and seasonal dormancy, which limit the availability of suitable plant materials [2,8]. Furthermore, stable transformation in gymnosperms, particularly in Pinus species, is highly inefficient and remains a bottleneck for functional genomics research. Given this limitation, a robust protoplast-based transient transfection system would provide a crucial alternative, enabling rapid functional characterization of gymnosperm-specific gene regulatory networks.

In this study, we aimed to establish a rapid, uncomplicated, and reliable method of isolating high-yield, viable protoplasts from Pinus densiflora (Korean red pine), a representative gymnosperm species. We hypothesized that by optimizing enzyme composition, buffer components, and incubation conditions for multiple tissue types, including mesophyll and woody tissues (developing xylem and whole stems), it would be possible to obtain healthy protoplasts with improved viability and transfection efficiency. Our objective was to develop a robust protoplast-based platform that enables functional genomics and transcriptional studies in gymnosperms, thereby facilitating research into gene regulation, wood formation, and potential genetic improvement in Pinus species.

2. Materials and Methods

2.1. Plant Materials and Growing Conditions

Korean red pine (Pinus densiflora) was used as the source material for protoplast isolation. Pine seeds (Aram Jongmyo Shrine, Seoul, Republic of Korea) were vernalized in water at 4 °C for one week. For in vitro germination, the vernalized seeds were sterilized and cultured on half-strength MS medium (Murashige and Skoog, Duchefa, Haarlem, The Netherlands) supplied with 1% (w/v) sucrose under controlled conditions in a plant growth room (14 h light; light intensity of 150 μmol m⁻² s⁻¹; 23 ± 2 °C). Additionally, vernalized seeds were planted in pots with soil and grown in a growth room. The soil (Topsoil for landscaping, Taeheung F&G, Gyeonggi-do, Republic of Korea) consisted of a mixture of coco peat (40%), peat moss (35%), vermiculite (24.5%), and humic acid (0.5%).

2.2. Protoplast Isolation from Various Pine Tissues

Protoplast isolation was performed using a modified protocol based on methods established for Arabidopsis [15]. Various tissue types, including cotyledons (1-month-old in vitro and potted seedlings), young needles (from 2-month-old potted seedlings, 2-year-old potted plants, and outdoor-grown mature trees collected at Kyung Hee University, Suwon, Republic of Korea; 37.243618, 127.079988), and young stems from adult trees, were tested to optimize protoplast isolation.

For needle tissues, 250 mg of tissue was longitudinally split or chopped, and then immediately placed in 2.5 mL of enzyme solution in a Petri dish. Debarked stem segments (~1 cm diameter) were incubated in 50 mL conical tubes containing enzyme solution, following previously described protocols for developing xylem protoplast isolation [6,19]. For whole-stem protoplast isolation, young stems (~3 mm diameter, 3.5 g) were cross-sectioned (~0.05 mm thick) under distilled water, washed carefully three times with distilled water, and then placed in 20 mL of enzyme solution in a Petri dish.

The enzyme solution for cell wall degradation was freshly prepared with 0.4 M mannitol, 20 mM KCl and 20 mM MES, heated to 65 °C for 5 min, and cooled to room temperature (~25 °C). After cooling, 10 mM CaCl₂, 0.1% (w/v) bovine serum albumin (A7906-100G, Sigma, St. Louis, MO, USA), and enzymes were supplied and passed through a 0.45 µm syringe filter. The enzymes used include cellulase R-10 (Yakult, Honsha, Japan), pectinase (P2401-1KU, Sigma), and hemicellulase (H2125-150KU, Sigma). Tissue samples were immersed in the enzyme solution, vacuum infiltrated for 30 min, and incubated with gentle shaking at 15 rpm at room temperature for 1 to 12 h to determine the optimal incubation duration. Protoplast release and viability were assessed at periodic times during processing. Samples were stained with Wiesner reagent (phloroglucinol-HCl) and toluidine blue O (TBO), as described by Li et al. (2019) [20], and observed under a microscope (KCS3-SS, Optinity, Korea Lab Tech, Seoul, Republic of Korea).

2.3. Protoplast Purification

To purify protoplasts from needle samples, the enzyme solution containing protoplasts was passed through a 150 μm nylon mesh (Miracloth, 475855-1RCN, Sigma) and diluted with an equal volume of wash solution. The mixture was placed on ice for 30 min to pellet the protoplasts. The wash solutions include W5 (154 mM NaCl, 125 mM CaCl₂, 5 mM KCl, and 2 mM MES [15]), W1 (80 mM KCl and 0.4 M sucrose [21]), W2 (half-strength MS medium, 4 mM CaCl₂, and 0.7 M glucose [13]), and W5-modified (W5 supplemented with 5 mM glucose [4]). For stem samples, the enzyme solution was filtered through a mesh and gently layered on sucrose solutions (13%, 15%, or 17%) in 15 mL conical tubes. The tubes were centrifuged (120× g, 15 min.) at room temperature, and the middle condensed-cell layer containing purified protoplasts was collected.

