Next Article in Journal
The Impact of Klotho in Cancer: From Development and Progression to Therapeutic Potential
Previous Article in Journal
Integration of GWAS and Co-Expression Network Analysis Identified Main Genes Responsible for Nitrogen Uptake Traits in Seedling Waxy Corn
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Genes
Volume 16
Issue 2
10.3390/genes16020127
genes-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Genome-Wide Identification and Characterization of Histone Acetyltransferases and Deacetylases in Cucumber, and Their Implication in Developmental Processes

by
Agnieszka Skarzyńska-Łyżwa
,
Szymon Turek
,
Maksymilian Pisz
,
Aparna
,
Wojciech Pląder
and
Magdalena Pawełkowicz
*
Department of Plant Genetics, Breeding and Biotechnology, Institute of Biology, Warsaw University of Life Sciences-SGGW, 159 Nowoursynowska Str., 02-776 Warsaw, Poland
*
Author to whom correspondence should be addressed.
Genes 2025, 16(2), 127; https://doi.org/10.3390/genes16020127
Submission received: 13 December 2024 / Revised: 18 January 2025 / Accepted: 21 January 2025 / Published: 23 January 2025
(This article belongs to the Section Plant Genetics and Genomics)
Browse Figures
Versions Notes

Abstract

:
Background/Objectives: Cucumber (Cucumis sativus) provides a model for exploring the molecular basis of sex determination, particularly the regulation of floral organ differentiation through gene expression. This complex process is modulated by epigenetic factors, such as histone acetyltransferases (HATs) and histone deacetylases (HDACs), which respectively activate and repress gene transcription by adding or removing acetyl groups from histone proteins. Despite their known functions, the roles of HATs and HDACs throughout cucumber’s floral developmental stages remain unclear. Methods: In this study, we conducted a genome-wide analysis of HAT and HDAC gene families in cucumber, examining their phylogenetic relationships, gene structures, protein domains, and expression profiles across various stages of floral development. Results: We identified 36 CsHAT and 12 CsHDAC genes, grouping them into families with evolutionary counterparts in other plant species. RNA sequencing revealed stage-specific expression patterns, suggesting dynamic roles for these gene families in floral organ development. Conclusions: These findings contribute valuable insights into the epigenetic regulation of gene expression in cucumber flower formation, presenting avenues for further research on the genetic control of plant reproductive development.

1. Introduction

Chromatin, a highly intricate and organized structure, is composed of densely packed DNA molecules within eukaryotic nuclei. The fundamental unit of chromatin is the nucleosome, which consists of approximately 146 base pairs of DNA intricately wound around an octamer of histone proteins [1]. This octamer comprises four distinct histone subunits, namely H2A, H2B, H3, and H4. Post-translational modifications (PTMs) occur on specific amino acid residues located on the unstructured tails of these histone proteins [2]. These PTMs, such as acetylation, methylation, phosphorylation, ubiquitination, ADP-ribosylation, and glycosylation, play pivotal roles in epigenetic regulation of gene expression [3,4].
Among these modifications, acetylation stands out as one of the extensively researched mechanisms. Notably, lysine residues in the histone proteins H3 and H4 are frequently targeted for acetylation [5,6]. It is important to note that epigenetic modifications do not involve changes in the underlying DNA sequence itself. Instead, they primarily involve modifications of histone proteins and DNA methylation. These epigenetic modifications exert profound influences on gene expression and are instrumental in facilitating adaptive responses in plants, enabling them to thrive in diverse developmental phases and under challenging environmental conditions [7,8].
The acetylation process involves combining two different counteracting enzymes, histone acetyltransferases (HATs) and histone deacetylases (HDACs). Both enzymes work in a reversible manner [9]. HATs, through their enzymatic action, facilitate the loosening of the interactions between DNA and histones. This enzymatic activity results in the relaxation of chromatin structure, rendering it more accessible for gene expression [10]. This chromatin relaxation phenomenon is instrumental in promoting gene transcription, as it frees the DNA from its tightly wound configuration, allowing transcriptional machinery to bind more effectively. Conversely, HDACs catalyze the removal of acetyl groups from histone proteins. This deacetylation process leads to the condensation of the chromatin structure, making it more compact. In addition to promoting chromatin condensation, HDACs also exert an inhibitory effect on gene transcription [11].
Both HATs and HDACs can be categorized into distinct families. In plants, HATs contain four families such as HAG of the GNAT (GCN5-related N-terminal acetyltransferases) superfamily, HAC of the p300/CBP (CREB-binding protein) family, HAM of the MYST (Moz, ybf2/saa2 and Tip60) superfamily, and HAF of the TAFII 250 (TATA-binding protein-associated factors) family, while HDACs are divided into three families such as RPD3/HDA1 (Reduced Potassium Dependency 3/Histone Deacetylase 1), SIR2 (Silent Information Regulator 2), and HD2 (Histone Deacetylase 2) [12,13].
HATs and HDACs perform crucial roles in the developmental processes of plants, including flowering timings and regulation of seed, flower, leaf, and root growth, as well as fruit development and ripening-related phytohormone metabolism and plant responses to biotic and abiotic environmental stresses [14,15].
This study aimed to identify and subsequently analyze histone modifiers (CsHATs and CsHDACs) found in the genome of the B10 line of cucumber. A comprehensive analysis of the structure, phylogeny, and composition of this family is presented in this research. We also identified the expression levels of different HATs genes (CsHATs) and HDACs genes (CsHDACs) at various developmental stages of the studied cucumis B10 line.

2. Materials and Methods

2.1. Identification of Histone Acetyltransferases and Deacetylase Protein Sequences

Protein sequences of cucumber acetyltransferases and histone deacetylases were identified by searching the B10v3 genome annotation, as established and published by Osipowski and coauthors [16].
Additionally, a BLASTP [17] analysis was performed on the B10 proteins against related cucumber proteomes, in particular the 9930 v3 line [18] and the Gy14 line [19]. These searches yielded a consolidated list of shared proteins across the three cucumber lines, focusing on those annotated as histone acetyltransferases and deacetylases [12,20]. The InterProScan program [21] was used to analyze the protein sequences of the B10v3. All databases available in the local version of the program were utilized, consisting of TIGRFAM, SFLD, SUPERFAMILY, PANTHER, Gene3D, Hamap, ProSiteProfile, Coils, SMART, CDD, PRINTS, PIRSR, ProSitePattern, AntiFam, Pfam, MobiDBLite, and PIRSF. This comprehensive search expanded the HATs list by identifying additional proteins associated with characteristic domains or families, such as GNAT, MYST, and CBP.
Subsequently, the results from InterProScan were used to refine the classification. Proteins were narrowed down to those containing the specific domains relevant to this category, ensuring a more targeted and accurate dataset.
Using the identified protein domains and the family classifications obtained from the Pfam and Superfamily programs, HAT proteins were categorized into the HAG, HAC, HAF, and HAM groups. This approach enabled the classification of cucumber HDACs into three distinct groups: HDA, SRT, and HDT, reflecting their sequence similarity and domain characteristics.

2.2. Multiple Sequence Alignment and Phylogenetic Analysis

The amino acid sequences of the HAT and HDAC genes were aligned using Clustal W 2.1 [22]. The neighbor-joining method was employed with 1000 replications of bootstrapping and partial deletion. Selected histone acetyltransferase and deacetylase sequences from cucumber, Arabidopsis thaliana, and Solanum lycopersicum were used to perform the MSA. In the next step, the IQ-TREE server [23] was used to construct a phylogenetic tree based on the sequence alignment file. Finally, the iTOL server [24] was used to visualize the resulting tree and present it in a suitable format. This process was then repeated to compare previously identified domain sequences in the same way.

