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. 2024 Jul 31;72(32):18257–18270. doi: 10.1021/acs.jafc.4c03614

Comparative Transcriptomic and Lipidomic Analysis of Fatty Acid Accumulation in Three Camellia oleifera Varieties During Seed Maturing

Dayu Yang §, Rui Wang †,‡, Hanggui Lai §, Yimin He §, Yongzhong Chen †,‡, Chengfeng Xun †,‡, Ying Zhang †,‡, Zhilong He †,‡,*
PMCID: PMC11328181  PMID: 39084609

Abstract

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Camellia oleifera, a major woody oil crop in China, produces tea oil rich in unsaturated fatty acids, earning it names like liquid gold and eastern olive oil. This study provides an integrated investigation of the transcriptome and lipidome within seeds at the maturing process across three C. oleifera varieties, revealing a significant relationship between fatty acid production and genes involved in lipid synthesis. Through transcriptomic analysis, 26,344 genes with varied expression were found. Functional enrichment analysis highlighted that pathways related to starch and sucrose metabolism, plant hormone signal transduction, and lipid accumulation were highly enriched among the differentially expressed genes. Coordinated high expression of key genes (ACCase, KAS I, KAS II, KAS III, KAR, HAD, EAR, SAD, LPAAT, LACS, DGAT, PDAT) during the late maturation stage contributes largely to high oil content. Additionally, expression variations of SAD and FADs among different varieties were explored. The analysis suggests that high expression of genes such as FAD3, FAD7, and FAD8 notably increased linolenic acid content. This research provides new insights into the molecular mechanisms of oil biosynthesis in C. oleifera, offering valuable references for improving yield and quality.

Keywords: Camellia oleifera, fatty acids, oil biosynthesis, transcriptome, lipidome

1. Introduction

Camellia oleifera Abel., a tiny tree or shrub belonging to the Theaceae family, is a key source of woody oil in Asia, widely grown in countries such as China, the Philippines, Thailand, Japan, and South Korea.1 The C. oleifera seeds, known for their rich oil content, produce tea oil highly valued for its superior quality, significantly contributing to the economy. The fatty acids (FA) in tea oil predominantly consist of unsaturated types, essential for their high nutritional value.2 Oleic acid, a monounsaturated FA, potentially plays a positive role in treating inflammation.3 Additionally, linoleic and linolenic acids in tea oil are crucial for lowering blood pressure.4 Tea oil is abundant in bioactive elements like squalene, phytosterols, polyphenols, and tocopherols.5 Byproducts like Camellia seed cake, saponins, and fruit husks are utilized across various industries, including papermaking, chemical fiber, textiles, and pesticides.6 The cultivation of C. oleifera not only holds high economic value but also has a crucial role in soil conservation, and biodiversity maintenance,7,8 thus promoting ecological stability and sustainable development.

FA are vital components of plant metabolism, found in various plant cell lipids. Common FA in plants include palmitic acid (C16:0), stearic acid (C18:0), oleic acid (C18:1), linoleic acid (C18:2), and linolenic acid (C18:3). They are crucial for biological membranes, energy storage, and as precursors for bioactive molecules, playing key roles in plant growth and development.9 Additionally, FA are raw materials for cutin, suberin, jasmonic acid, phenolic lipids, chloroplastic lipids, and sphingolipids, contributing to plant stress defense.10 Oil biosynthesis in plant seeds primarily includes the de novo synthesis of FA in plastids and the synthesis of triacylglycerol (TAG) in the endoplasmic reticulum.11 Acetyl-CoA carboxylase (ACCase), the initial enzyme in de novo FA synthesis, converts acetyl-CoA to malonyl-CoA and is widely regarded as the key rate-limiting step in this process.12 After Malonyl-CoA: acyl carrier protein transacylase (MCAT) converts malonyl-CoA to malonyl-ACP, malonyl-ACP can further undergo a series of reactions catalyzed by fatty acid synthase, including condensation, reduction, dehydration, and rereduction, thereby increasing the chain length. After seven cycles, palmitoyl-ACP is formed. Ketoacyl-ACP synthase II (KASII) further converts C16:0-ACP to stearoyl-ACP.13 Additionally, saturated acyl-ACP in plastids can be converted to unsaturated acyl-ACP under the desaturation effect of enzymes such as C18:0-ACP desaturase (SAD), fatty acid desaturase 6 (FAD6), fatty acid desaturase 7 (FAD7), and fatty acid desaturase 8 (FAD8).14 Fatty acyl-ACP thioesterase (FAT) is responsible for hydrolyzing acyl-ACP, ending the de novo synthesis of FA, and releasing free fatty acids (FFA). Different types of FAT have specific substrate selectivity. FAT A (FATA) mainly hydrolyzes unsaturated long-chain acyl-ACP, while FAT B (FATB) tends to hydrolyze saturated long-chain acyl-ACP.15 The released FFA are converted to acyl-CoA by long-chain acyl-CoA synthetase (LACS) and participate in the assembly of TAG in the endoplasmic reticulum.16 Glycerol-3-phosphate acyltransferase (GPAT), 1-acylglycerol-3-phosphate acyltransferase (LPAAT), phosphatidic acid phosphatase (PAP), and diacylglycerol acyltransferase (DGAT) catalyze four enzyme-catalyzed reactions in the Kennedy pathway, which produces TAG from glycerol 3-phosphate and acyl-CoAs.17 In addition to TAG synthesis through the Kennedy pathway, recent discoveries have revealed more complex pathways centered around phosphatidylcholine.18,19

The expression of genes related to FA synthesis affects not only plant oil production but also various aspects of seed growth and development. The accD gene encodes the β-carboxyl transferase subunit of ACCase. Knocking out the accD gene in tobacco reduces ACCase activity, leading to decreased TAG synthesis, impaired chloroplast division, and reduced fertility.20 Conversely, overexpressing the LuAccD gene in Arabidopsis increases total fatty acid content in seeds and enhances seed germination under salt and mannitol stress.21 Similarly, the absence of KASI in Arabidopsis inhibits chloroplast division and interrupts embryo development.22 Overexpression of SiKASI in Arabidopsis causes male sterility, characterized by defects in microspore and pollen wall development.23 Additionally, differential expression of FAD genes affects fatty acid composition and is associated with plant tolerance to abiotic stresses.24−27

