Comparison of fruit organic acids and metabolism-related gene expression between Cerasus humilis (Bge.) Sok and Cerasus glandulosa (Thunb.) Lois

Xiaopeng Mu; Pengfei Wang; Junjie Du; Yu Gary Gao; Jiancheng Zhang

doi:10.1371/journal.pone.0196537

Abstract

Cerasus humilis (Bge.) Sok and Cerasus glandulosa (Thunb.) Lois are economically important fruit-producing shrubs. Although these two species have similar looking fruits, their fruit organic acid contents differ drastically. In this study, we focused on comparing the organic acid content, activity of enzymes gene expression involved in organic acid metabolism in both C. humilis and C. glandulosa fruits. To investigate the differences of organic acid metabolism in fruits of these two species, a comparative transcriptome analysis was performed. Our results showed that temporal changes of two main organic acids exhibited different trends in these two species. Transcriptome sequencing of developing C. humilis and C. glandulosa fruits systematically revealed 6,594 differentially expressed genes. Compared with C. humilis, the expression levels of 3,469 and 3,125 genes were up- and down-regulated in C. glandulosa, respectively, including one PEPC gene, 12 malic acid metabolism genes, 25 citric acid cycle genes, and 194 NAD/NADP metabolism genes. The correlation analysis and principal component analysis of gene expression, enzymatic activity and organic acid content showed that differences in the expression of genes encoding the NAD-malate dehydrogenase (NAD-MDH) and NADP-malate enzyme (NADP-ME) contributed substantially to the observed differences in organic acid accumulation of two species. Our results provide a solid foundation for future elucidation of key mechanisms regulating organic acid biosynthesis in C. humilis and C. glandulosa fruits and could lead to efficient and highly targeted generation of more commercially accepted cultivars.

Citation: Mu X, Wang P, Du J, Gao YG, Zhang J (2018) Comparison of fruit organic acids and metabolism-related gene expression between Cerasus humilis (Bge.) Sok and Cerasus glandulosa (Thunb.) Lois. PLoS ONE 13(4): e0196537. https://doi.org/10.1371/journal.pone.0196537

Editor: Ji-Hong Liu, Key Laboratory of Horticultural Plant Biology (MOE), CHINA

Received: December 23, 2017; Accepted: April 14, 2018; Published: April 26, 2018

Copyright: © 2018 Mu et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Data Availability: Transcriptome data of C. humilis and C. glandulosa are available from the NCBI BioProject (accession number PRJNA417674) and the SRA database (accession numbers SRX3380120 and SRX3380121).

Funding: This work was supported by the Major Subject of Shanxi Science and Technology Research (Grant No. 20121101010), the China Postdoctoral Science Foundation (Grant No. 2017M621106) and the Doctoral Research Fund of Shanxi Agriculture University (Grant No. 2015ZZ19). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Competing interests: The authors have declared that no competing interests exist.

Introduction

The acidity of fleshy fruit is an important component of fruit organoleptic quality [1,2]. Fruit acidity is due to the presence of organic acids synthesized within fruits [3], with considerable changes in production during different developmental stages [4–6]. The variations in the production of organic acids during different developmental stages vary drastically among species. For example, the main organic acids in ‘Honeycrisp’ apple flesh increase gradually with fruit development [6], while the content of the main organic acids in sand pear fruits decline with fruit development [7].

Organic acids could be utilized as the substrates in the glycolytic pathway, trimesic acid cycle, and gluconeogenesis during fruit maturation process [8]. Therefore, fruit organic acid content is closely associated with changes in the activities of related enzymes. For example, phosphoenolpyruvate carboxylase (PEPC), NAD-dependent malate dehydrogenase (NAD-MDH), and NADP-malic enzyme (NADP-ME) are important for the synthesis of malic acid and citric acid [9–12]. Previous studies revealed that PEPC activity is positively correlated with navel orange organic acid content [13], while the PEPC gene expression level and associated protein content help regulate peach and pineapple fruit acidity [14]. Additionally, PEPC activity is also affected by malic acid, pyruvic acid, cytoplasmic pH, and reversible phosphorylation during fruit organic acid metabolism [15–17]. Thus, the effects of PEPC on fruit organic acid metabolism are complex. Furthermore, NAD-MDH activity is positively correlated with malic acid content in nectarines [18], while the expression of the NAD-MDH gene in peach and apple does not directly affect fruit acidity [19,20]. In most fruits, NADP-ME plays a major role in the degradation of malic acid. The organic acid content of loquat fruit is negatively correlated with NADP-ME activity [18]. In grapes, the expression level of the NADP-ME gene is relatively high in the young fruits, declines in the maturing fruits, and increases when the fruits mature [21].

The genetic regulation of organic acids is complex and most often involves polygenes. However, the organic acid content of some plant species that produce moderately or highly acidic fruits is controlled by one major gene (D model), with low acidity being a recessive trait [8]. Previous studies concluded there is no correlation or no significant correlation between the expression of organic acid metabolism-related genes and organic acid content [14,19,22]. However, it was reported that the expression of the cytosolic NADP-IDH gene in lemon fruits is well correlated with changes in NADP-IDH (NADP⁺ -isocitrate dehydrogenase) activity and promotes the degradation of organic acids [23].