2.3. Protein Motif Analysis and Determination of Physicochemical Properties of CsHATs and CsHDACs

The conserved motifs identification was performed with MEME [25]. For CsHATs, the number of searched motifs was set to 30, while for CsHDACs, this value was set to 20. The results of the motifs search were visualized with TBtools. The amino acid length, molecular weight (Mw), and isoelectric point (pI) of the identified CsHATs and CsHDACs proteins were estimated using the EMBOSS PEPSTATS tool [26]. Additionally, the WoLF PSORT online tool [27] was used to predict the sub-cellular localization of the CsHATs and CsHDACs proteins.

2.4. Chromosomal Localization and Gene Structure

Chromosomal localization was performed based on the B10v3 cucumber genome. Visualization was performed with the circlize [28] package for R with a custom script. Gene structure information was obtained from B10v3 genome annotation [16,29] and visualized using TBtools [30].

2.5. Expression Analysis of CsHATs and CsHDACs

The RNA-seq data analyzed in this study was sourced from the SRA database under the identifier PRJNA1166086. Reads were aligned to the B10v3 cucumber genome sequence (GenBank: LKUO00000000.3). Gene expression levels were quantified using the Salmon software [31], with sequence-specific and GC content bias correction. Normalization was performed to Transcripts Per Million (TPM), and differential expression analysis was conducted using the Limma package [32]. Genes identified as acetyltransferases and deacetylases were extracted from the TPM matrix, and a heatmap of their expression profiles was generated with the gplots package [33].
Differential gene expression results were filtered for genes of interest (acetyltransferases and deacetylases) using an adjusted p-value threshold of 0.05. Venn diagrams were employed to visualize differentially expressed genes (DEGs) across floral bud growth stages, including comparisons such as 1–2 mm vs. shoot apex, 3–5 mm vs. 1–2 mm, 6–8 mm vs. 3–5 mm, 9 mm vs. 6–8 mm, and comparison of each developmental stage relative to the leaf stage.

2.6. Cis-Acting Elements in Promotor Regions

To identify the cis-acting elements in the promoter region of CsHATs and CsHDACs genes, the upstream sequences of 1000 bp were extracted. The sequences were analyzed using the PlantCARE database [34] to predict cis-regulatory elements. The results were then organized and visualized using TBtools.

3. Results

3.1. Identification of CsHATs and CsHDACs in C. sativus

Analysis of the B10v3 cucumber database led to the identification of 36 CsHATs and 12 CsHDACs. Among the CsHATs, 28 of the proteins were classified as belonging to the HAG family, seven of them were assigned to the HAC family, and a single protein was assigned to the HAM family. None of the identified CsHAT proteins was found to be a part of the HAF family. For the CsHDACs, nine proteins were assigned to the HDA family, three proteins were identified in the HDT family, and no protein was found in the SRT family. Additionally, one CsHAT and one CsHDAC were found only in the 9930 line. The obtained results were compared with existing protein distribution data for A. thaliana and S. lycopersicum and compiled in Table 1.

3.2. Comparative Phylogenetic Analysis

We compared the 36 CsHAT proteins from cucumber that were identified in this study with 12 proteins previously identified in A. thaliana [12] and 29 proteins identified in the S. lycopersicum [20]. To investigate the protein relationships, all described HAT proteins were used to construct the phylogenetic tree based on the neighbor-joining method (Figure 1). Proteins that were classified as belonging to the same family (HAG, HAM, HAC, or HAF) were mostly grouped together on the same branch of the tree.
We performed the same analysis for HDACs proteins—12 proteins identified in the cucumber, 15 proteins from A. thaliana, and 12 proteins from S. lycopersicum. All the sequences were used to prepare a neighbor-joining phylogenetic tree (Figure 2). Proteins assigned to the same family (HDA, HDT, or SRT) were grouped together on the same branch, except for two HDA proteins (one from tomato and one from cucumber) that were grouped together with the HDT family.
Based on the InterProScan protein domain identification (Supplementary Table S1), we constructed the phylogenetic trees using only domain sequences specific to each protein. We used the neighbor-joining method. We used 77 sequences for HATs and 39 sequences for HDACs. The tree obtained for the HAT proteins showed similar clustering to the alignment prepared using whole protein sequences. The tree obtained for HDAC proteins also showed similar clustering. However, the domain alignment allowed for better grouping of the HDT family proteins, which were distributed on the same branch except for one protein from the cucumber. The resulting phylogenetic trees based on the specific domain sequences are presented in Figure S1 in the Supplementary Materials Section.

3.3. Characteristics of Motifs, Domains, Gene Structure of CsHATs and CsHDACs

For a better understanding of the structural diversity of the CsHAT and CsHDAC families, we identified and analyzed the conserved motifs using the MEME tool and analyzed the occurrence of conserved protein domains with InterProScan. We identified 30 conserved motifs in the CsHAT proteins, ranging from 7 aa to 50 aa in length, and 30 protein domains from the Pfam database (Figure 3). There is no clear pattern for each HAT class in terms of the predicted motifs. However, the proteins that clustered together on the same branches of the phylogenetic tree showed a similar motif distribution. For example, HAC family proteins G19498, G5059, and G13466 consist only of motif 5 and motif 10, which are related to the occurrence of the KIX domain, a coactivator of CBP, found only in these three proteins. Another group is the HAG family proteins G5634, G8133, G2255, G12163, and G1301, which showed a similar composition of conserved motifs, which are motifs 1–4, 8–9, 14, and 19. Motif 1 was found in 24 CsHAT proteins. This motif is most likely related to the occurrence of the acetyltransferase 1 domain.
Through analysis of the conserved motifs in the CsHDAC proteins, we identified 21 motifs ranging from 9 aa to 50 aa in length and seven domains from the Pfam database (Figure 4). Motifs 7, 10, and 19 are specific to the HDT protein family. They are associated with the presence of nucleoplasmin-like domain (NPL). The HDA proteins showed a similar composition of conserved motifs. Each of them has motifs 1, 2, 4, 5, and 14 (with the exception of protein G11372, which lacked motifs 5 and 14), which are most likely related to the histone deacetylase domain found in almost all protein sequences analyzed.
The gene structure analysis was conducted based on the cucumber B10v3 genome annotation (Supplementary Table S2). The CsHAT genes varied in length (the shortest transcript consisted of 406 bp, while the longest consisted of 15,795 bp) and number of exons, which ranged from 2 to 18 exons. Most genes (32 out of 36) have both untranslated regions (UTRs). The CsHDAC genes also differed in length (the shortest transcript consisted of 2932 bp, while the longest consisted of 20,916 bp), but less significantly than CsHATs. The number of exons also varied from 3 to 18 exons. All genes have both UTRs.

3.4. Physicochemical Properties of CsHATs and CsHDACs

Physicochemical analysis and subcellular localization of the identified proteins were performed using the EMBOSS PEPSTATS and WoLF PSORT tools, respectively (Table 2). The proteins belonging to the CsHDAC family ranged in length from 296 aa to 659 aa and in molecular weight from 31.7 kD to 73.2 kD. The theoretical isoelectric point (pI) of the CsHDACs exhibited a range of values, from 4.4 to 7.5. Proteins of the CsHAT family showed a greater variety of parameters, with lengths from 113 aa to 1707 aa and molecular weights ranging from 12.5 kD to 192.6 kD. The theoretical isoelectric point (pI) of CsHATs ranged from 4.8 to 9.9. The complete results of the analysis are included in Supplementary Table S3. In terms of subcellular localization, the majority of CsHDACs were identified in the nucleus, as well as in the cytoplasm and chloroplast, whereas CsHATs were mainly located in the nucleus and cytoplasm.