Protoplasts were observed under a microscope and counted using a hemacytometer (Hausser Scientific, Horsham, PA, USA). Viability was evaluated by staining live cells with fluorescein diacetate (FDA; Sigma) and dead cells with propidium iodide (PI; Sigma), following the method of Jones et al. (2016) [22].

2.4. Protoplast Transfection and Transient Activation Assay

Transient transfection of protoplasts was performed using a Polyethylene Glycol (PEG)-mediated method previously established for Arabidopsis [15,23,24]. Transfection efficiency was assessed using a fluorescent assay based on Green Fluorescent Protein (GFP). Healthy protoplasts were diluted in MMG solution (0.4 M Mannitol, 15 mM MgCl₂, 4 mM MES, pH 5.7) to a concentration of 3 × 10⁵ cells/mL. A mixture of 10 µg 35S::GFP/pK2GW7 plasmid (for GFP expression) or 10 µg pK2GW7 plasmid (for control), 100 µL protoplast-MMG solution, and 110 µL PEG-calcium 40% solution (PEG4000 40% w/v, 0.2 M Mannitol, 0.1 M CaCl₂) was incubated at room temperature for 5 min. The protoplasts were then washed with 400 µL W5-modified solution, diluted in 500 µL WI solution (0.5 M Mannitol, 4 mM MES, 20 mM KCl), and incubated overnight. Fluorescent signals were observed under blue light (450–490 nm) using a microscope.

For transient activation assay (TAA), 4 µg of a reporter plasmid (Pro_PdeCesA7::GUS/pTrOX or Pro_AtCesA4::GUS/pTrOX), 5 µg of an effector plasmid (35S::PdeMYB46/pTrOX or 35S::AtMYB46/pTrOX or DW), and 1 µg of NAN plasmid as an internal control were used [23,25,26]. After overnight incubation, GUS and NAN enzyme activities were measured using MUG (4-Methylumbelliferyl β-d-Glucuronide) and MUN (2′-(4-Methylumbelliferyl)-α-D-N-acetylneuraminic acid) as substrates, respectively. Activities were normalized to MU (4-Methylumbelliferone) standards and quantified using a Hoefer TK 100 fluorometer (excitation: 355 nm, emission: 460 nm; Hoefer Scientific Instruments, Bridgewater, MA, USA). The results were expressed as the ratio of GUS to NAN activity, with three biological replicates for each experiment.

3. Results

3.1. Optimization of Mesophyll Protoplast Isolation from P. densiflora

To improve the limitation of current protocols for protoplast isolation in Pinus species, we optimized protoplast isolation from P. densiflora mesophyll tissues, testing various tissue sources and enzymatic digestion conditions.

Initial experiments using young needles from 2-year-old potted plants demonstrated that splitting the needles yielded approximately 20 times more protoplasts than chopping (Figure 1a) in an enzyme solution that includes 3% (w/v) cellulase and 0.14% (w/v) pectinase, based on the Arabidopsis protocol [15]. Further optimization using different needle sources (Figure 1b) showed that 1-month-old cotyledons from potted seedlings yielded the greatest count of protoplasts (5.0 × 10⁶/g FW), followed by in vitro-grown 1-month-old cotyledons, which produced slightly lower yields (1.2 × 10⁶/g FW). Older needles (2-month-old and 2-year-old) produced significantly lower yields, and outdoor-grown adult needles were nearly undigestible under the same conditions (Figure 1b).

3.2. Enhanced Protoplast Isolation Through Optimized Enzymatic Hydrolysis

To improve protoplast release and minimize potential damage from prolonged enzymatic incubation, six enzyme combinations (E1–E6) and varying hydrolysis durations were evaluated using 1-month-old potted cotyledons (Figure 1c). E1 (3% cellulase and 0.14% pectinase) served as the control, while E2, E3, and E4 introduced hemicellulase (2%) with progressively increasing pectinase concentrations (0.14%, 0.5%, and 0.7%, respectively). E5 increased cellulase (4.5%) and hemicellulase (3%) concentrations by 1.5-fold compared to E4, while E6 further increased cellulase (6%) and hemicellulase (4%) concentrations.