Cerasus humilis is a small shrub grown in China for its fruit [24]. It is highly stress-resistant, especially to drought and low temperatures [25,26]. Additionally, its fruit contains more calcium, flavonoids, vitamins, and other nutrients than many other fruits [27], making it a very promising multi-purpose fruit tree species. It is rare for C. humilis cultivars to produce fruits with low acidity, which greatly limits the commercial potential of this species [28]. Cerasus glandulosa is closely related to C. humilis but produces fruits with lower organic acid content [29]. There have been no studies that examined the effects of enzyme activities and gene expression levels on organic acid content in C. humilis or C. glandulosa fruits. In this study, we investigated the temporal changes in organic acid content, the activities of key enzymes, and the expression levels of genes associated with malic acid metabolism in the developing fruits of C. humilis and C. glandulosa. Our results should be useful for improving the fruit quality of C. humilis and C. glandulosa, and for characterizing the molecular mechanisms regulating fruit organic acid biosynthesis.

Materials and methods

The fruits of C. humilis (cv. Jinou 1; high acidity) and C. glandulosa (cv. DS-1; low acidity) were used as the test materials in this study. Because of the asynchrony in the developmental processes of C. humilis and C. glandulosa fruits, C. humilis fruit samples were collected at seven developmental stages (8, 10, 12, 14, 16, 18, and 19 weeks after flowering), while C. glandulosa fruit samples were collected at five developmental stages (8, 10, 12, 13, and 14 weeks after flowering), from plants grown at the experimental farm of the Horticultural College, Shanxi Agricultural University, Taigu, China. Following collection, the fruit samples were immediately frozen in liquid nitrogen and stored at -80 °C until analysis.

Determination of total acid, malic acid, and citric acid contents

Fruit organic acid extraction and determination were carried out with some modifications to the methods described by Nergiz and Ergönül [30]. Samples were analyzed with HPLC (UltiMate3000 system, Thermo Fisher, USA), and the analysis conditions were as follows: chromatographic separation was performed on a Syncronis C18 column (5 μm, 250 mm × 4.6 mm); mobile phase was methanol:0.01mol/L K₂HPO₄ solution (10:90, pH = 2.0) with a flow rate of 0.5 mL/min; column temperature was set at 25 °C; sample injection volume was 15 μL; detection wavelength was set at 210 nm. Three replicates of each sample were analyzed.

Determination of enzyme activity

Enzyme samples were prepared as described by Chen et al [8]. Enzyme activity was determined as described by Hirai and Ueno [31], but the reaction volume was changed to 3 mL. The enzyme activity of NAD-MDH was measured in reaction mixture containing 300 μL Tris-HCl (800 mM/L, pH 8.2), 150 μL KHCO₃ (800 mM/L), 150 μL MgCl₂ (40 mM/L), 150 μL GSH (10 mM/L), 150 μL NADH (3 mM/L), 1600 μL OAA (4 mM/L), and 500 μL enzyme extract; The enzyme activity of NADP-ME was measured in reaction mixture containing 300 μL Tris-HCl (800 mM/L, pH 7.4), 150 μL MnSO₄ (4 mM/L), 150 μL NADP (3.4 mM/L), 300 μL ddH₂O, 1600 μL malic acid (4 mM/L), and 500 μL enzyme extract; The enzyme activity of PEPC was measured in reaction mixture containing 300 μL Tris-HCl (800 mM/L, pH 8.5), 150 μL KHCO₃ (200 mM/L), 150 μL MgCl₂ (40 mM/L), 150 μL GSH (10 mM/L),150 μL NADH (3 mM/L), 1600 μL PEP (4 mM/L), and 500 μL enzyme extract. The reaction components were mixed just before adding the substrate. The absorbance (at 340 nm) of the enzyme samples was measured using a UV-2450 spectrophotometer (Shimadzu Corporation, Japan) immediately after adding the substrate (1 s for the unit readings with a 3-min scan). Changes in absorbance were recorded and the analysis was repeated three times. One enzyme unit corresponded to an absorbance change of 0.01 in 1 min, and enzyme activity was expressed in units per gram of pulp (fresh weight) per minute (U g⁻¹ FW min⁻¹).

RNA extraction

Frozen fruit samples (1 g) from each of the different developmental stages (seven stages for C.humilis and five stages for C.glandulosa) were used for the total RNA extraction using a modified CTAB method [27]. RNA quality was then assessed on 1% agarose gels for degradation and contamination. The purity of the extracted RNA was checked using the NanoPhotometer^® spectrophotometer (Implen, CA, USA), while the concentration was measured using the Qubit^® RNA Assay Kit and the Qubit^® 2.0 Fluorometer (Life Technologies, CA, USA). Finally, RNA integrity was assessed using the RNA Nano 6000 Assay Kit and the 2100 Bioanalyzer system (Agilent Technologies, CA, USA).