3.5. Chromosomal Location of CsHDAC and CsHAT Genes

The genome annotation information (Table 2) was used to create a chromosome map of the CsHDACs and CsHATs, with all 48 members showing specific locations (Figure 5). The chromosomal distribution of 12 CsHDACs was found to be irregular, with the genes present on chromosomes 1, 3, 4, 6, and 7. The largest number—four CsHDAC genes—is present on chromosomes 1 and 6. Two CsHDACs are present on chromosome 4, while one gene each is present on chromosomes 3 and 7. The 36 CsHAT members were distributed on each chromosome. Seven genes are distributed on chromosome 3, six each on chromosomes 1 and 7, five CsHAT genes are present on chromosomes 2 and 5, and three genes each on chromosomes 4 and 5.

3.6. Expresional Analysis

As a result of the analysis of the RNA-seq data for genes classified as acetyltransferases and deacetylases (Supplementary Table S4), heatmaps showing the expression profiles of these two groups during flower bud development were prepared. Samples from leaves, shoot apex, and flower buds at different developmental stages (1–2 mm, 3–5 mm, 6–8 mm, and 9 mm), in triplicate, were used in the analysis. Figure 6 presents the expression profile of CsHATs, while Figure 7 shows the expression profile of CsHDACs.
Visualizations of the expression profile revealed differences between the early stages of flower bud development (shoot apex and 1–2 mm) compared to the later stages (3–5 mm, 6–8 mm, 9 mm) for both acetyltransferases and deacetylases. The leaves represent a distinct group whose expression profile shows similarities to both the early and late stages of flower bud development.
Among the genes classified as acetyltransferases and deacetylases, only a subset exhibited statistically significant changes in expression. Specifically, 26 acetyltransferase genes showed significant changes, compared to five genes among the deacetylases. Venn diagrams (Figure 8) illustrate the overlap of differentially expressed genes (DEGs) across various comparisons for both CsHATs and CsHADCs together. The corresponding gene identifiers are listed in Supplementary Table S5. Notably, 13 CsHATs showed unique changes in expression only in the comparison between the 1–2 mm flower buds and the shoot apexes (Figure 8A). The highest number of CsHAT or CsHDAC genes with differential expression during flower bud development was identified in the comparison between the 1–2 mm stages and the apexes, totaling 17 genes, of which 13 were unique to this comparison.

3.7. Promoter Analysis

The analysis of the promoter region of genes—identification of the cis-acting elements, their quantity and arrangement, may indicate the function and regulatory mechanisms of genes. The core promoter elements like TATA-box and CAAT-box, represented on average 65% of all elements found in CsHAT genes and 55% in CsHDACs genes (Figure 9). Besides core elements, we found cis-acting elements involved in the light response (for example ACE, Box4, GATA-motif, G-box), dehydration responsiveness (like MYB binding site or DRE), abscisic acid responsiveness (like ABRE, MYB), salicylic acid responsiveness (like as-1), low-temperature responsiveness (MYC), heat stress responsiveness (HSE1), defense and stress responsiveness (like W-box), wounding response (box S), and also elements involved in meristem expression (CAT-box) and differentiation of the palisade mesophyll cells (HD-Zip 1). The detailed results are presented in Supplementary Table S6.
The analysis of the promoter regions of CsHATs revealed that the cis-acting elements involved in ABA responsiveness were the most abundant and accounted for 33.1% of all cis-elements found, excluding core promoter elements (Figure 10). The next most abundant groups were light responsiveness elements (22.1%) and low-temperature responsiveness elements (21.1%).
The analysis of the promoter regions of CsHDACs showed that cis-acting elements involved in ABA responsiveness were also most abundant in these genes (Figure 11). They accounted for 35.2% of all cis-elements found (excluding core promoter elements). The next highly represented groups were elements involved in low-temperature responsiveness (19.7% of all elements) and light responsiveness elements (16.4%) (Figure 12).

4. Discussion

Histone modification plays a critical role in plant growth and development. Histone acetyltransferases and deacetylases contribute significantly to various plant physiological and metabolic processes, including chromatin remodeling and transcription regulation. These processes are essential not only for general growth but also for regulating genes involved in determining reproductive structures and processes [35]. Importantly, histone modifications are also implicated in plant sex determination. For example, reprogramming histone H3 lysine methylation has been shown to affect the expression of genes critical for sex-specific development during gametogenesis. Such modifications can influence the differentiation of male and female gametophytes, ultimately guiding sex determination and reproduction in plants [36,37].
In this study on Cucumis sativus, 12 HDAC and 36 HAT proteins were identified. Among these, the HDA family was the most prominent in the CsHDACs, while the HAG family predominated in the CsHATs. This distribution pattern is consistent with results from other plant species such as Arabidopsis thaliana [12], Solanum lycopersicum [20], Triticum aestivum [38], and Vitis vinifera [39]. Both CsHDACs and CsHATs contain family-specific structural domains [36,37]. In addition, the family members exhibit similar protein sequence lengths, motif compositions, and gene structures, suggesting a close phylogenetic relationship.
The expression profiles of acetyltransferase and deacetylase genes during flower bud development, visualized through RNA-seq heatmaps, provide insights into the regulatory dynamics of these two groups of genes. The findings reveal distinct stage-specific gene expression patterns, emphasizing the dynamic role of histone-modifying enzymes in floral development. Acetyltransferase genes showed significant shifts in expression between early and late developmental stages. Their activity was distinct during the shoot apex and 1–2 mm flower bud stages, suggesting a potential role in initiating developmental programs. In contrast, later stages (3–5 mm, 6–8 mm, and 9 mm flower buds) displayed altered acetyltransferase expression, possibly supporting differentiation and maturation processes. Similarly, deacetylase genes showed pronounced differences between the early and late stages of flower bud development, reflecting their involvement in repressing or fine-tuning gene expression as the flower bud progresses through development. It is worth mentioning that previous studies on cucumber flower bud development indicate that at early stages of development when flower buds are about 1 mm long, the initiation of particular primordia growth and flower sex differentiation begins. In the later stages, the growth of primordia is continued [40].
Interestingly, leaf samples displayed an intermediate pattern of expression, sharing similarities with both the early and late stages of flower bud development. This suggests overlapping regulatory mechanisms or retention of developmental cues common to vegetative and floral tissues. The different expression profiles of acetyltransferases and deacetylases during flower bud development are consistent with the literature findings that highlight their key roles in regulating developmental processes. For instance, studies have demonstrated that histone acetyltransferases are crucial for chromatin remodeling and gene expression during plant growth. In Vitis vinifera, HATs have been shown to regulate the expression of genes associated with developmental transitions, indicating their role in organ differentiation [39]. Studies on HATs in Triticum aestivum also indicate the role of these genes in plant development [1].
Similarly, histone deacetylases have been implicated in stage-specific expression during plant development. Research on rice HDAC families revealed distinct expression patterns, correlating with specific developmental stages and stress responses. This emphasizes the dual role of HDAC in controlling gene silencing and developmental fine-tuning [41].
These examples provide further evidence of the functional importance of dynamic acetylation and deacetylation processes, supporting the observed stage-specific expression profiles in flower bud development. Such regulation ensures proper progression through developmental stages, balancing activation and repression of critical genetic pathways [14,42].
Recent studies have revealed the involvement of HDAC proteins: HDA6 and HDA5 in the regulation of flowering by the autonomic pathway [43]. In particular, hda6 mutants exhibit delayed flowering phenotypes in both long-day and short-day conditions, with their flowering time being dependent on the presence of FLC [44].
In bananas, HDA1 has been shown to interact with ERF11, a component of the ethylene signaling pathway, to delay fruit ripening. This delay also occurs through repression of the ethylene biosynthesis gene MaACO1 and the expansin genes [45]. Another histone deacetylase, MaHDA6, plays a role in promoting ethylene signaling and fruit ripening by inhibiting ERF11/15 genes [46]. Similarly, in tomatoes, different HDACs have different effects on fruit ripening. SlHDA3 acts as a ripening inhibitor by regulating ethylene biosynthesis [47], whereas SlHDT3 stimulates ethylene and carotenoid accumulation, contributing to fruit ripening [48]. In papayas, CpHDA3, an RPD3-type histone deacetylase, forms a repressor complex with ERF9 [49]. The involvement of HDACs in ethylene biosynthesis and signaling pathways is also important in cucumber floral sex determination, in which ethylene plays a key role as a major hormone [50].
It can be seen that in the 1000 bp of the analyzed promoter region, the most abundant motifs are the motifs common to CsHAT/CsHDAC, which are 105/43 (32/11 genes), 70/20 (29/11 genes), and 67/24 (30/12 genes), respectively for the processes of ABA responsiveness, light responsiveness, and low-temperature responsiveness. The above three processes are mainly related to cell physiology and concern many aspects of ABA use, including sex development [51,52].
These results highlight the complex interaction between acetylation and deacetylation during developmental transitions, which may have a real connection with ethylene signaling during cucumber flower morphogenesis. This provides a foundation for further research into their specific functional roles in floral organogenesis.

