The sugarcane variety ROC22 planted in the sugarcane (Saccharum officinarum) experimental base of the College of Agriculture, Guangxi University, Guangxi, PR. China. The young leaves were sampled at the early elongation stage. Collected samples were quickly frozen in liquid nitrogen and stored at -80°C freezer for further use.

Protoplasts were isolated from sugarcane young leaves as previously described (Zhang et al., 2022). Briefly, the robust tail sheaths of sugarcane at the initial elongation stage wereselected as materials for enzymolysis. At first, outer two to three layers of the leaf sheaths were peeled off and then sterilized with 75% alcohol for 30 s. The outer layer and leaf sheaths at both ends were removed after three washes with sterile water to reveal the light-yellow central leaves. The young leaves 1−5 cm above the growing point were cut into thin slices with a thickness of approximately 1 mm. Next, 0.5 g of young leaves were collected and added to 5 mL of CPW solution (containing 13% mannitol, pH 5.8). After plasmolysis for 0.5–1 h, the CPW (containing 13% mannitol) solution was removed, and 5 mL of enzymolysis solution was added to allow enzymolysis at 28°C for 4 h. With 0.5M of mannitol in the digestion buffer, highest protoplast yield and viability was further observed using an enzyme composition containing 3% (m/v) cellulase R-10 and 1.7% (m/v) pectinase R-10. The protoplast suspension was then filtered through 100- and 200-mesh cell sieves subsequently, and the protoplasts were purified using gradient centrifugation (750 rpm/min, 5 min).

Protoplast viability was determined using fluorescein diacetate (FDA): 100 uL of protoplast suspension was aspirated in a 2 mL centrifuge tube, 2 uL of 5 mg/mL FDA was added, mixed, and incubated at room temperature for 5 min. Green protoplasts were observed and photographed under blue excitation light using a confocal laser microscope (Nikon, Beijing, China). The green- fluorescent active protoplasts were observed and photographed under blue excitation light using a confocal laser microscope (Nikon, Beijing, China). The FDA stock solution was dissolved in acetone to 5 g/L and stored at -20°C. The FDA working solution was made by diluting the above stock solution with phosphate buffer solution (PBS) to 5 mg/mL. The total number of cells in the samples measured in this study was 2.5 × 10⁶ g/FW.

The enzymatically digested protoplasts were put into 2.5% glutaraldehyde fixative (pH 7.2) at 4°C and fixed for more than 2 h; washed 3 times with 0.1 mol/L PBS (pH 7.2) for 10 min each; fixed for 3 h after 1% starvation acid; washed again 3 times with PBS for 10 min each; graded with 30%, 50%, 70%, 90% and 100% alcohol After dehydration, the samples were dried in a Leica (EM CPD300, Weztlar, GER) automatic critical point drier. The dried samples were fixed on the sample tray with conductive double-sided adhesive and dried under vacuum (1 × 10 ^-6 kPa) for 0.5 h to increase the adhesion of the sample to the conductive adhesive. The sample is gently held with forceps and pulled upward, and the pulled off tissue is partly adhered to the conductive adhesive and the other part is made to adhere to the conductive adhesive under the dissecting microscope with the fracture side up. After the samples were pasted, the samples were sprayed with gold in a Leica (EM ACE600, Weztlar, GER) ion sputter coater with a coating thickness of 22 nm, and the samples were observed and photographed using a Hitachi (S-3400N, Tokyo, JPN) scanning electron microscope (Lu et al., 2021).

Chromosomes were examined in a dark room. First, 30 µL of DAPI (0.5 µg/mL) drops per sheet were applied to the microscopically examined chromosome films, covered with coverslips, and observed using a fluorescence microscope (Leica DMRA2 Microscope, Weztlar,GER) and the images were captured using the SH DC 350F camera system (Leica QFI, Weztlar, GER). The average pixel values of chromosomes were measured using Leica (CW4000, Weztlar, GER).