Library preparation for transcriptome sequencing

The sequencing cDNA libraries were constructed for C.humilis and C.glandulosa respectively with the NEBNext^® Ultra^™ RNA Library Prep Kit for Illumina^® (NEB, USA) following the manufacturer’s recommendations. Index codes were added to attribute sequences to each sample. Briefly, mRNA was purified from total RNA using poly-T oligo-attached magnetic beads and then fragmented by heating in 5X NEBNext First Strand Synthesis Reaction Buffer containing divalent cations. First-strand cDNA was synthesized using a random hexamer primer and M-MuLV Reverse Transcriptase (RNase H−). Second-strand cDNA was synthesized with DNA Polymerase I and RNase H. The remaining overhanging ends were converted to blunt ends via exonuclease/polymerase activities. After adenylating the 3′ ends of cDNA fragments, the NEBNext Adaptor with a hairpin loop structure was ligated for a subsequent hybridization. To select cDNA fragments that were 150–200 bp long, the library fragments were purified with the AMPure XP system (Beckman Coulter, Beverly, USA). Then, 3 μL USER enzyme (NEB, USA) was added to the size-selected, adaptor-ligated cDNA samples, which were then incubated at 37 °C for 15 min and then at 95 °C for 5 min. The subsequent polymerase chain reaction (PCR) was completed with the Phusion High-Fidelity DNA polymerase, universal PCR primers, and the Index (X) Primer. Finally, the PCR products were purified with the AMPure XP system and the library quality was assessed using the 2100 Bioanalyzer system.

Clustering and sequencing

The index-coded samples were clustered using the cBot Cluster Generation System and the TruSeq PE Cluster Kit v4 cBot-HS (Illumina) according to the manufacturer’s instructions. The libraries were then sequenced with the HiSeq 2500 platform (Illumina) to generate paired-end reads.

Raw data (raw reads) in fastq format were first processed using in-house Perl scripts. Clean data (clean reads) were obtained by removing reads with an adaptor, reads containing multiple Ns, and low-quality reads. Additionally, the Q20, Q30, GC content, and sequence duplication level of the clean data were calculated. All downstream analyses were based on high quality clean data.

Transcriptome assembly and function annotation

For all libraries/samples, the left files (read1 files) were pooled into one left.fq file, while the right files (read2 files) were combined into one right.fq file. The transcriptome was assembled based on the left.fq and right.fq files using Trinity [32] with min_kmer_cov set to 2 by default. All other parameters were also set to default values. The transcriptome data of C. humilis and C. glandulosa were submitted to NCBI BioProject under the accession number PRJNA417674.

Gene function was annotated based on the following databases: Nr (NCBI non-redundant protein sequences); Nt (NCBI non-redundant nucleotide sequences); Pfam (Protein family); KOG/COG (Clusters of Orthologous Groups of proteins); Swiss-Prot (manually annotated and reviewed protein sequence database); KO (KEGG Ortholog database); and GO (Gene Ontology).

Quantification of gene expression levels and identification of differentially expressed genes

Gene expression levels were estimated by RSEM [33] using a two-step process. First, clean data were mapped back to the assembled transcriptome. Second, the read count for each gene was obtained based on the mapping results.

For samples with three biological replicates, the analysis of differentially expressed genes between two conditions/groups was completed using the DESeq R package (1.10.1), which detects differentially expressed genes using a model based on the negative binomial distribution. The resulting P-values were adjusted with the Benjamini and Yekutieli approach for controlling the false discovery rate [34]. Genes with an adjusted P-value < 0.05 were considered differentially expressed. Prior to the analysis of differentially expressed genes, the read counts for each sequenced library were adjusted by the edgeR program package with one scaling normalization factor. Genes differentially expressed between two samples were analyzed with the DEGseq (2010) R package. The P-value was adjusted using the q-value [35]. The following threshold was applied to identify differentially expressed genes: q-value < 0.005 and |log₂ (fold change)| ≥ 1. The WEGO online program (http://wego.genomics.org.cn/cgi-bin/wego/index.pl) was used for the GO functional classification of all unigenes and determination of the distribution of gene functions.

Quantitative real-time PCR analysis

Total RNA was used as the template to synthesize cDNA with the PrimeScript^™ RT reagent Kit with gDNA Eraser (Perfect Real Time) (TaKaRa, Japan). The 20-μL reverse transcription reaction mixture contained 5 μL total RNA (1 μg), 1 μL 10 mM dNTP, 1 μL random primer (9-mer) (50 mM), 1 μL oligo-d(T)₁₈ primer (50 mM) (TaKaRa, Japan), and 6 μL DEPC water. The quantitative real-time PCR (qPCR) was completed using qPCR SYBR^® Premix Ex Taq^™ II (Tli RNaseH Plus) (TaKaRa) and the 7500 Real-Time PCR System (Applied Biosystems, USA). The qPCR primers used to validate the differentially expressed genes were synthesized by Beijing Liuhe Genomics Technology Co., Ltd. (Table 1). Each gene was analyzed using four replicates, after which the average threshold cycle was calculated for each sample. An endogenous actin gene was used for data normalization [36]. Relative fold changes in gene expression were calculated using the 2^−ΔΔCt method.

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Table 1. Primers used for quantitative real-time PCR.

https://doi.org/10.1371/journal.pone.0196537.t001

Statistical analysis

Results are expressed here as mean ± SD. The differences among means were evaluated using Tukey’s multiple comparison test and the statistical significance was set at p < 0.05. Principal component analysis (PCA) was carried out using SAS 8.0 (SAS Institute, Cary, NC, USA).