In recent years, rising demand for healthy foods and premium edible oils has spotlighted the FA composition and content in C. oleifera seeds as a key research area. Consequently, unraveling the biosynthetic mechanisms of C. oleifera FA holds crucial practical value for enhancing tea oil quality to meet market demands. Advancements in omics technologies have spurred numerous studies into the biosynthesis and metabolism of FA in C. oleifera seeds. Zhang et al.28 revealed the developmental differences of C. oleifera seeds between different varieties through comparative transcriptome analysis of fruits from two C. oleifera varieties. Gong et al.29 found that the WRINKLED1 (WRI1) transcription factor (TF) interacts with 17 key genes for lipid biosynthesis. Lin et al.30 proposed that in the later stages of seed development, high SAD activity and low FAD2 activity may contribute to an increase in C18:1 levels. Yang et al.31 functionally characterized genes such as SAD, FAD2, FAD3, DGAT1, and DGAT2. Such research is pivotal for elucidating C. oleifera FA composition and metabolic pathways, thereby optimizing tea oil quality and boosting yields.

However, considering the complex polyploidy and diverse varieties of C. oleifera, extensive research is still needed to understand the biosynthesis and accumulation patterns of tea oil. It is noteworthy that during oil synthesis in plants, FA exist as free and bound forms. Previous studies have paid little attention to the dynamic changes of FFA, which are intermediates between de novo FA synthesis and TAG assembly. Observing their variations can help explore the relationship between oil accumulation and gene expression. Here, we conducted a combined transcriptome and lipidome analysis of mature seeds from three C. oleifera varieties to uncover the reasons for differences in oil content and fatty acid composition. This study enhances our understanding of the biosynthesis mechanism of tea oil and provides valuable references for breeding high-oil and high-quality C. oleifera varieties.

2. Materials and Methods

2.1. Plant Materials

The National Engineering Research Center for Oil-Tea Camellia (Changsha, China), which is located at latitude 28°06′ N and longitude 113°01′ E, provided the experimental materials for this study. The base has an altitude range of 80–100 m and is situated in an area with four different seasons, typical of a subtropical monsoon climate. Three eight-year-old C. oleifera varieties were selected in this study, namely Xianglin 192 (XL192), Xianglin 108 (XL108), and Huaihua 2 (HH2). Base on previous research,32 these three varieties show significant differences in oil content and fatty acid composition, making them ideal materials for exploring the mechanisms of oil synthesis. For clear and concise descriptions in the text, they will be referred to as L (XL192), M (XL108), and H (HH2), respectively. The peak oil accumulation period in C. oleifera fruits mainly occurs from mid-September to late October before harvest.33 Therefore, in this study, starting from September 14, 2022, fruits from Camellia trees were sampled every 14 days. The specific sampling dates were September 14 (T1), September 28 (T2), October 12 (T3) and October 26 (T4). Six fruits were selected from different positions of the tree each time, of which three fruits with a full appearance and uniform growth were selected as biological replicates. These samples were moved to the laboratory and stored at −80 °C after being instantly frozen in liquid nitrogen.

2.2. Measurement of Seed Oil Content

After removing the shells from the C. oleifera seeds, the kernels were dried in an oven at 60 °C until they reached a consistent weight. After drying, the kernels were processed in a grinder into a powder. A Soxhlet extractor was used to hold a filter paper bag containing 5 g of the powder (m0), which had been precisely weighed. Petroleum ether was poured into two-thirds of the oil cup after its weight (m1) was measured. The water bath temperature was set to 70 °C, and the sample was reflux extracted for 6 h. The weight of the oil cup (m2) was measured once the petroleum ether had evaporated to a consistent weight in the receiving bottle. The following formula was used to determine the amount of seed oil present: seed oil content (%) = (m2 – m1)/m0 × 100. In this case, m0 represents the sample weight (g), m1 the oil cup weight (g), and m2 the combined weight of the oil and oil cup (g).

2.3. Measurement of Fatty Acid Content Proportion

The FA composition was determined using a gas chromatograph, with each sample containing three biological replicates. A potassium hydroxide-methanol mixed solution (1.31 g potassium hydroxide, 10 mL methanol) was prepared. Four milliliters of n-heptane and 60 μL of oil were combined in a centrifuge tube and carefully stirred. Then, 200 mL of a potassium hydroxide-methanol solution were added, agitated for 30 s, and allowed to stand for approximately 30 min. One gram of sodium bisulfate was added to the centrifuge tube, shaken quickly, and allowed to settle for 20 min after standing. After the solution was clarified, the supernatant was drawn up with a syringe and added to the detection bottle. The detection bottles were loaded into the gas chromatograph in batches, and the FA content was quantitatively determined by the peak area of the chromatogram. The proportion of a certain FA was determined by comparing the peak area of the FA being analyzed to the total peak areas of all fatty acid components.

The determination parameters of the gas chromatograph are as follows: The flame ionization detector is set at 250 °C; the temperature of the sample injection port is 250 °C; the dimensions of the chromatographic column are 60 cm × 0.25 mm × 0.2 mm; the carrier gas is nitrogen; the split ratio is 1:50; the sample injection volume is 1 mL; the heating program is set to 50 °C (held for 2 min), 170 °C (rise by 10 °C/min, held for 10 min), 180 °C (rise by 2 °C/min, held for 10 min), and 220 °C (rise by 4 °C/min, held for 22 min).