The frozen sectioning method combined with indirect immunofluorescence technique and DAPI staining was used to observe the arrangement of the cellular microtubule array of sugarcane young leaves and protoplasts using fluorescence microscopy, as previously described Li et al. (2008).

Young leaves: The young leaves of ROC22 sugarcane were sampled at the early elongation stage, flash-frozen in liquid nitrogen, and stored at -80°C for later use. Three replicates were produced. Protoplasts: Following enzymolysis of ROC22 sugarcane young leaf cells, the density of the protoplasts was adjusted to 1×10⁶/mL. Next, 0.5 mL of the protoplast sample was taken, shaken well, frozen in liquid nitrogen, and stored at -80°C. Three replicates were produced.

The experiment was conducted based on previous methods of Wang et al. (2022). Briefly, 0.5 g of sugarcane young leaves and 5×10⁶ protoplasts were homogenized in 2 mL of lysis buffer containing 8 M urea, 50 mM Tris-HCl (pH 8), and 0.2% sodium dodecyl sulfate (SDS), followed by sonication on ice for 5 min. The samples were then centrifuged at 12,000 × g at 4°C for 15 min, and the supernatant was transferred to a new tube. The protein concentration was determined using Bradford protein assay (Shanghai yuanye Bio-Technology Co., Ltd, Shanghai, China). Extracts from each sample were reduced with 2 mM Dithiothreitol (DTT) for 1 min and alkylated with sufficient iodoacetic acetate for 1 min in the dark at 28°C. Next, samples were mixed with 4-fold volume of precooled acetone, incubated at 20°C for 1 min, and centrifuged. The precipitate was collected and washed three times with cold acetone. The pellets were dissolved in lysis buffer containing 0.1 M triethyl ammonium bicarbonate (pH 8.5) and 8 M urea. An equal number of proteins were digested with trypsin (Promega, Madison, WI, USA) at a ratio of 1:50 (w:w) for 16 min at 37°C. The digested proteins (100 µg) were labeled using the iTRAQ reagent kit (Applied Biosystems, Framingham, MA, USA) according to the manufacturer’s protocol.

A Shimadzu LC system equipped with a 20 AB column (Gemini C18 4.6 × 250 mm, 5 µm) was used for high-performance liquid chromatography analysis. LC-MS/MS assay was conducted using the UltiMate 3,000 UHPLC system (Thermo, Massachusetts, USA). Protein quantification was performed as previously described (Xu et al., 2022).

Total RNA extraction and reverse transcription of RNA into cDNA were conducted using Prime Script TMII 1stStrand cDNA Synthesis Kit and TB GreenTM Premix Ex TaqTMII (TaKaRa, Beijing, China) according to the manufacturer’s protocol. RT-qPCR was performed with ChamQ Universal SYBR qPCR Master Mix (Vazyme, Nanjing, China) and the QuantStudio 5 Real-time PCR system (Applied Biosystems, Waltham, MA, USA) (Xu et al., 2022). The amplification procedure was as follows: 95°C 30 s, 1 cycle; quantitative analysis: 95°C 5 s, 60°C 30 s, 40 cycles; melt curve: 95°C 5 s, 60°C 1 min, 95°C, 1 cycle; cooling: 95°C 30 s, 1 cycle (Zhang et al., 2022). Primer sequence is shown in Supplementary Table S6 .

The data were subjected to ANOVA using the SPSS statisticalsoftware SPSS (Chicago, IL,USA). According to Duncan’s Multiple Range Test, the mean values were statistically compared and separated at p < 0.05. After calculating the difference between the obtained. After obtaining the variance analysis results and making a graph with Origin 2021 (OriginLab USA).Bowtie2 software was used to compare clean reads of each sample to Unigene, while RSEM was used to calculate the gene expression level of each sample.

Functional analysis of identified proteins was performed using Gene Ontology (GO) (http://www.geneontology.org). Differentially accumulated proteins were then inputted into the Kyoto Encyclopedia of Genes and Genomes (KEGG) database (http://www.kegg.jp/kegg/pathway.html). To determine functional subgroups and metabolic pathways in which the differentially accumulated proteins were enriched, GO and KEGG pathway enrichment analyses were conducted. Cluster analysis of the differentially accumulated proteins was conducted using Cluster 3.0, and a heatmap was generated using TreeView version 1.6.