The enzyme combinations significantly influenced protoplast isolation efficiency. E5 (4.5% cellulase, 0.7% pectinase, and 3% hemicellulose) achieved the highest protoplast yield (approximately 1.2 × 10⁷/g FW) within 4–5 h of hydrolysis. This indicates that the optimized balance of cellulase, pectinase, and hemicellulase in E5 effectively breaks down cell wall components, maximizing protoplast release. In contrast, E1 and E2 produced significantly lower yields, demonstrating the critical role of hemicellulase and sufficient pectinase concentrations in improving cell wall digestion. Although E6 utilized higher concentrations of cellulase (6%) and hemicellulase (4%), it resulted in reduced protoplast yield and viability, likely due to enzyme saturation or increased cellular damage associated with excessive enzyme levels.

3.3. Improving Washing Buffer for High-Quality Protoplast Recovery

Following isolation, pine needle protoplasts showed a tendency to form sticky clusters when suspended in the standard W5 washing buffer. This clustering, likely caused by surface adhesion and cellular aggregation during the washing and pelleting steps, significantly reduced protoplast viability (Figure S1).

To address this issue, we tested various modifications to the washing buffer to minimize cell aggregation and improve protoplast quality. Among the buffers tested, a modified W5 buffer (W5-m), supplemented with 5 mM glucose, was the most effective, compared to the standard W5 buffer and other alternatives (W1 and W2) (Figure 1d).

Using the W5-m buffer, protoplast viability significantly improved, reaching 86.2% (Figure 1d and Figure S1). Figure 1e supports these findings by showing protoplasts under bright-field and FDA staining. The bright-field image indicates that protoplasts retain their spherical shape, while the FDA staining confirms their viability through the presence of fluorescent signals. Together, these results validate the effectiveness of W5-m in preserving both the structural integrity and functionality of protoplasts. Additionally, the size range of isolated pine needle protoplasts, 50–80 µm in diameter, aligned with previous studies on mesophyll protoplasts in gymnosperms [6,8,10] (Figure 1e).

3.4. Protoplast Isolation from Woody Tissues of P. densiflora

Building on the successful isolation of protoplasts from mesophyll tissue, we extended the method to woody tissues, specifically developing xylem (DX) and whole stems, using the same enzyme solution. DX protoplasts were successfully isolated, achieving a viability of 85.3% and a yield of 8.9 × 10⁵/g FW, with cell sizes ranging from 5 to 60 µm (Figure S2), following established methods [6,19].

For whole-stem protoplast isolation, young stems from mature P. densiflora trees were utilized. A primary challenge was the abundant resin released from large resin ducts, which impeded enzymatic digestion and introduced substantial debris into the isolation solution [8] (Figure S3). To address this, stem sections were cut under distilled water and subjected to repeated washing with fresh distilled water, effectively reducing resin contamination and enhancing enzyme accessibility, thereby facilitating cleaner protoplast preparations (Figure 2a). Subsequently, a time-course digestion experiment was performed to optimize protoplast release. Protoplasts began to emerge after 2 h, with a marked increase in yield observed at 5 h (7.4 × 10⁴/g FW, viability 68.4%). However, extending digestion beyond 7 h resulted in decreased viability, increased cellular lysis, and the formation of aggregates (Figure 2b). Microscopic analysis confirmed that a 5 h enzymatic digestion provided an optimal balance between protoplast yield and integrity, minimizing damage from prolonged enzyme exposure (Figure 2c).

3.5. Efficient Purification of Stem-Derived Protoplasts Using Sucrose Gradient Separation

Stem-derived protoplasts from P. densiflora presented challenges due to their heterogeneous cell sizes (ca. 5–60 µm) and significant debris, including undigested tissue fragments, resin residues, and other cellular components, which interfered with pelleting and reduced purity (Figure S2b). These issues necessitated an additional purification step to separate intact protoplasts from debris effectively.

To effectively separate intact protoplasts from contaminants, a sucrose-gradient purification method, modified from Jeong et al. (2021) [27], was employed. This technique leverages differences in cell density to selectively isolate viable protoplasts. Among the sucrose concentrations tested (13%, 15%, and 17%), the 17% gradient provided the clearest separation, forming a distinct protoplast layer while minimizing debris contamination (Figure 3 and Figure S4). We did not detect intact, wall-containing cells as contaminants in this fraction. The purified stem-derived protoplasts exhibited significantly improved quality, with minimal aggregation, a yield of 7.4 × 10⁴/g FW, and a viability of 81.9% (Figure 3). The high viability indicates that the sucrose gradient method effectively preserves protoplast integrity while removing contaminants.