Results

Analysis of organic acid content in developing C. humilis and C. glandulosa fruits

The C. humilis and C. glandulosa fruits reached maturity at 19 and 14 weeks after flowering (WAF), respectively (Fig 1). Malic acid and citric acid were the main organic acids in their fruits. Similar trends were observed for the total acid, malic acid, and citric acid content of C. humilis fruits, with an increase during the early fruit-developmental stage, followed by a rapid increase during the fruit-enlargement stage (FES), and then a decrease in the mature fruits. The highest total acid (28.62±1.05 mg/g), malic acid (23.81±1.20 mg/g), and citric acid (3.53±0.23 mg/g) content were observed at 18 WAF. In mature fruits, the total acid, malic acid, and citric acid content decreased to 22.58±1.07 mg/g, 19.27±0.98 mg/g, and 2.48±0.21 mg/g, respectively (Table 2). Moreover, the malic acid and citric acid content of mature fruits accounted for 85.34% and 10.98% of the total acid content, respectively.

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Fig 1. C. humilis (A: 8, 10, 12, 14, 16, 18, and 19 weeks after flowering) and C. glandulosa (B: 8, 10, 12, 13, and 14 weeks after flowering) fruits at different developmental stages.

https://doi.org/10.1371/journal.pone.0196537.g001

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Table 2. Changes to total acid, malic acid, and citric acid contents in C. humilis and C. glandulosa fruits during development.

https://doi.org/10.1371/journal.pone.0196537.t002

Similar trends among the total acid, malic acid, and citric acid content were also observed for C. glandulosa fruits. The total acid and citric acid content decreased throughout the fruit development period, while the malic acid content increased slightly before fruit maturation. Total acid (17.59±0.88 mg/g), malic acid (14.13±0.33 mg/g), and citric acid (3.15±0.18 mg/g) content were highest at 8 WAF. The total acid, malic acid, and citric acid content decreased to 7.39 mg/g, 6.57 mg/g, and 0.35 mg/g in mature fruits, respectively. The lowest malic acid content (6.12 mg/g) was recorded at 13 WAF (Table 2). In mature C. glandulosa fruits, malic acid and citric acid accounted for 88.90% and 4.74% of the total acid content, respectively.

Analysis of organic acid-related enzyme activities in developing C. humilis and C. glandulosa fruits

Phosphoenolpyruvate carboxylase (PEPC) catalyzes the β-carboxylation of PEP to produce oxaloacetic acid (OAA) and inorganic phosphoric acid. The OAA is reduced to malic acid in a reaction catalyzed by NAD-MDH [11]. The activity of PEPC in C. humilis fruits increased gradually from 8 to 14 WAF, then decreased slightly from 14 to 16 WAF, followed by a rapid increase to the maximum value of 70.91 U g⁻¹ FW min⁻¹ (Fig 2A). In contrast, in C. glandulosa fruits, the PEPC activity was 72.69 U g⁻¹ FW min⁻¹ at 8 WAF and then decreased to 39.75 U g⁻¹ FW min⁻¹ at maturity (Fig 2A).

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Fig 2. Changes in PEPC (A), NAD-MDH (B), and NADP-ME (C) activities in C. humilis and C. glandulosa during fruit development.

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NAD-MDH activity exhibited the opposite trend in the developing fruits of C. humilis and C. glandulosa (Fig 2B). In C. humilis, NAD-MDH activity at 8 WAF was 82.43 U g⁻¹ FW min⁻1, which then gradually increased to 214.32 U g⁻¹ FW min⁻¹ as the fruits developed. In C. glandulosa, NAD-MDH activity was 192.69 U g⁻¹ FW min⁻¹ at 8 WAF, which then gradually decreased to the minimum value of 76.46 U g⁻¹ FW min⁻¹ at 13 WAF, after which the activity slowly increased to 91.25 U g⁻¹ FW min⁻¹ in mature fruits.

In developing C. humilis fruits, the activity of NADP-ME exhibited a decreasing trend from 8 to 18 WAF and then increased as the fruits matured (Fig 2C). Meanwhile, NADP-ME activity in C. glandulosa fruits gradually increased from 8 to 13 WAF, followed by a slight decrease as the fruits matured.

Transcriptome sequencing and identification of differentially expressed genes

We obtained 38,969 unigenes following the transcriptome sequencing of C. glandulosa and C. humilis (S1 Table), including 6,594 differentially expressed genes. There were 3,469 up-regulated and 3,125 down-regulated genes in C. glandulosa compared with that in C. humilis, including 12 genes directly involved in malic acid metabolism (CL4433.Contig2_All, Unigene8844_All, CL188.Contig2_All, Unigene6244_All, Unigene18125_All, CL471.Contig2_All, CL738.Contig1_All, CL3719.Contig1_All, CL4438.Contig1_All, Unigene5127_All, CL4732.Contig2_All, and Unigene12965_All), 25 genes directly involved in the citric acid cycle, 194 genes involved in NAD/NADP metabolism, and one PEPC gene (CL2688.Contig3_All) (S2 Table).

The above-mentioned 223 genes involved in malic acid metabolism and citric acid metabolism were classified into GO categories (Fig 3). There were 16, 10, and 12 subclasses in the biological process, molecular function, and cellular component categories, respectively. In the biological process category, the metabolic process (211 genes), single-organism process (204 genes), and cellular process (153 genes) subgroups contained the most genes, while the rhythmic process (one gene) contained the fewest genes. In the molecular function category, the catalytic activity (202 genes) and binding (151 genes) subgroups contained the most genes, while the nucleic acid binding transcription factor activity, antioxidant activity, and molecular function regulator subgroups contained the fewest genes, each with only three genes. In the cellular component category, the cell part and cell subgroups included the most genes, both with 168 genes, while the supramolecular fiber subgroup contained only one gene.