2.4. Measurement of Free Fatty Acid Content

After thawing the seed samples, mix 0.05 g with 150 μL methanol, 200 μL methyl tert-butyl ether, and 50 μL 36% phosphoric acid/water (which has been chilled to −20 °C). Vortex the mixture vigorously for 3 min at a speed of 2500 rpm (r/min), followed by centrifugation at 12,000 rpm for 5 min at a temperature of 4 °C. Transfer 200 μL of the clear liquid (supernatant) into a fresh centrifuge tube, use a nitrogen blower to dry it completely, and then introduce 300 μL of a 15% boron trifluoride methanol solution. Vortex the mixture for 3 min at a speed of 2500 rpm, and subsequently incubate it in an oven set at 60 °C for 30 min. Once the mixture has returned to ambient temperature, carefully add 500 μL of n-hexane and 200 μL of a saturated sodium chloride solution. After mixing, the mixture underwent centrifugation at 12,000 rpm for 5 min at a temperature of 4 °C. Subsequently, 100 μL of the upper n-hexane phase were collected for subsequent analysis via gas chromatography–mass spectrometry (GC–MS).

Derivatized samples were analyzed using a GC-EI-MS/MS system (GC, Agilent 7890B; MS, 7000D system). GC conditions: column, DB-5MS capillary column (30 m × 0.25 mm × 0.25 μm, Agilent); carrier gas, high purity helium (purity >99.999%); The heating procedure was started at 40 °C (2 min), 30 °C/min increased to 200 °C (1 min), 10 °C/min increased to 240 °C (1 min), 5 °C/min increased to 285 °C (3 min); traffic: 1.0 mL/min; inlet temperature: 230 °C; injection volume: 1.0 μL. EI–MS/MS settings: Agilent 7890B-7000D GC–MS/MS system, temperature, 230 °C; ionization voltage, 70 eV; transmission line temperature, 240 °C; four-stage rod temperature, 150 °C; solvent delay, 4 min; scanning mode, SIM.

2.5. Analysis of RNA-Seq and Bioinformatics Exploration

TRIzol reagent (Invitrogen, CA, USA) was used to extract total RNA from 36 seed samples of three C. oleifera cultivars at four developmental phases. The purity of RNA was evaluated utilizing a NanoDrop spectrophotometer (Thermo Scientific, DE, USA), while its integrity was verified through an Agilent 2100 (Agilent Technologies, CA, USA). Finally, RNA degradation was assessed by conducting agarose gel electrophoresis (1%). RNA sequencing was performed by Beijing Allwegene Technology Company Limited (Beijing, China).

For each sample, 1.5 μg of RNA was utilized to enrich for mRNA using Oligo (dT) beads. 36 sequencing libraries were obtained using the NEBNext UltraTM RNA Library Prep Kit for Illumina (New England Biolabs, MA, USA), followed by PE 150 sequencing on the Illumina Novaseq 6000. Clean reads were generated by filtering out sequences from the raw data that contained sequencing adapters, had an N (unknown bases) ratio exceeding 10%, or were of low quality. The filtered clean reads were subsequently mapped to the C. oleifera reference genome34 using STAR (v2.5.2b). Genes were considered differentially expressed if they had a corrected P-value of less than 0.005 and an absolute log2 (fold change) greater than 1.

2.6. Weighted Gene Correlation Network Analysis

To study gene coexpression patterns, this study employed the weighted gene correlation network analysis (WGCNA) R package (version 1.72–5). The network creation and module identification were accomplished using default settings, with specific parameters as follows: an unsigned topological overlap matrix was used for network construction; a soft threshold β (power β) was set to 8 to enhance the interconnectivity between modules; each module contained at least 100 genes; the branch merge cut height for module merging was set at 0.25 (i.e., merging coexpression modules with at least 75% similarity). Module eigengene (ME), representing the first principal component of all gene expression data in a module, reflected the overall expression pattern of that module and was used to explore associations with oil amounts. Module membership (MM) was identified by matching a gene’s expression profile with the ME of its appropriate module and was significantly linked to the module’s internal connection (K.in). Utilizing the MM and K.in, Cytoscape (Version 3.10.1) was used to construct regulatory networks of genes within the module.

2.7. Real-Time Quantitative PCR Verification

Based on previous studies, we selected 12 structural genes and TFs related to oil synthesis for RT-qPCR experiments to validate the sequencing results.30,35 The tubulin gene was employed as an internal reference. Primer Premier 5 (Premier Biosoft, Palo Alto, CA, USA) was used to build the primers required to perform RT-qPCR assays (Table S1). Total RNA from C. oleifera seeds was extracted using the RN53-EASYspin Plus polysaccharide polyphenol/complex plant RNA Rapid Extraction Kit (Aidlab, Beijing, China), and first-strand cDNA was obtained using the HiScript III first strand cDNA synthesis kit (+gDNA wiper) (Vazyme, Nanjing, China). The cDNA was then diluted 5-fold and used as the template for RT-qPCR, with each sample including three technical replicates. The 2–ΔΔCt method was used to calculate the relative transcription levels of the genes.

2.8. Statistical Analysis Method

The data for oil content, fatty acid composition, and free fatty acid content were presented as the mean ± standard deviation (SD) of three biological replicates. The significance of differences among different groups was determined using one-way analysis of variance followed by Tukey’s post hoc test in IBM SPSS Statistics 27 software (IBM Corp, Armonk, NY, USA). Differences were considered statistically significant at p < 0.05. The correlation between RT-qPCR and RNA-seq data was assessed using Pearson correlation coefficients, and the significance of these correlations was evaluated using p-values. Both RT-qPCR and RNA-seq data were log2-transformed for normalization before analysis.

3. Results

3.1. Total Oil Content and Proportions of Major FA Components in Seeds of Different Varieties

The results showed the H variety had a considerably higher oil content, exceeding that of the L and M varieties by 34.97 and 34.30%, respectively (Figure 1). While the M variety exhibited a marginally higher oil content than the L variety, this difference was not statistically significant. C18:1 emerged as the dominant FA component in all varieties’ kernels, making up over 70% of the FA composition, with C16:0, C18:0, C18:2, and C18:3 following in sequence. Notably, an inverse relationship was observed between oil content and C18:1 proportion, with a corresponding increase in C18:2 and C18:3.