The contents of MDA and O2- were determined using a kit (Nanjing Jiancheng Biotechnology Co., LTD, Nanjing, China). Enzyme activity was measured using the conventional method. Peroxidase (POD) activity was determined using the guaiacol method (Pu et al., 2021), whereas ascorbic acid peroxidase (APX) and (catalase) CAT activities were measured using the UV absorption method (Sun et al., 2008; Zhang and Chen, 2010). SOD activity was measured in pure enzyme preparations using the Francexo Paoletti method, which uses a stable reagent for rapid and highly sensitive measurement (Paoletti et al., 1986).

The sugarcane young leaf cells ( Figure 1A ) were rounded after enzymatic digestion and showed smooth cell membranes ( Figures 1B, F ). The FDA detection results showed that the viability of the enzymatically digested sugarcane protoplasts of > 90% (p < 0.05) ( Figure 1C ). The cell membranes of sugarcane young leaves were intact before enzymolysis ( Figure 1D ), but the cell membranes of protoplasts were perforated to different degrees post enzymolysis ( Figure 1E ). DAPI fluorescence staining showed that the nucleolus of the nuclear membrane was clear; the DAPI fluorescence was strong prior to enzymolysis ( Figures 1G, H ). The nuclear membrane of protoplasts remained intact following enzymatic hydrolysis, while the blue fluorescence (DAPI) and nuclear activity were weakened, and the nucleolus structure was not obvious ( Figure 1I ). There was a connection between the microtubules and the plasma membrane in sugarcane young leaf cells, and the microtubule array was well organized ( Figures 1J, K ), However, the microtubules depolymerized and form many short rod-like structures in the protoplasts, and a large number of adhesions of periplasmic microtubules were observed on the plasma membrane of newly isolated protoplasts in a radial or fan-shaped distribution ( Figure 1L ).

After enzymolysis, malondialdehyde (MDA) content in the protoplasts was significantly increased, 4.7 times higher than that in the young leaves ( Figure 2A ). In contrast, the content of O^2- in protoplasts was significantly lower, only 1.2% of that in the young leaves ( Figure 2B ). Compared with sugarcane young leaves, SOD content decreased to 93.7% post enzymolysis ( Figure 2C ). However, compared with that in sugarcane young leaves, the content of Peroxidase POD, Catalase CAT, and Ascorbate peroxidase APX in the protoplasts was significantly reduced to 17.7%, 6.5%, and 17.5%, respectively ( Figures 2D–F ).

Following enzymolysis, Gu/ZnSOD and CAT expression levels in the protoplasts were downregulated compared with those in the young leaves, which accounted for only 1.6% and 2.8% of that in the young leaves, respectively ( Figure 3 ).

The expression of the DREB, WRKY, MAPK4, and NAC genes in sugarcane young leaves and protoplasts following enzymolysis was detected using qPCR, with GADPH as an internal reference gene. DREB, WRKY, MAPK4, and NAC gene expression in sugarcane protoplasts was significantly upregulated, being 21-, 57,184-, 6,100-, and 200,050-fold higher than that in the young leaves, respectively ( Figures 4A–D ).

Proteins with a fold change ≥ 2 and a p < 0.05 were considered significantly differentially expressed proteins (DEPs). A total of 2,287 DEPs in sugarcane protoplasts were identified through statistical analysis of sugarcane young leaves and protoplasts following enzymatic digestion. Of these DEPs, 810 were upregulated and 1,477 were downregulated ( Figure 5A ). In addition, Figure 5B shows that six samples were large, and the experimental data were effective and reasonable. In addition, the results in Figure 5B . shows that the correlation coefficient between the six samples is close to 1, indicating a high degree of similarity between the samples and a low number of differential proteins between the samples. This suggests that the experimental data are valid and reasonable.