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Fig 3. Gene Ontology classifications of the differentially expressed genes related to malic acid and citric acid metabolism.

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The KEGG pathway analysis indicated that 233 differentially expressed genes were involved in 10 metabolic pathways, with the carbohydrate metabolism (42 genes), global and overview (37 genes), and energy metabolism (26 genes) pathways associated with the most genes. The pathways with the fewest genes were the metabolism of other amino acids (four genes) and nucleotide metabolism (two genes). The genetic information processing group contained folding, sorting, and degradation, as well as translation and transcription metabolic pathways. The environmental information processing group comprised signal transduction and membrane transport metabolic pathways. The cellular processes group contained transport and catabolism metabolic pathways, while the organismal systems group included environmental adaptation metabolic pathways (Fig 4).

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Fig 4. KEGG pathway classifications of the differentially expressed genes related to malic acid and citric acid metabolism.

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Expression analysis of genes encoding key enzymes for malic acid metabolism

Mature C. humilis and C. glandulosa fruits contained more malic acid than citric acid (Table 2), implying that malic acid is the main organic acid responsible for the differences in taste. The expression levels of genes encoding key enzymes involved in organic acid metabolism during different fruit-developmental stages were analyzed based on the transcriptome sequencing results and homologous sequences in the NCBI database.

In C. humilis fruits, the PEPC gene expression level increased slightly from the young fruit stage (YFS) to the pit-hardening stage (PHS), followed by a rapid increase from the PHS to the fruit-enlargement stage (FES) (Fig 5A). The highest and lowest PEPC gene expression levels were observed at the FES and the color changing stage (CCS), respectively. The PEPC gene had a similar expression trend in C. glandulosa fruits. The PEPC gene expression level at the FES was much lower in C. glandulosa fruits than in C. humilis fruits. In contrast, at all other fruit-development stages, the PEPC gene expression level was slightly higher in C. glandulosa fruits than in C. humilis fruits. These results implied that the PEPC gene affected the accumulation of organic acids in C. humilis fruits primarily at the FES (Fig 5A).

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Fig 5. Changes in the PEPC (A), NAD-MDH (B), NADP-ME (C) and CS (D) gene expression levels in C. humilis and C. glandulosa during fruit development.

YFS: young fruit stage; PHS: pit-hardening stage; FES: fruit-enlargement stage; CCS: color changing stage; FRS: fruit-ripening stage.

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In C. humilis fruits, the expression of the NAD-MDH gene decreased slightly from the YFS to the PHS, and then increased rapidly to reach the maximum value at the FES, which was followed by a rapid decrease at the CCS and a drastic increase at the FRS (Fig 5B). The NAD-MDH gene was highly expressed at the FES and at the FRS, with expression levels that were 5.58 and 7.42 times that of fruits at the YFS, respectively. The lowest expression level observed at the PHS was 0.81 times that of fruits at the YFS. Meanwhile, the highest NAD-MDH gene expression in C. glandulosa fruits was found at the FRS, which was 9.05 times that of fruits at the YFS, and the lowest NAD-MDH gene expression was observed at the PHS (Fig 5B). The NAD-MDH gene expression level was higher in C. humilis fruits than in C. glandulosa fruits throughout the fruit-developmental stage, but there was relatively little difference prior to the FES.

In C. humilis fruits, the NADP-ME gene expression level was the highest at the FES (2.17 times that of fruits at the YFS) and the lowest at the PHS (0.83 times that of fruits at the YFS). Meanwhile, in C. glandulosa fruits, the NADP-ME gene expression level was the highest at the FES (4.55 times that of fruits at the YFS) and the lowest at the YFS. Overall, the NADP-ME gene was more highly expressed in C. glandulosa fruits than in C. humilis fruits, except at the CCS. The relatively high NADP-ME gene expression levels induced the degradation of malic acid, especially at the FES and at the FRS (Fig 5C). The Citrate synthase (CS) gene expression levels were similar in two species and the CS gene was more highly expressed in C. humilis fruits than in C. glandulosa fruits (Fig 5D).

Correlation analysis and principal component analysis

The correlation between gene expression, enzymatic activity and organic acid content in developing fruits of C. humilis and C. glandulosa was analyzed. Strong correlations are apparent among total acid, malic acid, and citric acid content in both C. humilis and C. glandulosa fruits (Fig 6). The p-value was the probability and was used to compare the confidence coefficient among the data groups in the the paired Tukey’s-test [37]. Significant correlation between data groups is indicated if p-value is less than 0.05 (p < 0.05). In this study, the levels of total acid, malic acid, and citric acid contents are all less than 0.05, indicating significant correlations. The correlation of organic acid contents with enzyme activities and the gene expression is also shown in Fig 6. PEPC and NAD-MDH are significantly positively correlated with the contents of organic acid while NADP-ME is significantly negatively correlated with the organic acid contents, PEPC and NAD-MDH in both C. humilis and C. glandulosa (Fig 6A and 6B).