5. Conclusions

The role of histone modifications, particularly acetylation and deacetylation, is fundamental for plant growth, development, and reproduction. Histone acetyltransferases (HATs) and deacetylases (HDACs) coordinate critical regulatory processes through chromatin remodeling and transcriptional control. The stage-specific expression of CsHAT and CsHDAC genes during flower bud development underscores their dynamic involvement in initiating, fine-tuning, and completing developmental transitions. In Cucumis sativus, the identification of diverse HDAC and HAT families highlights conserved patterns and unique subfamily features that are consistent with other plant species. By modulating histone H3 lysine methylation and other epigenetic marks, they influence gametophyte differentiation and gene expression critical for male and female organogenesis. The results of RNAseq analysis showed that HATs and HDACs play an important role in flower bud development and growth. Overall, the interplay between histone acetylation and deacetylation ensures a balanced progression of gene expression programs, enabling plants to adapt their developmental pathways to internal and external signals. These findings provide a basis for deeper investigations into epigenetic mechanisms regulating floral organogenesis and sex determination, with potential implications for crop improvement and yield optimization.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/genes16020127/s1, Table S1: Results of the InterProScan search, Figure S1, Results of phylogenetic analysis for HAT and HDAC domains, Table S2: Gene structure analysis, Table S3: Results of EMBOSS PEPSTATS analysis, Table S4: Results of expressional analysis, Table S5: Venn diagram results tables, Table S6: Results of the PlantCARE promoter analysis.

Author Contributions

Conceptualization, M.P. (Magdalena Pawełkowicz) and A.; methodology, S.T. and M.P. (Maksymilian Pisz); formal analysis, S.T., M.P. (Maksymilian Pisz) and A.S.-Ł., writing—original draft preparation, M.P.; writing—review and editing, W.P.; visualization, A.S.-Ł., S.T. and M.P. (Maksymilian Pisz); supervision, M.P. (Magdalena Pawełkowicz); project administration, M.P. (Magdalena Pawełkowicz); funding acquisition, M.P. (Magdalena Pawełkowicz). All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by a project from the National Science Center UMO-2020/37/B/NZ9/00586.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