Transcriptome studies before and after enzymatic digestion of sugarcane young leaves showed that the expression levels of genes closely associated with cell cycle (CyclinD3, CyclinA, CyclinB, and cdc2), cell proliferation (phytosulfokine gene, PSK), cell wall regeneration (Galacturonosyltransferase gene, GAUT; cellulose synthase gene, CESA), which are closely linked to cell wall regeneration, showed significant changes in the expression levels. As shown in Figure 8 , the expression of Cyclin D3, Cyclin A, Cyclin B, and cdc2 in sugarcane protoplasts was only 52%, 21.32%, 18.60%, and 45% of the young leaves, respectively; the expression of PSK in sugarcane protoplasts was significantly lower, accounting for only 2% of the young leaves; the expression of GAUT and CESA was only 65% and 47% of the young leaves, respectively ( Figure 8 ).

The degree of viability is used as a criterion to determine the quality of protoplasts after enzymatic digestion. However, enzymatic structural variability, which results in anomalies in digestion during cell division and in plants regenerated from protoplasts (Cambecedes et al., 1988). In this study, although high-yield (5×10⁶ protoplasts/g FW) and high-vitality (> 90%) protoplasts were obtained by optimizing the enzymatic hydrolysis conditions, the cell membranes of the protoplasts perforated to different degrees post enzymolysis. The nucleolus was intact following enzymatic hydrolysis, but the blue fluorescence (DAPI) and nuclear activity were weakened. Prior to enzymolysis, the microtubules were tightly connected to the plasma membrane in young sugarcane cells, and a large number of periplasmic microtubules stuck to the plasma membrane of all newly isolated protoplasts in a fan-like pattern. These anomalies often hinder the introduction of new plant varieties obtained via in vitro protoplast fusion (Handley et al., 1986). The highly viable sugarcane protoplasts obtained via enzymatic digestion of sugarcane young leaves using the optimal mannitol concentration showed severe browning at a later stage, and cells could not continuously divide, which greatly hindered the regeneration of sugarcane protoplasts (Li et al., 2020). Investigation of the tubulin cytoskeleton of protoplasts isolated from Medicago sativa and Nicotiana tabacum indicates that the perinuclear and radial cytoskeleton significantly limit the capacity for proper cell division. These factors play a key role in the migration of the nucleus to the center of the cell and the maintenance of the proper position of the nucleus just before division (Meijer and Simmonds, 1988). Therefore, enzymatic digestion influences the subcellular structure and microtubule array of sugarcane protoplasts and may be the cytological reason for the difficulty of highly viable sugarcane protoplasts to regenerate.

Higher or lower osmotic pressure can cause osmotic stress in protoplasts, thereby reducing protoplast viability. Therefore, the enzymatic process requires the addition of mannitol and sucrose to regulate cellular osmolality, (Bai et al., 2020). Osmotic stress increases ROS and oxidative stress (Noctor et al., 2015). Our results showed that in sugarcane protoplasts, MDA content significantly increased by 4.7 times that in the young leaves, whereas antioxidant anion O^2- content significantly decreased to only 1.2% that in the young leaves. Moreover, sugarcane protoplasts were subjected to oxidative stress, which activated the expression of the resistance gene MAPK and resulted in significant changes in ROS and high levels of accumulated MDA, and is one of the primary causes of protoplast browning, reduced cell viability, and death (Apel and Hirt, 2004). In addition, SOD, CAT, POD, and APX reduce oxidative stress in maize (Zea mays) (Ma et al., 2015), reduce membrane damage during enzymolysis in peanut (Arachis hypogaea) protoplasts (He et al., 1994), and scavenge free radicals in tomato (Solanum lycopersicum) (Bai et al., 2020). Our study showed that the activities of the antioxidant enzymes POD, CAT, and APX decreased to 17.7%, 6.5%, and 17.5%, respectively, in sugarcane protoplasts. In addition, the expression levels of Gu/ZnSOD and CAT in sugarcane protoplasts were only 1.6% and 2.8%, respectively, lower than those in the young leaves. During protoplast isolation, cell wall removal and the consequent abiotic stress decrease the level and activity of antioxidant enzymes, thus disrupting the dynamic balance between intracellular ROS production and scavenging, which in turn leads to protoplast browning (Chen et al., 2006).