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Fig 6. The correlation analysis of organic acid content, enzyme activity and the gene expression in developing fruits of C. humilis (A) and C. glandulosa (B).

** represents p< 0.01; * represents p< 0.05.

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In C. humilis fruits, the expression of PEPC gene is significantly positively correlated with organic acid contents and enzyme activity of NAD-MDH while it is significantly negatively correlated with the expression of the NADP-ME gene (Fig 6A). The expression of NAD-MDH gene is significantly positively correlated with total acid, malic acid and PEPC expression, but its correlations with citric acid content, PEPC activity, NAD-MDH activity and NADP-ME activity are not significant. The correlations between the expression of NADP-ME gene and other data are not significant which indicate that NADP-ME gene doesn’t have a decisive effect on the organic acid accumulation in the fruits of C. humilis. However, NADP-ME gene plays a very important role in the organic acid metabolism of C. glandulosa fruits. For example, the NADP-ME gene expression is significantly negtively correlated with the organic acid contents and enzyme activity of NAD-MDH.

The PCA results identified four components that explained 100% of the total variation in organic acid metabolism of C. humilis and C. glandulosa (Table 3). The first principal component (PC1) contributed 99.65% and 91.85% of the total variation in organic acid metabolism of C. humilis and C. glandulosa respectively, suggesting that the variation in PC1 could explain the differences in organic acid metabolism of these two species. In C. humilis, NADP-ME gene has the largest eigenvalue meaning that the NADP-ME gene plays the most important part in the organic acid metabolism of the fruits. In C. glandulosa, NAD-MDH gene and NADP-ME gene have the largest and the second largest eigenvalues respectively, suggesting that in addition to NAD-MDH gene, NADP-ME gene also has a substantial effect on the organic acid metabolism.

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Table 3. Eigenvectors and eigenvalues of the first four principal components grouping the factors attributing to the organic acid metabolism in C. humilis and C. glandulosa fruits.

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Discussion

To the best of our knowledge, we systematically report for the first time, the comparative study on the organic acid accumulation in fruits of C. humilis and C. glandulosa. In this study, the malic acid and citric acid content in C. humilis fruits accounted for 85.34% and 10.98% of the total acid content, respectively. In contrast, the malic acid and citric acid content in C. glandulosa fruits accounted for 88.90% and 4.74% of the total acid content, respectively. These results suggest that malic acid and citric acid are the main organic acids in both fruits, and likely affect fruit taste. The organic acid contents decreased in mature fruits, which is consistent with the results of earlier studies on nectarine, pear, and apple [6,7,18]. Additionally, the organic acid content decreased continuously throughout the fruit-developmental stages of C. glandulosa. However, in the developing fruits of C. humilis, the organic acid content increased gradually until one week before maturation. Although it underwent a sharp decrease in the fruit-ripening stage, the organic acid content in the fruits of C. humilis was still high, making the fruits very sour.

In our study, 6,594 differentially expressed genes were detected based on transcriptome sequencing results, including 12 genes involved in malic acid metabolism, 25 genes involved in the citric acid cycle, 194 genes related to NAD/NADP metabolism, and one PEPC gene. These genes may be involved in the biosynthesis or catabolism of organic acids in C. humilis and C. glandulosa fruits, with potential consequences for fruit quality. There are at present relatively few reports describing the transcriptional regulation and cloning of genes regulating organic acid contents in C. humilis and C. glandulosa fruits. Therefore, the differences in gene expression levels described herein may be useful for future studies.

Through correlation analysis, we discovered strong correlations between organic acid content, enzyme activity and gene expression in both C. humilis and C. glandulosa fruits. First of all, PEPC, NAD-MDH and NADP-ME are significantly correlated with the organic acid contents in both C. humilis and C. glandulosa. In addition, NADP-ME gene doesn’t have a decisive effect on the organic acid accumulation in C. humilis while it plays a very important role in the organic acid metabolism of C. glandulosa. The PCA analysis was carried out and the results verified the important role of NADP-ME gene in the organic acid metabolism in C. glandulosa.

Our results suggest that NAD-MDH, NADP-ME, and PEPC are all involved in regulating organic acid biosynthesis in C. humilis and C. glandulosa fruits. However, the gene expression levels were not directly related to changes in malic acid content.

Supporting information

S1 Table. List of all unigenes obtained through transcriptome sequencing of Cerasus humilis and Cerasus glandulosa.

https://doi.org/10.1371/journal.pone.0196537.s001

(XLS)

S2 Table. List of differentially expressed unigenes between Cerasus humilis and Cerasus glandulosa which were related to organic acid metabolism.

https://doi.org/10.1371/journal.pone.0196537.s002

(XLS)

Acknowledgments

The authors would like to acknowledge Ms. Luting Jia, Ms. Xuanxuan Han and Dr. Zhifeng Chen who provided invaluable assistance in the samples collection and picture preparation. Dr. Juan Wang provided critical comments on an earlier version of the manuscript.

References

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- View Article
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- View Article
- Google Scholar
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- View Article
- Google Scholar
28. Mu XP, Aryal N, Du JM, Du JJ. Oil Content and Fatty Acid Composition of the Kernels of 31 Different Cultivars of Chinese Dwarf Cherry [Cerasus humilis (Bge.) Sok.]. J. Hortic. Sci. Biotech. 2015, 90: 525–529.
- View Article
- Google Scholar
29. Wang P, Xue X, Mu X, Zhang J, Cao Q, Du J. Analysis of Organic Acid Accumulation Characteristics and Organic Acid-Metabolizing Enzyme Activities of Chinese Dwarf Cherry (Cerasus humilis Bunge) Fruit. Scientia Agricultura Sinica. 2013, 46 (19): 4101–4109.
- View Article
- Google Scholar
30. Nergiz C, Ergönül PG. Organic acid content and composition of the olive fruits during ripening and its relationship with oil and sugar. Sci. Hortic. 2009, 122 (2): 216–220.
- View Article
- Google Scholar
31. Hirai M, Ueno I. Development of citrus fruits: Fruit development and enzymatic changes in juice vesicle tissue. Plant & Cell Physiology, 1977, 18 (4): 791–799.
- View Article
- Google Scholar
32. Grabherr MG, Haas BJ, Yassour M, Levin JZ, Thompson DA, Amit I, et al. Trinity: reconstructing a full-length transcriptome without a genome from RNA-Seq data. Nat. Biotechnol. 2011, 29 (7): 644–65.
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View Article
PubMed/NCBI
Google Scholar

[2] View Article

[3] PubMed/NCBI

[4] Google Scholar

[ref2] 2. Saradhuldhat P, Paull RE. Pineapple organic acid metabolism and accumulation during fruit development. Sci. Hortic. 2007, 112 (3): 297–303.
View Article
Google Scholar

[6] View Article

[7] Google Scholar

[ref3] 3. Yen C, Koch KE. Developmental changes in translocation and localization of ¹⁴C-labeled assimilates in grapefruit: Light and dark CO₂ fixation by leaves and fruit. Amer. Soc. Hort. Sci. 1990, 115 (5): 815–819.
View Article
Google Scholar

[9] View Article

[10] Google Scholar

[ref4] 4. Guo RZ, Guo WL, Li XY, Kou XH. Changes of organic acid contents and relative enzyme activities during the development of Huangguan pear fruit. Jiangsu Journal of agriculture science. 2013, 29 (1): 157–161.
View Article
Google Scholar

[12] View Article

[13] Google Scholar

[ref5] 5. Wang P, Xue X, Mu X, Zhang J, Cao Q, Du J. Analysis of Organic Acid Accumulation Characteristics and Organic Acid-Metabolizing Enzyme Activities of Chinese Dwarf Cherry (Cerasus humilis Bunge) Fruit. Scientia Agricultura Sinica. 2013, 46 (19): 4101–4109.
View Article
Google Scholar

[15] View Article

[16] Google Scholar

[ref6] 6. Zhang Y, Li P, Cheng L. Developmental changes of carbohydrates, organic acids, amino acids, and phenolic compounds in ‘Honeycrisp’ apple flesh. Food. Chem. 2010, 123 (4):1013–1018.
View Article
Google Scholar

[18] View Article

[19] Google Scholar

[ref7] 7. Huo YQ, Hu HJ, Peng SA, Chen QL. Contents and changes of organic acid in Sand pears from different germplasm resources. Scientia Agricultura Sinica, 2009, 42 (1): 216–223.
View Article
Google Scholar

[21] View Article

[22] Google Scholar

[ref8] 8. Chen MX, Zhao CK, Chen XS, Fang SM, Zhang XS. Relationship between accumulation of sugar and sucrose-metabolizing enzymes during apricot fruit development. Journal of Fruit Science. 2009, 02: 471–474.
View Article
Google Scholar

[24] View Article

[25] Google Scholar

[ref9] 9. Chollet R, Vidal J, O’Leary MH. Phosphoenolpyruvate carboxylase: A ubiquitous, highly regulated enzyme in plants. Annu. Rev. Plant. Phys. 1996, 47: 273–298.
View Article
Google Scholar

[27] View Article

[28] Google Scholar

[ref10] 10. Notton BA, Blanke MM. Phosphoenolpyruvate carboxylase in avocado fruit: purification and properties. Phytochemistry. 1993, 33:1333–1337.
View Article
Google Scholar

[30] View Article

[31] Google Scholar

[ref11] 11. Miller SS, Driscoll BT, Gregerson RG, Gantt JS, Vance CP. Alfalfa malate dehydrogenase (MDH): molecular cloning and characterization of five different forms reveals a unique nodule enhance MDH. Plant J. 1998, 15: 173–184.
View Article
Google Scholar

[33] View Article

[34] Google Scholar

[ref12] 12. Scheibe R. Malate valves to balance cellular energy supply. Physiologia Plantarum. 2004, 120: 21–26. pmid:15032873
View Article
PubMed/NCBI
Google Scholar

[36] View Article

[37] PubMed/NCBI

[38] Google Scholar

[ref13] 13. Wen T, Xiong QE, Zeng WG, Liu PY. Changes of Organic Acid Synthetase Activity during Fruit Development of Navel Orange (Citrus sinesis Osbeck). Acta Horticulturae Sinica. 2001, 28 (2):161–163.
View Article
Google Scholar