Publicly available datasets were analyzed in this study and are referenced in the cited articles.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Gao, S.; Li, L.; Han, X.; Liu, T.; Jin, P.; Cai, L.; Xu, M.; Zhang, T.; Zhang, F.; Chen, J.; et al. Genome-Wide Identification of the Histone Acetyltransferase Gene Family in Triticum aestivum. BMC Genom. 2021, 22, 49. [Google Scholar] [CrossRef]
  2. Xu, J.; Xu, H.; Liu, Y.; Wang, X.; Xu, Q.; Deng, X. Genome-Wide Identification of Sweet Orange (Citrus sinensis) Histone Modification Gene Families and Their Expression Analysis during the Fruit Development and Fruit-Blue Mold Infection Process. Front. Plant Sci. 2015, 6, 151116. [Google Scholar] [CrossRef]
  3. Berger, S.L. The Complex Language of Chromatin Regulation during Transcription. Nature 2007, 447, 407–412. [Google Scholar] [CrossRef] [PubMed]
  4. Kouzarides, T. Chromatin Modifications and Their Function. Cell 2007, 128, 693–705. [Google Scholar] [CrossRef] [PubMed]
  5. Cozzolino, F.; Iacobucci, I.; Monaco, V.; Angrisano, T.; Monti, M. Lysines Acetylome and Methylome Profiling of H3 and H4 Histones in Trichostatin A—Treated Stem Cells. Int. J. Mol. Sci. 2021, 22, 2063. [Google Scholar] [CrossRef]
  6. Ding, P.; Ma, Z.; Liu, D.; Pan, M.; Li, H.; Feng, Y.; Zhang, Y.; Shao, C.; Jiang, M.; Lu, D.; et al. Lysine Acetylation/Deacetylation Modification of Immune-Related Molecules in Cancer Immunotherapy. Front. Immunol. 2022, 13, 865975. [Google Scholar] [CrossRef] [PubMed]
  7. Akhter, Z.; Bi, Z.; Ali, K.; Sun, C.; Fiaz, S.; Haider, F.U.; Bai, J. In Response to Abiotic Stress, DNA Methylation Confers EpiGenetic Changes in Plants. Plants 2021, 10, 1096. [Google Scholar] [CrossRef]
  8. Sharma, M.; Sidhu, A.K.; Samota, M.K.; Gupta, M.; Koli, P.; Choudhary, M. Post-Translational Modifications in Histones and Their Role in Abiotic Stress Tolerance in Plants. Proteomes 2023, 11, 38. [Google Scholar] [CrossRef] [PubMed]
  9. Gallinari, P.; Di Marco, S.; Jones, P.; Pallaoro, M.; Steinkühler, C. HDACs, Histone Deacetylation and Gene Transcription: From Molecular Biology to Cancer Therapeutics. Cell Res. 2007, 17, 195–211. [Google Scholar] [CrossRef] [PubMed]
  10. Liu, R.; Zhang, L.; Zhang, K. Histone Modification in Psoriasis: Molecular Mechanisms and Potential Therapeutic Targets. Exp. Dermatol. 2024, 33, e15151. [Google Scholar] [CrossRef]
  11. Milazzo, G.; Mercatelli, D.; Di Muzio, G.; Triboli, L.; De Rosa, P.; Perini, G.; Giorgi, F.M. Histone Deacetylases (HDACs): Evolution, Specificity, Role in Transcriptional Complexes, and Pharmacological Actionability. Genes 2020, 11, 556. [Google Scholar] [CrossRef] [PubMed]
  12. Pandey, R.; Müller, A.; Napoli, C.A.; Selinger, D.A.; Pikaard, C.S.; Richards, E.J.; Bender, J.; Mount, D.W.; Jorgensen, R.A. Analysis of Histone Acetyltransferase and Histone Deacetylase Families of Arabidopsis Thaliana Suggests Functional Diversification of Chromatin Modification among Multicellular Eukaryotes. Nucleic Acids Res. 2002, 30, 5036–5055. [Google Scholar] [CrossRef] [PubMed]
  13. Zhao, L.; Lu, J.; Zhang, J.; Wu, P.Y.; Yang, S.; Wu, K. Identification and Characterization of Histone Deacetylases in Tomato (Solanum lycopersicum). Front. Plant Sci. 2015, 5, 119871. [Google Scholar] [CrossRef] [PubMed]
  14. Liu, X.; Yang, S.; Yu, C.W.; Chen, C.Y.; Wu, K. Histone Acetylation and Plant Development. Enzymes 2016, 40, 173–199. [Google Scholar] [CrossRef] [PubMed]
  15. Cai, Y.; Xu, M.; Liu, J.; Zeng, H.; Song, J.; Sun, B.; Chen, S.; Deng, Q.; Lei, J.; Cao, B.; et al. Genome-Wide Analysis of Histone Acetyltransferase and Histone Deacetylase Families and Their Expression in Fruit Development and Ripening Stage of Pepper (Capsicum annuum). Front. Plant Sci. 2022, 13, 971230. [Google Scholar] [CrossRef]
  16. Osipowski, P.; Pawełkowicz, M.; Wojcieszek, M.; Skarzyńska, A.; Przybecki, Z.; Pląder, W. A High-Quality Cucumber Genome Assembly Enhances Computational Comparative Genomics. Mol. Genet. Genom. 2020, 295, 177–193. [Google Scholar] [CrossRef]
  17. Camacho, C.; Coulouris, G.; Avagyan, V.; Ma, N.; Papadopoulos, J.; Bealer, K.; Madden, T.L. BLAST+: Architecture and Applications. BMC Bioinform. 2009, 10, 421. [Google Scholar] [CrossRef]
  18. Li, Q.; Li, H.; Huang, W.; Xu, Y.; Zhou, Q.; Wang, S.; Ruan, J.; Huang, S.; Zhang, Z. A Chromosome-Scale Genome Assembly of Cucumber (Cucumis sativus L.). Gigascience 2019, 8, giz072. [Google Scholar] [CrossRef] [PubMed]
  19. Yang, L.; Koo, D.H.; Li, Y.; Zhang, X.; Luan, F.; Havey, M.J.; Jiang, J.; Weng, Y. Chromosome Rearrangements during Domestication of Cucumber as Revealed by High-Density Genetic Mapping and Draft Genome Assembly. Plant J. 2012, 71, 895–906. [Google Scholar] [CrossRef] [PubMed]
  20. Aiese Cigliano, R.; Sanseverino, W.; Cremona, G.; Ercolano, M.R.; Conicella, C.; Consiglio, F.M. Genome-Wide Analysis of Histone Modifiers in Tomato: Gaining an Insight into Their Developmental Roles. BMC Genom. 2013, 14, 57. [Google Scholar] [CrossRef] [PubMed]
  21. Zdobnov, E.M.; Apweiler, R. InterProScan—An Integration Platform for the Signature-Recognition Methods in InterPro. Bioinformatics 2001, 17, 847–848. [Google Scholar] [CrossRef] [PubMed]
  22. Larkin, M.A.; Blackshields, G.; Brown, N.P.; Chenna, R.; Mcgettigan, P.A.; McWilliam, H.; Valentin, F.; Wallace, I.M.; Wilm, A.; Lopez, R.; et al. Clustal W and Clustal X Version 2.0. Bioinformatics 2007, 23, 2947–2948. [Google Scholar] [CrossRef]
  23. Trifinopoulos, J.; Nguyen, L.T.; von Haeseler, A.; Minh, B.Q. W-IQ-TREE: A Fast Online Phylogenetic Tool for Maximum Likelihood Analysis. Nucleic Acids Res. 2016, 44, W232–W235. [Google Scholar] [CrossRef]
  24. Letunic, I.; Bork, P. Interactive Tree of Life (ITOL) v6: Recent Updates to the Phylogenetic Tree Display and Annotation Tool. Nucleic Acids Res. 2024, 52, W78–W82. [Google Scholar] [CrossRef]
  25. Bailey, T.L.; Johnson, J.; Grant, C.E.; Noble, W.S. The MEME Suite. Nucleic Acids Res. 2015, 43, W39–W49. [Google Scholar] [CrossRef] [PubMed]
  26. Rice, P.; Longden, L.; Bleasby, A. EMBOSS: The European Molecular Biology Open Software Suite. Trends Genet. 2000, 16, 276–277. [Google Scholar] [CrossRef] [PubMed]
  27. Horton, P.; Park, K.J.; Obayashi, T.; Fujita, N.; Harada, H.; Adams-Collier, C.J.; Nakai, K. WoLF PSORT: Protein Localization Predictor. Nucleic Acids Res. 2007, 35, W585. [Google Scholar] [CrossRef] [PubMed]
  28. Gu, Z.; Gu, L.; Eils, R.; Schlesner, M.; Brors, B. Circlize Implements and Enhances Circular Visualization in R. Bioinformatics 2014, 30, 2811–2812. [Google Scholar] [CrossRef] [PubMed]
  29. Turek, S.; Pląder, W.; Hoshi, Y.; Skarzyńska, A.; Pawełkowicz, M. Insight into the Organization of the B10v3 Cucumber Genome by Integration of Biological and Bioinformatic Data. Int. J. Mol. Sci. 2023, 24, 4011. [Google Scholar] [CrossRef]