Figure 1.

Figure 1

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Total oil content and proportions of major FA components in seeds of three varieties (T4, October 26). (A) Total oil content of each variety. (B–F) Proportion of palmitic acid (C16:0), stearic acid (C18:0), oleic acid (C18:1), linoleic acid (C18:2), and linolenic acid (C18:3), respectively. Significance levels: *P < 0.05, **P < 0.01. L, XL192; M, XL108; H, HH2.

Overall, significant variations in FA composition were observed among the different C. oleifera varieties examined in this study. Specifically, in FA composition, the proportions of C18:0 and C18:1 were considerably greater in the L variety than in the M and H varieties. In contrast, the M variety showed a higher preference for C18:1 and C18:2 proportions. H variety had significantly higher proportions of C18:2 and C18:3 than the M and L varieties, indicating significant differences in unsaturated FA preference among these varieties.

3.2. Dynamic Changes of Free Fatty Acid Content in Kernels of Different Varieties

This study further explored FA accumulation patterns across different varieties by analyzing FFA composition and content in kernels at four maturation stages. As shown in Figure 2A, across all varieties, the contents of free C16:0, free C18:0, free C18:1, and free C18:2 exhibited a general upward trend over time. Conversely, free C18:3 content demonstrated a general decline throughout the period. It is noteworthy that at T2, the L and M varieties experienced a slight decrease in free C18:2 content compared to T1. Furthermore, in the H variety, free C18:3 content surged by 41.04% from T1 to T2, then plummeted by 60.05% from T2 to T3. During the T3–T4 stage, the L variety saw significant increases in free C18:0 and C18:1 content.

Figure 2.

Figure 2

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Content of free fatty acid components in different stages of seeds in three varieties. (A) Temporal trends of major FFA from T1 to T4. (B) Levels of major FFA in kernels of each variety at T4. Significance levels: *P < 0.05, **P < 0.01. L, XL192; M, XL108; H, HH2; T1, September 14; T2, September 28; T3, October 12; T4, October 26.

As shown in Figure 2B, at T4, alongside an increase in overall oil content, there was a decreasing trend in the contents of free C18:0 and C18:1 in the kernels. Interestingly, at T4, despite the C18:2 proportion being significantly higher in the H variety compared to the L and M varieties (Figure 1), the M variety exhibited the highest free C18:2 content, followed by the H and L varieties (Figure 2B). Moreover, at T4, the free C18:3 content in the H variety was comparable to that in the L and M varieties, showing no significant differences (Figure 2B). However, in terms of C18:3 proportion, the H variety was much higher than the L and M varieties (Figure 1). These observations indicate that at T4 in the H variety, a substantial portion of C18:2 and C18:3 may be present as bound FA.

3.3. Characterization of the Transcriptomes and Identification of Differentially Expressed Genes

The molecular mechanisms of oil biosynthesis during C. oleifera seed development were investigated through RNA-seq analysis on samples from various developmental stages. Three biological replicates were established for each developmental stage of the seed samples to ensure data reliability. From the 36 libraries, approximately 1.571 billion high-quality clean reads were generated in total (Table S2), which were used for subsequent alignment analysis. Of these, approximately 1.405 billion (89.45%) high-quality reads were successfully mapped, of which 1.235 billion (78.61%) reads were uniquely mapped (Table S3). Regarding gene expression, the number of valid expressed genes with FPKM values exceeding 1.0 in L variety at T1, T2, T3, and T4 stages was 28,781, 26,870, 27,151, and 24,384, respectively; the corresponding numbers for M variety were 26,431, 25,573, 25,582, and 25,374; and those for H variety were 25,062, 28,617, 24,779, and 24,833 (Table S4).

In the pairwise comparisons between varieties at identical stages, 36,244 DEGs were identified between L and M varieties (L1 vs M1, L2 vs M2, L3 vs M3, and L4 vs M4) (Figure 3A), while 36,918 DEGs were identified between H and M varieties (H1 vs M1, H2 vs M2, H3 vs M3, and H4 vs M4) (Figure 3A). Furthermore, more DEGs were identified in the pairwise comparisons of adjacent stages within the variety with higher oil content (Figure 3C). Specifically, 24,031 DEGs were identified in L variety (L2 vs L1, L3 vs L2, and L4 vs L3) (Figure 3B), 29,815 DEGs were identified in M variety (M2 vs M1, M3 vs M2, and M4 vs M3) (Figure 3B), and the most DEGs were identified in H variety (H2 vs H1, H3 vs H2, and H4 vs H3), with a total of 33,505 DEGs (Figure 3B).

Figure 3.

Figure 3

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Differential gene expression statistics. (A) Venn diagram of differential genes between groups. (B) Venn diagram of differential genes within groups. (C) Number of differential genes between different groups. Red indicates upregulated genes, and blue indicates downregulated genes.

Across all same-stage pairwise comparisons between L and M varieties and H and M varieties (L1 vs M1, H1 vs M1, L2 vs M2, H2 vs M2, L3 vs M3, H3 vs M3, L4 vs M4, H4 vs M4), a total of 27,981 annotated DEGs were identified (Table S5). At the same time, 27,600 DEGs were found in the pairwise comparisons of adjacent stages within L, M, and H varieties (L2 vs L1, L3 vs L2, L4 vs L3, M2 vs M1, M3 vs M2, M4 vs M3, H2 vs H1, H3 vs H2, H4 vs H3) (Table S6). A total of 26,344 DEGs, common across both comparison categories, were designated as significant differentially expressed genes (SigDEGs). These SigDEGs were identified in both same-stage pairwise comparisons and adjacent-stage pairwise comparisons (Table S7), highlighting their substantial value in exploring gene expression dynamics.