3.6. Functional Analysis of Gene Transfection and Transcriptional Activation in Pine Protoplasts

Gene transfection in gymnosperm protoplasts has been notoriously difficult, with conventional methods yielding poor efficiency and inconsistent expression. We optimized a PEG-mediated transfection method for P. densiflora cotyledon protoplasts, achieving a GFP transformation rate of 94.1% with 82.9% viability, comparable to Arabidopsis leaf protoplasts (Figure 4a,b). To assess background fluorescence, protoplasts transfected with an empty vector were used as negative controls (Figure 4a,b). Under identical imaging conditions, these controls exhibited negligible autofluorescence, indicating that the observed GFP signal was not confounded by endogenous fluorescence from resin ducts or phenolic compounds. Arabidopsis protoplast was used solely as a well-characterized reference system to validate assay performance and facilitate comparative context.

To assess the functionality of the purified pine protoplasts, we conducted a transient activation assay (TAA) using effector and reporter constructs derived from Arabidopsis and P. densiflora genes [23,26]. The Arabidopsis MYB46 (AtMYB46) effector (35S::AtMYB46) strongly activated its native ProAtCesA4::GUS reporter in Arabidopsis protoplasts (Figure 4c), confirming its robust activity in its native cellular context. In pine protoplasts, AtMYB46 also induced reporter activation, albeit at a lower level, indicating partial functionality in the gymnosperm system. Conversely, the P. densiflora MYB46 (PdeMYB46) effector (35S::PdeMYB46) strongly activated its native ProPdeCesA7::GUS reporter in pine protoplasts (Figure 4d), demonstrating robust functionality within the gymnosperm system. Notably, PdeMYB46 also activated the reporter in Arabidopsis protoplasts, showing its ability to function in an angiosperm cellular environment.

4. Discussion

4.1. Optimization of Protoplast Isolation from P. densiflora

Developing an efficient protoplast isolation method for Pinus species is crucial due to longstanding challenges in achieving successful gene transfection. Conventional pine protoplast preparation methods are often hindered by low yields, reduced viability, and poor transfection efficiency, significantly limiting their application in functional genomics. In this study, we developed an improved approach to isolate high-yield, viable protoplasts from both mesophyll and woody tissues of P. densiflora.

A major limitation of earlier protocols lies in the structural and biochemical features of mature pine needles, such as thick epidermis, cuticular waxes, and abundant secondary metabolites, that hinder enzymatic digestion and reduce protoplast yield and viability [8,28,29]. We overcame these barriers by longitudinally splitting young cotyledons, which significantly improved enzyme accessibility and enhanced both protoplast yield and transfection efficiency (Figure 1a,b). Notably, this splitting method also produced far more viable protoplasts from hypocotyls than conventional chopping (Figure 1a), likely due to reduced physical damage and better enzyme penetration. While splitting preserves cell integrity by exposing internal surfaces with minimal stress, chopping often ruptures cells and releases phenolics and resins that interfere with digestion. Similar benefits of gentle tissue handling have been observed in Populus [30] and Arabidopsis [15]. Additionally, Gupta and Durzan [8] reported that excessive disruption in conifers exacerbates resin-related issues. Thus, splitting young tissues serves as a critical refinement for high-quality protoplast isolation in woody gymnosperms like P. densiflora.

The choice and concentration of cell wall-degrading enzymes, as well as digestion time, were critical for balancing protoplast yield with cell viability [12,13]. Our optimized enzyme mix (E5) significantly enhanced yield while minimizing cell damage by shortening the hydrolysis duration (Figure 1c). Shorter digestion times also prevented cellular deterioration commonly caused by prolonged enzyme exposure [11,12,13]. Additional improvements were achieved through the inclusion of vacuum infiltration before enzymatic digestion. This step enhances enzyme penetration and accelerates tissue softening, which has proven particularly beneficial for woody tissues [27]. Moreover, we modified the washing buffer (W5-m) by supplementing it with glucose, which improved protoplast viability and reduced aggregation during pelleting (Figure 1d). Glucose likely acts as an osmotic stabilizer, mitigating membrane stress and reducing electrostatic interactions during centrifugation [31].

In the context of gymnosperm systems, the highest protoplast yield reported for Ginkgo biloba is approximately 4 × 10⁶/g FW [32]. By contrast, the method developed here for P. densiflora routinely delivers up to 1.2 × 10⁷/g FW from cotyledon tissue, an approximately three-fold increase, while maintaining cell viability above 85%. These results position our system among the most efficient protoplast isolation methods developed for gymnosperms to date. This optimized method is critical for downstream applications in genetic and molecular studies, providing a reliable foundation for protoplast-based research in P. densiflora. Additionally, given the significant number and high survivability of protoplasts that result from this method, the system is well-suited for downstream applications such as single-cell RNA sequencing, which typically requires 10³–10⁴ viable cells per sample [33].