  30. Chen, C.; Wu, Y.; Li, J.; Wang, X.; Zeng, Z.; Xu, J.; Liu, Y.; Feng, J.; Chen, H.; He, Y.; et al. TBtools-II: A “One for All, All for One” Bioinformatics Platform for Biological Big-Data Mining. Mol. Plant 2023, 16, 1733–1742. [Google Scholar] [CrossRef]
  31. Patro, R.; Duggal, G.; Love, M.I.; Irizarry, R.A.; Kingsford, C. Salmon: Fast and Bias-Aware Quantification of Transcript Expression Using Dual-Phase Inference. Nat. Methods 2017, 14, 417. [Google Scholar] [CrossRef] [PubMed]
  32. Ritchie, M.E.; Phipson, B.; Wu, D.; Hu, Y.; Law, C.W.; Shi, W.; Smyth, G.K. Limma Powers Differential Expression Analyses for RNA-Sequencing and Microarray Studies. Nucleic Acids Res. 2015, 43, e47. [Google Scholar] [CrossRef] [PubMed]
  33. Warnes, G.R.; Bolker, B.; Bonebakker, L.; Gentleman, R.; Huber, W.; Liaw, A.; Lumley, T.; Maechler, M.; Magnusson, A.; Moeller, S.; et al. Various R Programming Tools for Plotting Data. CRAN Contrib. Packages. 2022. Available online: https://cran.r-project.org/web/packages/gplots/index.html (accessed on 12 December 2024).
  34. Lescot, M.; Déhais, P.; Thijs, G.; Marchal, K.; Moreau, Y.; Van De Peer, Y.; Rouzé, P.; Rombauts, S. PlantCARE, a Database of Plant Cis-Acting Regulatory Elements and a Portal to Tools for in Silico Analysis of Promoter Sequences. Nucleic Acids Res. 2002, 30, 325–327. [Google Scholar] [CrossRef]
  35. Zhao, T.; Zhan, Z.; Jiang, D. Histone Modifications and Their Regulatory Roles in Plant Development and Environmental Memory. J. Genet. Genom. 2019, 46, 467–476. [Google Scholar] [CrossRef] [PubMed]
  36. Sena, S.; Prakash, A.; Van Staden, J.; Kumar, V. Epigenetic Control of Plant Regeneration: Unraveling the Role of Histone Methylation. Curr. Plant Biol. 2024, 40, 100408. [Google Scholar] [CrossRef]
  37. Brukhin, V.; Albertini, E. Epigenetic Modifications in Plant Development and Reproduction. Epigenomes 2021, 5, 25. [Google Scholar] [CrossRef]
  38. Li, H.; Liu, H.; Pei, X.; Chen, H.; Li, X.; Wang, J.; Wang, C. Comparative Genome-Wide Analysis and Expression Profiling of Histone Acetyltransferases and Histone Deacetylases Involved in the Response to Drought in Wheat. J. Plant Growth Regul. 2022, 41, 1065–1078. [Google Scholar] [CrossRef]
  39. Aquea, F.; Timmermann, T.; Arce-Johnson, P. Analysis of Histone Acetyltransferase and Deacetylase Families of Vitis Vinifera. Plant Physiol. Biochem. 2010, 48, 194–199. [Google Scholar] [CrossRef] [PubMed]
  40. Bai, S.L.; Peng, Y.B.; Cui, J.X.; Gu, H.T.; Xu, L.Y.; Li, Y.Q.; Xu, Z.H.; Bai, S.N. Developmental Analyses Reveal Early Arrests of the Spore-Bearing Parts of Reproductive Organs in Unisexual Flowers of Cucumber (Cucumis sativus L.). Planta 2004, 220, 230–240. [Google Scholar] [CrossRef]
  41. Fu, W.; Wu, K.; Duan, J. Sequence and Expression Analysis of Histone Deacetylases in Rice. Biochem. Biophys. Res. Commun. 2007, 356, 843–850. [Google Scholar] [CrossRef]
  42. Kumar, V.; Thakur, J.K.; Prasad, M. Histone Acetylation Dynamics Regulating Plant Development and Stress Responses. Cell Mol. Life Sci. 2021, 78, 4467. [Google Scholar] [CrossRef]
  43. Luo, M.; Tai, R.; Yu, C.W.; Yang, S.; Chen, C.Y.; Lin, W.D.; Schmidt, W.; Wu, K. Regulation of Flowering Time by the Histone Deacetylase HDA5 in Arabidopsis. Plant J. 2015, 82, 925–936. [Google Scholar] [CrossRef] [PubMed]
  44. Yu, C.W.; Liu, X.; Luo, M.; Chen, C.; Lin, X.; Tian, G.; Lu, Q.; Cui, Y.; Wu, K. HISTONE DEACETYLASE6 Interacts with FLOWERING LOCUS D and Regulates Flowering in Arabidopsis. Plant Physiol. 2011, 156, 173–184. [Google Scholar] [CrossRef] [PubMed]
  45. Han, Y.C.; Kuang, J.F.; Chen, J.Y.; Liu, X.C.; Xiao, Y.Y.; Fu, C.C.; Wang, J.N.; Wu, K.Q.; Lu, W.J. Banana Transcription Factor MaERF11 Recruits Histone Deacetylase MaHDA1 and Represses the Expression of MaACO1 and Expansins during Fruit Ripening. Plant Physiol. 2016, 171, 1070–1084. [Google Scholar] [CrossRef] [PubMed]
  46. Fu, C.C.; Han, Y.C.; Guo, Y.F.; Kuang, J.F.; Chen, J.Y.; Shan, W.; Lu, W.J. Differential Expression of Histone Deacetylases during Banana Ripening and Identification of MaHDA6 in Regulating Ripening-Associated Genes. Postharvest Biol. Technol. 2018, 141, 24–32. [Google Scholar] [CrossRef]
  47. Guo, J.E.; Hu, Z.; Yu, X.; Li, A.; Li, F.; Wang, Y.; Tian, S.; Chen, G. A Histone Deacetylase Gene, SlHDA3, Acts as a Negative Regulator of Fruit Ripening and Carotenoid Accumulation. Plant Cell Rep. 2018, 37, 125–135. [Google Scholar] [CrossRef]
  48. Guo, J.E.; Hu, Z.; Li, F.; Zhang, L.; Yu, X.; Tang, B.; Chen, G. Silencing of Histone Deacetylase SlHDT3 Delays Fruit Ripening and Suppresses Carotenoid Accumulation in Tomato. Plant Sci. 2017, 265, 29–38. [Google Scholar] [CrossRef] [PubMed]
  49. Fu, C.; Chen, H.; Gao, H.; Han, Y. Histone Deacetylase CpHDA3 Is Functionally Associated with CpERF9 in Suppression of CpPME1/2 and CpPG5 Genes during Papaya Fruit Ripening. J. Agric. Food Chem. 2019, 67, 8919–8925. [Google Scholar] [CrossRef] [PubMed]
  50. Pawełkowicz, M.E.; Skarzyńska, A.; Pląder, W.; Przybecki, Z. Genetic and Molecular Bases of Cucumber (Cucumis sativus L.) Sex Determination. Mol. Breed. 2019, 39, 50. [Google Scholar] [CrossRef]
  51. Su, Z.; Ma, X.; Guo, H.; Sukiran, N.L.; Guo, B.; Assmann, S.M.; Ma, H. Flower Development under Drought Stress: Morphological and Transcriptomic Analyses Reveal Acute Responses and Long-Term Acclimation in Arabidopsis. Plant Cell 2013, 25, 3785–3807. [Google Scholar] [CrossRef]
  52. Li, W.F.; Mao, J.; Su, J.; Li, X.W.; Dawuda, M.M.; Ma, Z.H.; Zuo, C.W.; An, Z.S.; Chen, B.H. Exogenous ABA and Its Inhibitor Regulate Flower Bud Induction of Apple Cv. ‘Nagafu No. 2′ Grafted on Different Rootstocks. Trees—Struct. Funct. 2021, 35, 609–620. [Google Scholar] [CrossRef]
Genes 16 00127 g001
Figure 1. Phylogenetic analysis of HAT proteins from Cucumis sativus (36 proteins, indicated by green circles), Arabidopsis thaliana (12 proteins, indicated by blue circles), and Solanum lycopersicum (29 proteins, indicated by red circles). The phylogenetic tree was built using the neighbor-joining method. The HAG, HAM, HAF, and HAC families have been marked with different colors.
Figure 1. Phylogenetic analysis of HAT proteins from Cucumis sativus (36 proteins, indicated by green circles), Arabidopsis thaliana (12 proteins, indicated by blue circles), and Solanum lycopersicum (29 proteins, indicated by red circles). The phylogenetic tree was built using the neighbor-joining method. The HAG, HAM, HAF, and HAC families have been marked with different colors.
Genes 16 00127 g001
Genes 16 00127 g002
Figure 2. Phylogenetic analysis of HDAC proteins from Cucumis sativus (12 proteins, indicated by green circles), Arabidopsis thaliana (15 proteins, marked with blue circles), and Solanum lycopersicum (12 proteins, marked with red circles). The phylogenetic tree was built using the neighbor-joining method. The HDA, HDT, and SRT families were indicated by different colors.