The identified SigDEGs underwent further analysis through GO and KEGG enrichment (Figure 4). GO enrichment analysis revealed that, within the biological process category, the predominant GO terms included metabolic process, cellular metabolic process, and organonitrogen compound metabolic process. For the cellular component category, leading GO terms comprised membrane, intracellular, and organelle. In the molecular function category, the top three GO terms were catalytic activity, small molecule binding, and nucleotide binding (Figure 4A). Moreover, KEGG enrichment analysis identified 51 significantly enriched pathways, encompassing plant hormone signal transduction, starch and sucrose metabolism, photosynthesis, fatty acid elongation, steroid biosynthesis, ABC transporters, arachidonic acid metabolism, pantothenate and CoA biosynthesis, among others (Figure 4B).

Figure 4.

Figure 4

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GO and KEGG enrichment analysis of SigDEGs. (A) GO enrichment analysis of SigDEGs. (B): KEGG enrichment analysis of SigDEGs.

3.4. Weighted Gene Coexpression Network Analysis of SigDEGs

This study employed the median absolute deviation (MAD) to filter out noise among SigDEGs, selecting 14,608 SigDEGs with MAD values ≥1.0 for WGCNA analysis. The analysis divided these 14,608 SigDEGs into 12 coexpression modules (Figure 5A), of which the blue module (4,943 genes) was most correlated with C18:2, while the green-yellow (435 genes) and dark-green modules (4,232 genes) both showed high correlations with C18:3 (Figure 5B). Despite the green-yellow module showing the highest correlation with C18:3, it contained only a few genes involved in lipid syntheses, such as FATB, GPAT, and LPCAT (Table S8). In contrast, large numbers of lipid synthesis-related genes were found in the blue and dark-green modules, including ACCase, KAR, KAS I, KAS II, KAS III, SAD, FAD6, FAD7, FAD8, FATB, LACS, GPAT, and PDAT genes (Tables S9 and S10).

Figure 5.

Figure 5

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WGCNA analysis of SigDEGs. (A) Clustering dendrograms of SigDEGs. (B) Heatmap showing the correlation between modules and free fatty acid content. (C) Network relationship of genes related to oil synthesis and TFs in the blue module. (D) Network relationship of genes related to oil synthesis and TFs in the dark-green module.

Notably, both modules contained FAD6 and FAD7, while FAD8 only appeared in the dark-green module (Figure 5C,D). Furthermore, TFs identified in these modules underscore their crucial regulatory roles in FA synthesis. The TFs identified in the two modules included MYB, bZIP, DOF, WRI1, LEC, ABI3, WRKY, bHLH, GRF, ARF, and ERF. Protein kinases related to abscisic acid signaling, such as PP2C and SNRK, were also found in the two modules. These findings highlight the significant roles of phytohormones, including ethylene, auxin, and abscisic acid, in lipid synthesis.

3.5. Expression Patterns of Genes Related to Oil Synthesis Pathways during the Maturation of Kernels of Different Varieties

Within the de novo FA synthesis pathway, genes such as ACCase, KAS I, KAS II, KAS III, KAR, HAD, EAR, and SAD in the H variety were significantly upregulated at T4 (Figure 6), indicating that FA synthesis activity may be relatively more active in the T3–T4 stage of the H variety. Specifically, FAD6 experienced significant upregulation during the T2–T3 stage in the H variety. Notably, the three transcripts of FAD7 in the H variety were mainly upregulated in the T1–T2 and T2–T3 stages, while FAD8 was upregulated in the T3–T4 stage. In comparison, the FAD7 gene in the M variety was mainly upregulated in the T2–T3 stage, but FAD8 gene expression levels were almost unchanged across stages. In the L variety, FAD7 and FAD8 only showed slight upregulation in the T2–T3 stage.

Figure 6.

Figure 6

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Oil synthesis pathway map and heatmap of gene expression related to oil synthesis. Heatmaps were generated using log2 FC values of gene expression between adjacent stages of each variety (Table S11). Reaction abbreviations: accA, acetyl-coenzyme A carboxylase carboxyl transferase subunit alpha; accB, biotin carboxyl carrier protein of acetyl-CoA carboxylase; ACCase, Acetyl-CoA carboxylase; ACS, acetyl-coenzyme A synthetase; DGAT, diacylglycerol acyltransferase; EAR, enoyl-[acyl-carrier-protein] reductase; FAD2, fatty acid desaturase 2; FAD3, fatty acid desaturase 3; FAD6, fatty acid desaturase 6; FAD7, fatty acid desaturase 7; FAD8, fatty acid desaturase 8; FATA, acyl acyl-carrier-protein thioesterase type A; FATB, acyl acyl-carrier-protein thioesterase type B; G3PDH, glycerol-3-phosphate dehydrogenase; GK, glycerol kinase; GPAT, sn-glycerol-3-phosphate acyltransferase; HAD, 3-hydroxyacyl-[acyl-carrier-protein] dehydratase; KAR, ketoacyl-ACP reductase; KASI, ketoacyl-ACP synthase I; KASII, ketoacyl-ACP synthase II; KASIII, ketoacyl-ACP synthase III; LACS, long chain acyl-CoA synthetase; LEC1, leafy cotyledon 1; LPAAT, lysophosphatidic Acid Acyltransferase; LPCAT, 1-acylglycerol-3phosphate acyltransferase; MCAT, malonyl-CoA-acyl carrier protein transacylase; Ole I, oleosin I; Ole II, oleosin II, Ole III, oleosin III; Ole IV, oleosin IV; Ole V, oleosin V; PAP, phosphatidate phosphatase; PDAT, phospholipid: diacylglycerol acyltransferase; PLA, phospholipase A; PLC, phospholipase C; PLD, phospholipase D; SAD, stearoyl-ACP desaturase; WRI1, wrinkled1.