Osmotic stress has multiple effects on cell physiology and protein and gene expression. Osmotic pressure reduces cell volume, thereby elevating the concentrations of ions and macromolecules. This indicates that several multivalent proteins and genes remain dispersed at the physiological conditions and reversibly condense to microscopic granules during enzymolysis (Majumder and Jain, 2020). Enzymatic removal of the cell wall not only causes osmotic stress on the protoplasts (Wang et al., 2022) but also inevitably alters the expression of NAC secondary wall thickening promoter (NST)/secondary wall-associated NAC structural domain protein (SND) and SOMBRERO (SMB) subfamily proteins, thus hindering the ability of NST, SND, and SMB to participate in the formation of the secondary cell wall (Kim et al., 2021). In this present study, the expression levels of the stress-resistant genes DREB, WRKY, MAPK4, and NAC were significantly upregulated. In addition, stimulation of DREB resistance gene expression and enhancement of the DREB/CBF-COR pathway improved plant tolerance to various abiotic stresses. Omotic stress during protoplast separation altered the expression of several resistance genes, leading to browning and difficulty in the regeneration of protoplasts (Hayat et al., 2022)

Plant protoplasts constitute unique single-cell systems that can be subjected to genomic, proteomic, and metabolomic analyses (Xu et al., 2021). Proteomic studies have shown that proteins such as ascorbate peroxidase (Holmes et al., 2006), dehydroascorbate reductase, glutathione transferase, and mitochondrial manganese superoxide dismutase (Shi et al., 2008) are associated with high cytokinesis activity. Using iTRAQ proteomic strategies coupled with LC-MS/MS, Wang et al. (2017)examined global changes in the proteome following protoplast development and identified 162 proteins involved in defense responses, energy production, translation, metabolism, protein destination and storage, transport, transcription, cell growth/division, cell structure, and signal transduction. Zhao et al. (2019) used a label-free quantitative proteomic approach to determine the protein accumulation profiles of protoplasts and chloroplasts under infection with Rice stripe virus (RSV) and established a method to elucidate the change in the localization of nucleus-encoded ChRPs. Our proteomic analysis identified 2,287 DEPs following the enzymatic digestion of sugarcane young leaves, of which 810 were upregulated and 1,477 were downregulated. The main biological processes of the DEPs included cellular processes (1,101 proteins), metabolic processes (1,049 proteins), stimulus responses (320 proteins), bioregulation (267 proteins), and cellular component organization or biogenesis (259 proteins). Abiotic stresses generated by enzymatic processes in sugarcane young leaves can cause continuous trauma to protoplasts. Since plants are constantly threatened by wounding throughout their lives, understanding the biological responses to wounds at the cellular level is critical (Son et al., 2021). These proteins are part of a dynamic networks that change in response to enzymatic digestion.

Oxidative stress affects gene expression in addition to cellular physiological and biochemical metabolism. Knockout or knockdown of SlMAPK3 expression inhibits the activities of antioxidant enzymes (APX, POD, SOD, and CAT) and induce the accumulation of H₂O₂ (Shu et al., 2022). However, osmotic stress caused by enzymatic hydrolysis causes an imbalance in the production and elimination of ROS in sugarcane young leaves, resulting in the accumulation of ROS, the main reason for the browning and death of protoplasts (Khatri and Rathore, 2022). Peroxisome proliferation and CAT activity contribute to ROS homeostasis and the subsequent induction of protoplast division (Tiew et al., 2015). In addition, the overexpression of TaWRKY46 in wheat has been reported to increase the activities of SOD, CAT, and POD in osmotic balance regulation and ROS scavenging (Yu and Zhang, 2021). Our current study showed that sugarcane heterozygous cells were subjected to oxidative stress during enzymatic digestion, during which most oxidase-related proteins and genes were downregulated. The expression of Gu/ZnSOD and CAT, as well as DEPs such as ascorbate peroxidase, glutathione peroxidase, peroxidase, and catalase, was significantly downregulated ( Figure 5 ). These results showed that protoplasts could not overcome oxidative stress after enzymolysis.