Woody tissues posed particular challenges for protoplast isolation in P. densiflora. Protoplasts extracted from developing xylem and whole stems exhibited considerable heterogeneity in cell size (5–60 µm) and were frequently contaminated with resin and undigested tissue fragments. These issues underscore resin exudation and extended digestion times as major limitations in conventional pine protoplast protocols, especially when applied to lignified tissues. By optimizing enzymatic digestion time and implementing pre-treatment steps to reduce resin leakage, our method substantially improved the yield and viability of protoplasts from woody tissues, enabling more consistent and reliable downstream applications (Figure 2). Furthermore, the application of a 17% sucrose gradient effectively removed residual debris, selectively enriching intact protoplasts and enhancing the reproducibility of subsequent assays, particularly gene transfection (Figure 3).

In summary, this improved method (outlined in Figure 5) successfully addresses critical obstacles in pine protoplast preparation—including resin interference, excessive digestion, and low purity—offering a robust and scalable platform for molecular analyses in gymnosperm tissues. It is especially advantageous for gene transfection studies, which have long been impeded by suboptimal protoplast quality in Pinus species [12,13].

4.2. Examining Gene Transfection and Transcriptional Activation in Pine Protoplasts

The high transfection efficiency achieved using PEG-mediated transformation in P. densiflora cotyledon protoplasts marks a significant advancement over previous gymnosperm studies, which often reported poor transformation outcomes without providing quantifiable data [6,14]. Our system, demonstrating a GFP transformation efficiency of over 90%, rivals that of well-established angiosperm systems such as Arabidopsis thaliana, highlighting the robustness of the optimized protocol for functional assays.

To evaluate transcriptional activation, we employed transient activation assays (TAAs) using MYB46 effector and CesA reporter constructs from both Arabidopsis and P. densiflora. The Arabidopsis MYB46 effector (35S::AtMYB46) efficiently activated its native ProAtCesA4::GUS reporter in Arabidopsis protoplasts, but its activity was markedly reduced in P. densiflora protoplasts. In contrast, the P. densiflora MYB46 effector (35S::PdeMYB46) robustly activated its native ProPdeCesA7::GUS reporter in both P. densiflora and Arabidopsis protoplasts (Figure 4c,d), suggesting broader cross-species functionality.

Both effector–reporter combinations demonstrated activation in the two protoplast systems, showcasing the versatility of Arabidopsis and pine protoplasts as platforms for studying transcription factor functionality. The difference in activation levels between Arabidopsis and pine protoplasts, however, highlights the influence of cellular context, including differences in transcriptional machinery, co-factors, or promoter recognition between the two species. These results suggest a degree of compatibility between gymnosperm transcription factors and angiosperm regulatory machinery, providing insights into conserved regulatory mechanisms. These findings align with previous studies showing variability in species-specific promoter activity across systems, such as the higher activity of the AtUBQ10 promoter compared to the CaMV 35S promoter in apple protoplasts [34].

Collectively, these results validate P. densiflora protoplasts as a flexible platform for transient gene expression studies. Importantly, given the persistent inefficiency and genotype dependence of stable transformation in gymnosperm species, the development of a reproducible and high-efficiency transient transfection platform represents a critical advancement. This system enables the rapid testing of transcription factors, regulatory elements, and potential genome editing components in a transient context. It also provides an accessible approach for exploring gymnosperm-specific gene regulation, including pathways related to wood formation, stress responses, and secondary metabolism, which are of growing interest in forest biotechnology.

5. Conclusions

This study presents an optimized and reproducible method for isolating and purifying protoplasts from P. densiflora (Figure 5), achieving high yields and viability from both mesophyll and woody tissues. By fine-tuning enzymatic digestion, washing, purification steps, and tissue selection, we effectively addressed key limitations of conventional pine protoplast protocols, including resin interference, low viability, and debris contamination. The resulting protoplasts supported high-efficiency gene transfection and functional assays, demonstrating their reliability for molecular applications in gymnosperms. Notably, cross-species transcriptional activation between Arabidopsis and pine constructs underscores the platform’s utility for studying both conserved and lineage-specific gene regulation. This method provides a reliable system for transient gene expression in conifers, offering a useful platform for gene discovery and regulatory analyses, with potential applications in studies of wood formation, stress adaptation, and forest biotechnology.