Within the TAG synthesis pathway, the FAD3 gene in the H variety was observed to be upregulated at both the T1–T2 and T3–T4 stages, up-regulated in the M variety at the T2–T3 stage, and showed no significant changes in all stages in the L variety. The difference in FAD expression between different varieties may explain the difference in C18:3 content. In addition, compared to the L variety and M variety, the corresponding transcripts of the DGAT gene in the H variety were up-regulated in all stages, indicating that TAG assembly activity in the H variety was also relatively active in all stages. Particularly at the T3–T4 stage, significant upregulation was observed for the LPAAT, LACS, DGAT, and PDAT genes in the H variety. Notably, the Ole gene in the H variety was significantly upregulated at the T3–T4 stage, suggesting a pivotal role for oleosin in lipid synthesis. G3PDH provides the glycerol backbone for TAG biosynthesis, and its high expression at the T3–T4 stage provides sufficient raw materials for TAG biosynthesis. Furthermore, the significant upregulation of WRI1 and LEC1 in the H variety at the T3–T4 stage underscores their critical roles in regulating lipid synthesis pathways.

In conclusion, during the T3–T4 stage, multiple genes related to oil synthesis in the H variety were upregulated, which may be the main reason why its oil content is higher than that of the L variety and the M variety. The enzymes encoded by genes such as FAD3, FAD7, and FAD8 are involved in the synthesis of C18:3. The high expression of these genes in the H variety effectively increased the content of C18:3 in the seeds.

3.6. Verification of RNA-Seq Data with RT-qPCR

Twelve DEGs associated with oil synthesis were selected for RT-qPCR experiments to confirm the accuracy of the RNA-seq data. The outcomes revealed that the expression patterns of the majority of these genes were closely aligned between the RT-qPCR results and the RNA-seq data (Figure 7). This high level of similarity confirms the reliability of the RNA-seq data, underscoring its value in identifying and analyzing gene expression related to oil production.

Figure 7.

Figure 7

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Comparison and validation of RT-qPCR and RNA-seq results. The bar chart shows the expression level of FPKM (right Y-axis), and the line chart shows the relative expression level of RT-qPCR (left Y-axis). The error bars in the figure represent the SD. The correlation coefficients and P-values in the figure represent the mean values for each group. The abbreviations and full names of the related genes are as follows: accA, acetyl-coenzyme A carboxylase carboxyl transferase subunit alpha; accB, biotin carboxyl carrier protein of ACCase; KAR, ketoacyl-ACP reductase; KASIII, ketoacyl-ACP synthase III; FAD6, fatty acid desaturase 6; FAD7, fatty acid desaturase 7; FAD8, fatty acid desaturase 8; FATA1, acyl acyl-carrier-protein thioesterase type A1; PDAT, phospholipid: DGAT; GK, glycerol kinase; MYB1, myeloblastosis TF 1; bZIP1, basic leucine zipper 1.

4. Discussion

In this study, the oil content of the three C. oleifera varieties on October 26 (T4) was highest in HH2 (H), followed by XL108 (M), and lowest in XL192 (L). Notably, the L variety exhibited a significantly higher C18:0 content compared to the M and H varieties. An intriguing inverse relationship emerged between the oil content and C18:1 proportion, while a direct relationship was observed with the proportion of C18:2 and C18:3. Overall, the differences in FA composition among the varieties mainly reflect their preference for FA desaturation. The L group tended to produce less desaturated FA, such as C18:0 and C18:1. In contrast, the M group preferred moderately desaturated FA, such as C18:1 and C18:2. The H group tended to generate highly unsaturated FA, particularly C18:2 and C18:3. Transcriptome sequencing of the seeds identified 43,865 DEGs, among which 26,344 genes were commonly identified across both the pairwise comparisons between varieties within the same period (L1 vs M1, H1 vs M1, L2 vs M2, H2 vs M2, L3 vs M3, H3 vs M3, L4 vs M4, H4 vs M4) and the pairwise comparisons between adjacent stages within each group (L2 vs L1, L3 vs L2, L4 vs L3, M2 vs M1, M3 vs M2, M4 vs M3, H2 vs H1, H3 vs H2, H4 vs H3). In-depth mining of these 26,344 DEGs uncovered significant varietal differences in key genes, including SAD and FAD. Additionally, TFs and phytohormones were identified as playing crucial roles in FA biosynthesis in the kernels. In the high-oil group (H), the coordinated high expression of multiple genes involved in de novo FA synthesis and TAG assembly pathways was identified as a primary reason for the high oil content in seeds. Overall, the insights gained from this research offer valuable guidance and constitute an important reference for breeding C. oleifera with high oil content and improved quality.

4.1. The Coordinated Expression of SAD and FAD Genes Affects the FA Composition in C. oleifera Kernels

During seed oil biosynthesis, SAD, FAD6, FAD7, and FAD8 in plastids, alongside FAD2 and FAD3 in the endoplasmic reticulum, enhance FA desaturation by introducing double bonds.11 In plastids, C18:0 undergoes successive introduction of one double bond by SAD, FAD6, and FAD7/FAD8 to form C18:1, C18:2, and C18:3, respectively.36 In the endoplasmic reticulum, C18:1 in phosphatidylcholine (C18:1 PC) can be introduced with one double bond by FAD2 and FAD3 to generate C18:2 PC and C18:3 PC, respectively (Figure 6). Previous studies have demonstrated that the elevated C18:1 level in the kernels of C. oleifera is associated with sustained high levels of transcription in the SAD gene and coordinated inadequate transcription of the FAD2, FAD3, FAD7, and FAD8 genes.30,31,37

In this study, SAD gene in the L variety had corresponding transcript upregulation at different stages, while genes including FAD3, FAD7, and FAD8 were expressed at low levels at most stages, limiting the conversion from C18:1 to C18:2 and C18:3, leading to C18:1 accumulation. Such findings align with those from previous research. For the H variety, the higher activity of FAD3, FAD7, and FAD8 indicates that C18:3 synthesis was more active at all stages, resulting in a lower proportion of C18:1 and a higher proportion of C18:3. The differential expression of these FAD genes is the main reason for the differences in FA composition among the seeds of different C. oleifera varieties.

4.2. Potential Feedback Regulation in FA Biosynthesis

In this study, almost all genes in the de novo FA synthesis pathway in the H group showed a coordinated upregulation effect from T3 to T4, promoting the entry of carbon sources into the FA biosynthesis pathway during this period. The lipidomic data further supports this view, as we found that most of the C18:2 and C18:3 in the H group during the T3–T4 stage existed in the form of bound FA, indicating highly active FA synthesis during this period. Therefore, the coordinated upregulation of multiple key genes during the late maturation stage of the seed is the main reason for the high oil content. It is worth mentioning that this phenomenon has rarely been reported in previous studies. Additionally, we observed that the levels of free C18:1 in the H group remained low at all stages. From T3 to T4, the free C18:1 content in the H group even decreased (Figure 2). Furthermore, the proportion of C18:1 in the H group was the lowest among all groups (Figure 1). We speculate that the coordinated upregulation of multiple key genes in the later stages in the H group may be related to the low levels of C18:1. In biosynthetic pathways, the accumulation levels of downstream products can feedback regulate the activity of initial enzymes, thereby adjusting the synthesis rate of the pathway and maintaining metabolic balance.38 Previous studies have also shown that the accumulation of 18:1-ACP inhibits the activity of ACCase in rapeseed without affecting FA elongation.39 However, further research is needed to verify whether the low levels of C18:1 in seeds might activate ACCase activity through potential feedback regulation mechanisms, thereby promoting the entry of carbon sources into the fatty acid synthesis pathway.

TAG usually remains in oil bodies after assembly, with ole being the most abundant protein on the phospholipid membrane.40,41 Despite the ability of oil bodies to form without oleosin, numerous pieces of evidence imply that ole serves an essential part of preserving oil body stability.42−44 In this research, multiple ole were significantly elevated within the H variety at the T3–T4 stages, accompanied by a significant up-regulation of TAG biosynthesis genes such as LPAAT, DGAT, and PDAT (Figure 6). Ole1 overexpression has been shown to promote oil accumulation by inhibiting TAG degradation-associated transport proteins and lipases, and by activating enzymes related to TAG assembly.45−47 TAG is one of the ultimate products for the Kennedy pathway. However, there is currently no evidence whether the content of “free TAG” can feedback-regulate the enzymes or TFs involved in TAG biosynthesis.

4.3. Regulatory Roles of TFs in FA Biosynthesis

The WRI1 acts as a pivotal regulator within oilseed FA synthesis pathways, controlling multiple enzymes for glycolysis and de novo FA synthesis.48,49 Additionally, LEC1 regulates FA synthesis depending on FUS3, ABI, and WRI1.50 Elevated levels of WRI1 and LEC1 are strongly connected to enhanced seed oil production.51,52 Thus, the simultaneous upregulation of ACCase, EAR, HAD, KAR, KASI, KASII, KASIII, and SAD genes in the H variety during T3–T4 stages may correlate with elevated LEC1 and WRI1 gene expression. Furthermore, although previous studies have suggested that TAG biosynthesis in the endoplasmic reticulum is not regulated by WRI1,53 recent research has indicated that WRI1 can target genes related to TAG biosynthesis, such as G3PHD, DGAT2, and PDAT.54,55 The data from this research indicate that G3PHD, DGAT, and PDAT also exhibited significant up-regulation in the H variety at the T3–T4 stage, which further supports the potential regulatory role of WRI1 in TAG biosynthesis.

Earlier research indicates that R2R3-type MYB TFs generally exert a negative influence on oil biosynthesis in the C. oleifera seeds, with a minority being implicated in long-chain FA synthesis.29,56 In this study, 13 MYB TFs were identified as potentially involved in the regulation of fatty acid synthesis based on WGCNA (Tables S9 and S10). Moreover, research has demonstrated that the overexpression of DOF stimulates oil buildup in many experiments.57−59 In this study, 9 DOFs were identified from the blue and dark-green modules based on WGCNA (Tables S9 and S10), which may be closely related to oil synthesis. The production of oil and the development of seeds are strongly influenced by bZIP TFs. In Arabidopsis,60bZIP67 can increase the content of C18:3 in Arabidopsis seeds by activating the desaturase FAD3. Further research shows that bZIP67 can cooperate with LEC1, AREB3, and ABI3 to oversee gene expression throughout the seed development process.61 In addition, transgenic Arabidopsis seedlings have also been shown to have higher lipid contents when GmbZIP123 is overexpressed.62 This suggests that bZIP positively contributes to seed lipid accumulation. In this work, according to WGCNA, a total of 10 bZIPs that might be associated with oil biosynthesis were found (Tables S9 and S10).

4.4. Crucial Role of Phytohormones in FA Production

Phytohormones are essential for the production of FA.63,64 According to the result of WGCNA in this study, FA biosynthesis is significantly impacted by ARF, ERF, SNRK, and PP2C. ARF and ERF stand out as crucial TFs within the auxin as well as ethylene signaling pathways, respectively. By attaching to certain DNA sequences, they influence target gene transcription, thus impacting plant development and growth.65,66 Auxin has been shown to enhance FA synthesis in microalgae and increase monounsaturated FA levels.67,68 For research on wheat69 and Arabidopsis,70 auxin is connected with the production of very long-chain FA. These very long-chain FAs play a function in plant reactions to stress from both abiotic and biotic sources.71 On the other hand, ERF is also a key TF when plants respond to stress.72 Camellia fruit has a long development period, during which it is susceptible to abiotic stresses. Hence, these hormonal response factors significantly contribute to Camellia fruit’s growth and development. Moreover, recent findings have demonstrated ethylene’s capacity to boost the levels of C18:2 and C18:3 within C. oleifera fruit.73 This also indicates that there is a complex relationship between plant hormones and plant lipids. Particularly under stress conditions, lipid remodeling emerges as a critical strategy for plant response to abiotic stressors.74,75 Studies have shown that spraying exogenous ABA on C. oleifera leaves under shortages of water significantly promotes the activity of genes such as accA, accB, FAD6, FAD7, and FAD8 in the leaves.76 According to another study, the application of external ABA promoted the transcription of FAD2 within oil palm fruit, which resulted in the accumulation of C18:2.77 This suggests the involvement of ABA in FA synthesis, with SNRK and PP2C as key elements in ABA signaling.