Enzymatic hydrolysis of sugarcane young leaves reduces the expression of genes related to protoplast regeneration, which is also a major obstacle to protoplast regeneration. In this study, the transcriptional levels of CycD2 and CDC2 (two genes regulating cell cycle progression) decreased, thereby inhibiting the activity of vacuole invertase after heat stress recoveryand shortening the cell length (Luo et al., 2021). In addition, CyclinD3, CyclinA, CyclinB, and CyclinE regulate the cell cycle during cell proliferation (Ahn et al., 2018). Our study also showed that the expression levels of CyclinD3, CyclinA, CyclinB, cdc2, PSK, CESA, and GAUT, which are genes related to plant regeneration, were significantly downregulated following enzymatic hydrolysis, reaching only 52, 21.32, 18.60, 45, 2, 47, and 65% of those in the young leaves, respectively. Phytosulokine (PSK) is a plant hormone involved in transmitting information between plant cells, and a decline in its expression inevitably affects plant development and growth (de Souza et al., 2021). Seed plants use different CESA isoforms for primary and secondary cell wall deposition (Li et al., 2022). The GAUT gene family may affect fiber development, including elongation and fiber thickening, in cotton (Senmiao et al., 2021). Compared with that in sugarcane young leaves, the expression of these key genes changed significantly following enzyme digestion. Therefore, the degradation of somatic cells by enzymes affects protoplast regeneration.

In conclusion, our study shows that enzymatic hydrolysis induces osmotic and oxidative stress in sugarcane protoplasts, which in turn alters the expression of proteins involved in bioenergetic metabolism, cell wall synthesis, and cell cycle regulation); reduces the activities of stress-related enzymes (SOD, POD, CAT, and APX), thereby hindering the elimination of oxidation products (O^2-), elevates the physiological indexes of MDA, and alters the expression of genes related to protoplast regeneration (GAUT, CESA, CyclinA, CyclinB, CyclinD3, cdc2, and PSK) compared with those in sugarcane young leaves ( Figure 9 ).

In the study of plant molecular biology processes, the establishment of a suitable protocol for protoplast transformation will enable a detailed analysis of early signs of protoplast regeneration (e.g., chloroplast division and cell wall reconstruction) and expand the prospects for functional studies of plants (Neubauer et al., 2022). Fluorescent dye labeling and qPCR can also be used to examine the effect of abiotic stress on the expression of protoplast-related genes after enzymatic digestion (Pasternak et al., 2005). This present study showed that enzymatic digestion caused osmotic stress in sugarcane protoplasts resulted in significantly up-regulated expression of related resistance genes and significantly down-regulated expression of regenerated genes. Thus, future research can use the protoplast transient gene expression system to locate the location and function of differential proteins during enzymatic digestion. This may, to some extent, elucidate the mechanism underlying how changes in protein expression during enzymatic digestion (such as changes in antioxidant enzyme activity) hinder protoplast regeneration. In addition, enzymatic digestion affects osmotic stress resistance as well as oxidative stress-related and regenerative genes. Therefore, it is possible to establish molecular markers for the enzymatic digestion of protoplasts, calibrate the degree of enzymatic digestion, and screen the conditions of enzymatic digestion (selection of optimal materials, enzymatic solution composition, osmotic pressure, time, and concentration) to obtain protoplasts with high yield and quality ( Figure 10 ). The findings of this study providenovel insights into the molecular, physiological, and cytological mechanisms hindering the regeneration of sugarcane protoplasts. The study present the relevant parameters for establishing a standard system for regenerated protoplasts using molecular and antibody markers for enzymolysis detection.