Figure 2. Phylogenetic analysis of HDAC proteins from Cucumis sativus (12 proteins, indicated by green circles), Arabidopsis thaliana (15 proteins, marked with blue circles), and Solanum lycopersicum (12 proteins, marked with red circles). The phylogenetic tree was built using the neighbor-joining method. The HDA, HDT, and SRT families were indicated by different colors.
Genes 16 00127 g002
Genes 16 00127 g003
Figure 3. Phylogenetic relationship, conserved motifs, protein domain distribution, and gene structure of identified CsHAT genes. (A) The phylogenetic tree was performed with the N-J method. (B) Motif composition predicted using MEME. The 30 identified motifs are indicated by boxes. (C) Protein domain distributions were constructed based on the pfam database. The 30 domains are indicated by colored boxes. (D) CsHAT gene structures were constructed based on the B10v3 genome annotation. Yellow boxes indicate exons, black lines indicate introns, and green boxes indicate UTRs.
Figure 3. Phylogenetic relationship, conserved motifs, protein domain distribution, and gene structure of identified CsHAT genes. (A) The phylogenetic tree was performed with the N-J method. (B) Motif composition predicted using MEME. The 30 identified motifs are indicated by boxes. (C) Protein domain distributions were constructed based on the pfam database. The 30 domains are indicated by colored boxes. (D) CsHAT gene structures were constructed based on the B10v3 genome annotation. Yellow boxes indicate exons, black lines indicate introns, and green boxes indicate UTRs.
Genes 16 00127 g003
Genes 16 00127 g004
Figure 4. Phylogenetic relationship, conserved motifs, protein domain distribution, and gene structure of identified CsHDAC genes. (A) The phylogenetic tree was made with the N-J method. (B) Motif composition predicted using MEME. The 21 identified motifs are indicated by boxes. (C) Protein domain distribution was created using the pfam database. Eight domains are marked with colored boxes. (D) CsHDAC gene structures were constructed based on the B10v3 genome annotation. Yellow boxes indicate exons, black lines indicate introns, and green boxes indicate UTRs.
Figure 4. Phylogenetic relationship, conserved motifs, protein domain distribution, and gene structure of identified CsHDAC genes. (A) The phylogenetic tree was made with the N-J method. (B) Motif composition predicted using MEME. The 21 identified motifs are indicated by boxes. (C) Protein domain distribution was created using the pfam database. Eight domains are marked with colored boxes. (D) CsHDAC gene structures were constructed based on the B10v3 genome annotation. Yellow boxes indicate exons, black lines indicate introns, and green boxes indicate UTRs.
Genes 16 00127 g004
Genes 16 00127 g005
Figure 5. Chromosomal location of CsHAT and CsHDAC genes in cucumber B10v3. The position of the genes on each chromosome is indicated by bars—CsHATs in purple and CsHDAC in orange.
Figure 5. Chromosomal location of CsHAT and CsHDAC genes in cucumber B10v3. The position of the genes on each chromosome is indicated by bars—CsHATs in purple and CsHDAC in orange.
Genes 16 00127 g005
Genes 16 00127 g006
Figure 6. Heatmap illustrating CsHAT expression profiles across different samples: leaves (LV), shoot apexes (SA), and flower buds at different developmental stages—1–2 mm (FB12), 3–5 mm (FB35), 6–8 mm (FB68), and 9 mm (FB9). The color gradient ranges from red, indicating higher expression levels, to green, representing lower expression levels, with values in TPM.
Figure 6. Heatmap illustrating CsHAT expression profiles across different samples: leaves (LV), shoot apexes (SA), and flower buds at different developmental stages—1–2 mm (FB12), 3–5 mm (FB35), 6–8 mm (FB68), and 9 mm (FB9). The color gradient ranges from red, indicating higher expression levels, to green, representing lower expression levels, with values in TPM.
Genes 16 00127 g006
Genes 16 00127 g007
Figure 7. Heatmap illustrating CsHDAC expression profiles across different samples: leaves (LV), shoot apexes (SA), and flower buds at various developmental stages—1–2 mm (FB12), 3–5 mm (FB35), 6–8 mm (FB68), and 9 mm (FB9). The color gradient ranges from red, indicating higher expression levels, to green, representing lower expression levels, with values in TPM.
Figure 7. Heatmap illustrating CsHDAC expression profiles across different samples: leaves (LV), shoot apexes (SA), and flower buds at various developmental stages—1–2 mm (FB12), 3–5 mm (FB35), 6–8 mm (FB68), and 9 mm (FB9). The color gradient ranges from red, indicating higher expression levels, to green, representing lower expression levels, with values in TPM.
Genes 16 00127 g007
Genes 16 00127 g008
Figure 8. Venn diagram showing distribution of CsHATs and CsHDACs with differential expression exhibiting significant changes in comparisons between: (A) different developmental stages of flower buds: shoot apexes (SA) and flower buds of following lengths: 1–2 mm (FB12), 3–5 mm (FB35), 6–8 mm (FB68), and 9 mm (FB9) (B) developmental stages of flower buds (shoot apexes (SA) and flower buds of following lengths: 1–2 mm (FB12), 3–5 mm (FB35), 6–8 mm (FB68), and 9 mm(FB9)), and leaves (LV).
Figure 8. Venn diagram showing distribution of CsHATs and CsHDACs with differential expression exhibiting significant changes in comparisons between: (A) different developmental stages of flower buds: shoot apexes (SA) and flower buds of following lengths: 1–2 mm (FB12), 3–5 mm (FB35), 6–8 mm (FB68), and 9 mm (FB9) (B) developmental stages of flower buds (shoot apexes (SA) and flower buds of following lengths: 1–2 mm (FB12), 3–5 mm (FB35), 6–8 mm (FB68), and 9 mm(FB9)), and leaves (LV).
Genes 16 00127 g008
Genes 16 00127 g009
Figure 9. Cis-acting elements in the promoters of CsHAT genes. Distribution of cis-acting elements divided into major classes according to their function, in regions of 1000 bp upstream of the TSS. To improve clarity, the core elements have been omitted from the visualization.
Figure 9. Cis-acting elements in the promoters of CsHAT genes. Distribution of cis-acting elements divided into major classes according to their function, in regions of 1000 bp upstream of the TSS. To improve clarity, the core elements have been omitted from the visualization.
Genes 16 00127 g009
Genes 16 00127 g010
Figure 10. Number of cis-active elements in the promoters of CsHAT genes. The colors indicate the more abundant cis-active elements, green color indicate cis-elements, red color indicates core elements.
Figure 10. Number of cis-active elements in the promoters of CsHAT genes. The colors indicate the more abundant cis-active elements, green color indicate cis-elements, red color indicates core elements.