In the realm of agricultural practices, the strategic use of certain plant hormones has begun to show promising results in the growth and development of C. oleifera fruits.73 These approaches not only contribute to an increase in oil accumulation within the fruit but also aids in enhancing its overall quality. Such advancements highlight the potential of targeted hormone application in optimizing crop yields and improving the nutritional value of agricultural products. Given the intricate polyploidy and the wide array of C. oleifera varieties, there’s a pressing need for more in-depth research to uncover the molecular mechanisms behind how plant hormones influence oil accumulation in its fruit. Additionally, fine-tuning the application of these hormones and assessing their long-term impacts are essential measures to ensure the economic sustainability of agricultural practices. This dual approach will not only pave the way for maximizing oil yield and quality but also bolster the economic feasibility of cultivating C. oleifera on a larger scale.

5. Conclusion

This research undertook an in-depth investigation of the transcriptome and lipidome in C. oleifera seeds at the maturing process, unveiling the FA accumulation patterns and their intricate association with gene expression regulation. The transcriptome analysis led to the identification of 26,344 genes that are expressed differently. Functional enrichment analysis indicated the involvement of phytohormone signaling transduction as well as starch and sucrose metabolism in oil accumulation. The notable high oil content in the kernel is largely due to the combined elevated expression of genes involved in the de novo fatty acid synthesis and TAG assembly pathways. Additionally, varietal expression variations in fatty acid desaturases result in differing unsaturated FA component levels among C. oleifera varieties. Particularly in high-oil varieties (HH2), the high expression of genes such as FAD3, FAD7, and FAD8 during maturation increases the proportion of C18:3 at the expense of the proportion of C18:1. In conclusion, the findings of this study provide new insights into the molecular mechanisms of oil biosynthesis in C. oleifera seeds, offering important guidance for improving oil content and optimizing its quality.

Data Availability Statement

Availability of Data The RNA-Seq data of Camellia oleifera seeds were uploaded to SRA with accession PRJNA1073581 (https://www.ncbi.nlm.nih.gov/bioproject/?term=PRJNA1073581).

Supporting Information Available

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acs.jafc.4c03614.

  • Table S1, primers used in RT-qPCR (XLSX)

  • Table S2, quality assessment metrics for RNA sequencing of C. oleifera varieties across different maturation stages (XLSX)

  • Table S3, ,apping statistics for RNA-Seq reads in C. oleifera across developmental stages (XLSX)

  • Table S4, gene expression distribution by FPKM intervals in the seeds of C. oleifera (XLSX)

  • Table S5, annotated DEGs from adjacent stage comparisons within L, M, and H varieties of C. oleifera (XLSX)

  • Table S6, annotated DEGs from L-M and H-M same-stage pairwise comparisons in C. oleifera (XLSX)

  • Table S7, SigDEGs common to same-stage and adjacent-stage comparisons in C. oleifera (XLSX)

  • Table S8, gene characteristics of the greenyellow module from WGCNA in C. oleifera(XLSX)

  • Table S9, gene characteristics of the blue module from WGCNA in C. oleifera (XLSX)

  • Table S10, gene characteristics of the garkgreen module from WGCNA in C. oleifera (XLSX)

  • Table S11, detailed gene expression Log2 fold changes for oil synthesis pathway genes in the seeds of C. oleifera (XLSX)

Author Contributions

∥ D.Y., R.W., and H.L. have contributed equally to this work.

This work was supported by the Hunan Province Science and Technology Innovation Project (2021JC0007), Hunan Province Special Project for Innovative Province Construction (2022JJ30325), Seed Industry Innovation Project of Hunan Province (2021NK1007), and Hunan Provincial Forestry Science and Technology Innovation Fund Project (XLK202101-1).

The authors declare no competing financial interest.

Supplementary Material

jf4c03614_si_001.xlsx (10.5KB, xlsx)
jf4c03614_si_002.xlsx (13.8KB, xlsx)
jf4c03614_si_003.xlsx (15.6KB, xlsx)
jf4c03614_si_004.xlsx (13.7KB, xlsx)
jf4c03614_si_005.xlsx (2.2MB, xlsx)
jf4c03614_si_006.xlsx (2.2MB, xlsx)
jf4c03614_si_007.xlsx (2.1MB, xlsx)
jf4c03614_si_008.xlsx (52.6KB, xlsx)
jf4c03614_si_009.xlsx (470.7KB, xlsx)
jf4c03614_si_010.xlsx (405.6KB, xlsx)
jf4c03614_si_011.xlsx (26.3KB, xlsx)

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

jf4c03614_si_001.xlsx (10.5KB, xlsx)
jf4c03614_si_002.xlsx (13.8KB, xlsx)
jf4c03614_si_003.xlsx (15.6KB, xlsx)
jf4c03614_si_004.xlsx (13.7KB, xlsx)
jf4c03614_si_005.xlsx (2.2MB, xlsx)
jf4c03614_si_006.xlsx (2.2MB, xlsx)
jf4c03614_si_007.xlsx (2.1MB, xlsx)
jf4c03614_si_008.xlsx (52.6KB, xlsx)
jf4c03614_si_009.xlsx (470.7KB, xlsx)
jf4c03614_si_010.xlsx (405.6KB, xlsx)
jf4c03614_si_011.xlsx (26.3KB, xlsx)

Data Availability Statement

Availability of Data The RNA-Seq data of Camellia oleifera seeds were uploaded to SRA with accession PRJNA1073581 (https://www.ncbi.nlm.nih.gov/bioproject/?term=PRJNA1073581).

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