[40] View Article

[41] Google Scholar

[ref14] 14. Moing A, Rothan C, Svanella L, Just D, Diakou P, Raymond P, et al. Role of phosphoenolpyruvate carboxylase in organic acid accumulation during peach fruit development. Physiol. Plant. 2000, 108: 1–10.
View Article
Google Scholar

[43] View Article

[44] Google Scholar

[ref15] 15. Moing A, Svanella L, Monet R, Rothan C, Just D, Diakou P, et al. Organic acid metabolism during the fruit development of two peach cultivars. Acta Horticulturae. 1998, 465: 425–432.
View Article
Google Scholar

[46] View Article

[47] Google Scholar

[ref16] 16. Law RD, Plaxton WC. Purification and Characterization of a novel Phosphoenolpyruvate carboxylase from banana fruit. Biochem. J. 1995, 307: 807–816. pmid:7741712
View Article
PubMed/NCBI
Google Scholar

[49] View Article

[50] PubMed/NCBI

[51] Google Scholar

[ref17] 17. Diakou P, Svanella L, Raymond P, Gaudillère JP, Moing A. Phosphoenolpyruvate carboxylase during grape berry development: protein level, enzyme activity and regulation. Aust. J. Plant. Physiol. 2000, 27: 221–229.
View Article
Google Scholar

[53] View Article

[54] Google Scholar

[ref18] 18. Zhao YH, Li XL, Jiang ZS, Wang CJ, Yang FL. Organic acid metabolism in nectarine fruit development under protected cultivation.China Journal of Eco-Agriculture. 2007, 15 (5): 87–89.
View Article
Google Scholar

[56] View Article

[57] Google Scholar

[ref19] 19. Etienne A, Moing A, Dirlewanger E, Raymond P, Monet R, Rothan C. Isolation and characterization of six peach cDNAs encoding key proteins in organic acid metabolism and solute accumulation: involvement in regulating peach fruit acidity. Physiologa Plantarum. 2002, 114: 259–270.
View Article
Google Scholar

[59] View Article

[60] Google Scholar

[ref20] 20. Yao YX, Li M, Liu Z, You CX, Wang DM, Zhai H, et al. Molecular cloning of three malic acid related genes MdPEPC, MdVHA-A, MdcyME and their expression analysis in apple fruits. Sci. Hortic. 2009, 122 (3): 404–408.
View Article
Google Scholar

[62] View Article

[63] Google Scholar

[ref21] 21. Or E, Baybik J, Sadka A, Saks Y. Isolation of mitochondrial malate dehydrogenase and phosphoenolpyruvate carboxylase cDNA clones from grape berries and analysis of their expression pattern throughout berry development. J. Plant. Physiol. 2000, 157: 527–534.
View Article
Google Scholar

[65] View Article

[66] Google Scholar

[ref22] 22. Palomo J, Gallardo F, Suarez MF, Canovas FM. Purification and characterization of NADP⁺-linked isocitrate dehydrogenase from Scots pine. Evidence for different physiological roles of the enzyme in primary development, Plant. Physiol. 1998, 118: 617–626. pmid:9765548
View Article
PubMed/NCBI
Google Scholar

[68] View Article

[69] PubMed/NCBI

[70] Google Scholar

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View Article
PubMed/NCBI
Google Scholar

[72] View Article

[73] PubMed/NCBI

[74] Google Scholar

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View Article
Google Scholar

[76] View Article

[77] Google Scholar

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View Article
Google Scholar

[79] View Article

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View Article
PubMed/NCBI
Google Scholar

[82] View Article

[83] PubMed/NCBI

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View Article
Google Scholar

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[87] Google Scholar

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View Article
Google Scholar

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[90] Google Scholar

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View Article
Google Scholar

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[93] Google Scholar

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View Article
Google Scholar

[95] View Article

[96] Google Scholar

[ref31] 31. Hirai M, Ueno I. Development of citrus fruits: Fruit development and enzymatic changes in juice vesicle tissue. Plant & Cell Physiology, 1977, 18 (4): 791–799.
View Article
Google Scholar

[98] View Article

[99] Google Scholar

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View Article
Google Scholar

[101] View Article

[102] Google Scholar

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View Article
PubMed/NCBI
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Google Scholar

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Google Scholar

[111] View Article

[112] Google Scholar

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View Article
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[115] PubMed/NCBI

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View Article
PubMed/NCBI
Google Scholar

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Figures

Abstract

Introduction

Materials and methods

Determination of total acid, malic acid, and citric acid contents

Determination of enzyme activity

RNA extraction

Library preparation for transcriptome sequencing

Clustering and sequencing

Transcriptome assembly and function annotation

Quantification of gene expression levels and identification of differentially expressed genes

Quantitative real-time PCR analysis

Statistical analysis

Results

Analysis of organic acid content in developing C. humilis and C. glandulosa fruits

Analysis of organic acid-related enzyme activities in developing C. humilis and C. glandulosa fruits

Transcriptome sequencing and identification of differentially expressed genes

Expression analysis of genes encoding key enzymes for malic acid metabolism

Correlation analysis and principal component analysis

Discussion

Supporting information

S1 Table. List of all unigenes obtained through transcriptome sequencing of Cerasus humilis and Cerasus glandulosa.

S2 Table. List of differentially expressed unigenes between Cerasus humilis and Cerasus glandulosa which were related to organic acid metabolism.

Acknowledgments

References