Genes 16 00127 g010
Genes 16 00127 g011
Figure 11. Cis-acting element analysis in the promoters of CsHDAC genes. Distribution of cis-acting elements divided into major classes according to their function, in regions of 1000 bp upstream of the TSS. To improve clarity, the core elements have been omitted from the visualization.
Figure 11. Cis-acting element analysis in the promoters of CsHDAC genes. Distribution of cis-acting elements divided into major classes according to their function, in regions of 1000 bp upstream of the TSS. To improve clarity, the core elements have been omitted from the visualization.
Genes 16 00127 g011
Genes 16 00127 g012
Figure 12. Number of cis-active elements in the promoters of CsHDAC genes. The colors indicate the more abundant cis-active elements, green color indicate cis-elements, red color indicates core elements.
Figure 12. Number of cis-active elements in the promoters of CsHDAC genes. The colors indicate the more abundant cis-active elements, green color indicate cis-elements, red color indicates core elements.
Genes 16 00127 g012
Table 1. Summary of CsHAT and CsHDAC proteins identified in B10v3 cucumber genome, compared to available data for Arabidopsis thaliana and Solanum lycopersicum.
Table 1. Summary of CsHAT and CsHDAC proteins identified in B10v3 cucumber genome, compared to available data for Arabidopsis thaliana and Solanum lycopersicum.
TypeFamilySpecies
Cucumis sativus
B10v3		Arabidopsis thalianaSolanum lycopersicum
HATHAG28323
HAM121
HAC754
HAF021
Total361229
HDACHDA997
HDT343
SRT022
Total121512
Table 2. List of CsHATs and CsHDACs genes identified in B10v3 genome and the physicochemical properties of the encoded proteins.
Table 2. List of CsHATs and CsHDACs genes identified in B10v3 genome and the physicochemical properties of the encoded proteins.
FamilyGene IDChr. NoChr Location Start/StopStringLength GenomicLength PeptideMolecular Weight
(kD)		Isoelectric Point (Mw)Localization
Histone acetyltransferases (HATs)
HAG family
HAGCucsat.G7439Chr1ctg1528:2195363-2199742+438037443.37.5nuclear
HAGCucsat.G7491Chr1ctg1528:3357562-3360155+259429934.69.3cytoplasmic
HAGCucsat.G7550Chr1ctg1528:4490591-4496659+606951658.55.9nuclear
HAGCucsat.G8133Chr1ctg1557:892275-894304+203039745.38.3cytoplasmic
HAGCucsat.G11039Chr1ctg1740:323653-327391+373956463.68.5cytoplasmic
HAGCucsat.G14614Chr1ctg1869:2371776-2380341+856626930.39.4nuclear
HAGCucsat.G4728Chr2ctg1227:2564029-2569251+522316418.69.0cytoplasmic
HAGCucsat.G19032Chr2ctg35:283689-291641−795325529.18.7cytoplasmic
HAGCucsat.G19739Chr2ctg4:329471-333655+418546452.14.8cytoplasmic
HAGCucsat.G1301Chr3ctg1:11167598-11170238+264141547.38.4cytoskeleton
HAGCucsat.G505Chr3ctg1:9926620-9928112−149316919.36.0cytoplasmic
HAGCucsat.G12163Chr3ctg1837:1420082-1422790−270940946.48.8cytoplasmic
HAGCucsat.G13099Chr3ctg1838:168257-179247+10,99116318.55.7nuclear
HAGCucsat.G8005Chr4ctg1556:2247813-2251525−371311312.59.0nuclear
HAGCucsat.G8051Chr4ctg1556:3460680-3465076−439724827.86.1chloroplast
HAGCucsat.G8928Chr4ctg1635:1972081-1987877−15,7971403155.36.1nuclear
HAGCucsat.G9940Chr5ctg1673:1987918-1994420+650329934.28.3chloroplast
HAGCucsat.G16316Chr5ctg2246:313272-316828−355719622.46.2cytoplasmic
HAGCucsat.G1790Chr6ctg1002:2300165-2307083−691963470.18.3cytoplasmic
HAGCucsat.G2199Chr6ctg1002:985444-1000064+14,6211030116.18.0cytoplasmic
HAGCucsat.G2255Chr6ctg1002:2192294-2194872+257941046.28.9cytoplasmic
HAGCucsat.G5632Chr6ctg1299:318309-325444+71361432161.58.1nuclear
HAGCucsat.G5634Chr6ctg1299:396719-398897+217941547.88.7nuclear,
cytoplasmic
HAGCucsat.G17933Chr6ctg3345:1507356-1519041+11,686972107.26.2nuclear
HAGCucsat.G2757Chr7ctg1041:822751-825505−275527130.78.1chloroplast
HAGCucsat.G3385Chr7ctg1041:6256432-6258342+191122024.87.9extracellular
HAGCucsat.G10849Chr7ctg1681:902951-910263−731322525.95.7cytoplasmic
HAGCucsat.G10916Chr7ctg1681:2018378-2026590−821329733.18.8chloroplast
HAC family
HACCucsat.G5059Chr2ctg1227:2823211-2832569−935939544.29.5nuclear
HACCucsat.G19498Chr2ctg4:1282651-1287430−478042347.87.0nuclear
HACCucsat.G716Chr3ctg1:42742-56672+13,9311707192.68.1nuclear
HACCucsat.G13206Chr3ctg1838:2203445-2207740+429672180.59.5nuclear
HACCucsat.G13466Chr3ctg1838:7194802-7198169+336835540.48.6cytoplasmic
HACCucsat.G9211Chr7ctg1658:1590288-1598827+854012714.44.8nuclear
HACCucsat.G9212Chr7ctg1658:1599861-1614805+149451365149.89.9nuclear
HAM family
HAMCucsat.G7714Chr5ctg1546:298050-304738+668949557.38.6chloroplast
Histone deacetylases (HDACs)
HDA family
HDACucsat.G11084Chr1ctg1740:159401-162484−308446552.35.8cytoplasmic
HDACucsat.G11217Chr1ctg1780:533431-536277+284737541.05.8cytoplasmic, cytoskeleton
HDACucsat.G7096Chr1ctg1528:914219-935116−20,89860566.46.8nuclear
HDACucsat.G14544Chr1ctg1869:945967-954231+826565973.25.3chloroplast
HDACucsat.G1690Chr4ctg1002:373456-379733−627844248.26.6chloroplast
HDACucsat.G11372Chr4ctg1798:64712-72066−735535238.97.5cytoskeleton
HDACucsat.G18033Chr6ctg3345:3960732-3965777+504649255.65.0nuclear
HDACucsat.G18140Chr6ctg3345:1224920-1231792−687344050.45.1mitochondrial
HDACucsat.G10868Chr7ctg1681:1172904-1178169−526647153.35.7cytoplasmic
HDT family
HDTCucsat.G12291Chr3ctg1837:3557001-3565310−831031834.64.4nuclear
HDTCucsat.G17927Chr6ctg3345:1319359-1323178+382029631.74.4nuclear
HDTCucsat.G17928Chr6ctg3345:1323822-1327471+365057161.64.8nuclear
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Skarzyńska-Łyżwa, A.; Turek, S.; Pisz, M.; Aparna; Pląder, W.; Pawełkowicz, M. Genome-Wide Identification and Characterization of Histone Acetyltransferases and Deacetylases in Cucumber, and Their Implication in Developmental Processes. Genes 2025, 16, 127. https://doi.org/10.3390/genes16020127

AMA Style

Skarzyńska-Łyżwa A, Turek S, Pisz M, Aparna, Pląder W, Pawełkowicz M. Genome-Wide Identification and Characterization of Histone Acetyltransferases and Deacetylases in Cucumber, and Their Implication in Developmental Processes. Genes. 2025; 16(2):127. https://doi.org/10.3390/genes16020127

Chicago/Turabian Style

Skarzyńska-Łyżwa, Agnieszka, Szymon Turek, Maksymilian Pisz, Aparna, Wojciech Pląder, and Magdalena Pawełkowicz. 2025. "Genome-Wide Identification and Characterization of Histone Acetyltransferases and Deacetylases in Cucumber, and Their Implication in Developmental Processes" Genes 16, no. 2: 127. https://doi.org/10.3390/genes16020127

APA Style

Skarzyńska-Łyżwa, A., Turek, S., Pisz, M., Aparna, Pląder, W., & Pawełkowicz, M. (2025). Genome-Wide Identification and Characterization of Histone Acetyltransferases and Deacetylases in Cucumber, and Their Implication in Developmental Processes. Genes, 16(2), 127. https://doi.org/10.3390/genes16020127

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop