Next Article in Journal
IPM Adoption in Common Beans in Brazil
Previous Article in Journal
From Sun to Snack: Different Drying Methods and Their Impact on Crispiness and Consumer Acceptance of Royal Gala Apple Snacks
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Horticulturae
Volume 11
Issue 6
10.3390/horticulturae11060609
horticulturae-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

A Genome-Wide Characterization of the Xyloglucan Endotransglucosylase/Hydrolase Family Genes and Their Functions in the Shell Formation of Pecan

by
Mengyun Wen
1,†,
Zekun Zhou
1,†,
Jing Sun
1,
Fanqing Meng
1,
Xueliang Xi
2,
Aizhong Liu
1,* and
Anmin Yu
1,3,4,*
1
Key Laboratory for Forest Resources Conservation and Utilization in the Southwest Mountains of China, Ministry of Education, Southwest Forestry University, Kunming 650224, China
2
Yunnan Academy of Forestry and Grassland, Kunming 650201, China
3
Yunnan Provincial Key Laboratory for Conservation and Utilization of In-Forest Resource, Southwest Forestry University, Kunming 650224, China
4
Yunnan Key Laboratory of Crop Wild Relatives Omics, Kunming Institute of Botany, Chinese Academy of Sciences, Kunming 650201, China
*
Authors to whom correspondence should be addressed.
†
These authors contributed equally to this work.
Horticulturae 2025, 11(6), 609; https://doi.org/10.3390/horticulturae11060609
Submission received: 9 April 2025 / Revised: 21 May 2025 / Accepted: 28 May 2025 / Published: 29 May 2025
(This article belongs to the Section Genetics, Genomics, Breeding, and Biotechnology (G2B2))
Browse Figures
Review Reports Versions Notes

Abstract

Xyloglucan endotransglucosylases/hydrolases (XTHs) are key enzymes involved in cell wall remodeling by modifying xyloglucan–cellulose networks, thereby influencing plant growth, development, and secondary cell wall formation. While the roles of XTHs have been extensively studied in primary and secondary growth, their functions in the formation and thickening of lignified nut shells remain largely unknown. Pecan (Carya illinoinensis), an economically important nut crop, develops a hard, lignified shell that protects the seed during fruit maturation. In this study, we performed a comprehensive genome-wide characterization of the XTH gene family in pecan and identified 38 XTH genes, which were categorized into four distinct phylogenetic groups. Structural analyses of the deduced proteins revealed conserved catalytic residues alongside divergent loop regions, suggesting functional diversification. Expression profiling across various tissues and among pecan cultivars with contrasting shell phenotypes indicated that specific XTH genes may play critical roles in shell structure formation. Moreover, gene regulatory networks in thin- and thick-shelled pecans provided new insights into the molecular mechanisms underlying shell development and thickness regulation. These findings lay a foundation for future genetic improvement strategies targeting nut shell traits in woody perennials.

1. Introduction

Plant cell walls, including the primary and secondary cell walls (SCWs), are highly dynamic structures. The formation of the primary cell wall occurs after cell division with cell growth and expansion, and it is composed of cellulose, hemicellulose, and pectins [1,2]. In contrast, the deposition of the SCW starts after the cessation of cell expansion to provide mechanical strength and protection, and it is composed of cellulose, hemicellulose, and lignin [3]. Among hemicelluloses, xyloglucan (XyG) is one of the most abundant components in dicotyledonous plants, accounting for approximately 20–25% of the primary cell wall [4]. Structurally, XyG consists of a β-1,4-linked D-glucose backbone, substituted with D-xylose residues, which may be further decorated with galactose and fucose, resulting in structural diversity among plant species [5]. In the plant cell wall, XyG interacts closely with cellulose microfibrils through hydrogen bonding, forming a key structural network that regulates wall extensibility, mechanical strength, and integrity, thereby linking cell wall architecture with developmental and environmental signaling pathways [6].
Xyloglucan endotransglucosylase/hydrolase (XTH) is a multifunctional enzyme family crucial for cell wall remodeling through dynamic XyG modification [7]. These enzymes exhibit two distinct activities: xyloglucan endotransglucosylase (XET) activity, which catalyzes the cleavage and subsequent transglycosylation of XyG chains, thereby modulating cell wall extensibility, and xyloglucan hydrolase (XEH) activity, which hydrolyzes XyG molecules, contributing to cell wall loosening and degradation in processes such as fruit ripening and seed germination [8,9]. In addition, XTH also participates in reinforcing the wall matrix during SCW biosynthesis, linking cell expansion to subsequent strengthening [4]. XTHs play a central role in balancing wall extensibility and reinforcement by coordinating with cellulose synthases, expansins, and peroxidases during SCW biosynthesis [10].
XTHs belong to the glycoside hydrolase family 16 (GH16), a diverse group that also includes microbial endoglucanases and other plant-specific cell wall-modifying enzymes [11]. The genomes of land plants typically encode around 30 XTH genes, with 33 members in Arabidopsis thaliana [12], 29 in Oryza sativa [13], and 43 in Populus trichocarpa [14]. The XTH family genes are classified into three (I/II, IIIA, and IIIB) or four major groups (I, II, IIIA, and IIIB). Group IIIA members exhibit xyloglucan hydrolase (XEH) activity, hydrolyzing XyG polymers [8], whereas Groups I/II and IIIB function as xyloglucan endotransglucosylases (XETs).
XTHs dynamically remodel XyG–cellulose networks, influencing cell wall elasticity and extensibility [10]. Their activity is closely linked to cell expansion, as they mediate the targeted hydrolysis and transglycosylation of XyG, disrupting hydrogen bonds with cellulose microfibrils to promote cell wall loosening. This function is particularly evident in rapidly growing tissues such as hypocotyls and roots [15]. For example, the overexpression of AtXTH9 in A. thaliana leads to a 25–30% increase in internodal length, demonstrating the direct impact of XTH enzymatic activity on tissue elongation [16]. Similarly, PcBRU1 in pear (Pyrus communis) regulates plant height and internode elongation by modulating cell wall extensibility and facilitating cell elongation [17]. Beyond their roles in primary cell wall dynamics, XTHs are also integral to SCW formation and lignification [4]. In A. thaliana, AtXTH4 is highly expressed in stems and seedlings, facilitating xylem cell expansion and lignified SCW formation through XyG–cellulose remodeling [18]. Additionally, AtXTH18 and AtXTH19 regulate lignified SCW assembly during vascular differentiation, orchestrating XyG–cellulose interactions and lignin integration [6,19]. AtXTH27 plays a crucial role in tracheary element differentiation, mediating XyG–cellulose network restructuring and lignin deposition during xylem development [20]. Similarly, in P. trichocarpa, PtXTH16 and PtXTH34 are highly expressed during SCW formation in xylem tissues, suggesting their importance in vascular differentiation and woody biomass accumulation [21]. In fruit crops, such as tomato (Solanum lycopersicum), SlXTH4 regulates fruit firmness and ripening by modulating XyG–pectin interactions through its XET/XEH activities, facilitating cell wall loosening and the starch-to-sugar transition during climacteric ripening, thereby directly influencing texture and postharvest quality [22]. Furthermore, in barley (Hordeum valgare L.), the overexpression of HvXTH1 significantly alters cell wall architecture and reduces lignin content [23]. Despite these findings, the precise roles of XTHs in the development of lignified protective tissues remain largely unexplored, particularly regarding their involvement in polysaccharide–lignin crosslinking during SCW biogenesis.
Pecan (Carya illinoinensis), a member of the hickory genus (Carya spp., Juglandaceae), is distributed across tropical and temperate regions, and it is widely cultivated on six continents, although it is native to the southern United States and northern Mexico [24]. Pecan is a major economic tree, and the pecan nut is rich in unsaturated fatty acids, antioxidant polyphenols, and essential vitamins, making it a highly sought-after ingredient consumed worldwide in food, confectionery products, and cooking oil production [25]. Given the increasing consumer demand and expanding pecan industry, breeding efforts have primarily focused on improving yield-related traits. However, the genetic improvement of pecan has been largely constrained by its long growth cycle. The pecan seed is encased by a lignified shell, composed of thick-walled sclerenchyma cells and a suberized periderm, which serves as a crucial protective barrier against mechanical damage, pathogen invasion, and environmental stress during dormancy and dispersal [26]. The thickness of the shell is a key trait in pecan breeding, influencing not only nut processing and consumer preference but also seed protection and dispersal efficiency. While thin-shelled varieties are often favored for easier cracking and commercial use, thick-shelled varieties may confer advantages in terms of pest resistance and environmental adaptability. Despite the agronomic significance of shell thickness, the genetic and molecular mechanisms governing shell development and thickness variation remain poorly understood.
Pecan shell formation is a highly coordinated process regulated by genetic, environmental, and hormonal factors. However, the underlying molecular mechanisms, particularly the roles of cell wall-modifying enzymes in shell lignification and thickening, have yet to be fully elucidated. XTH genes play a crucial role in cell wall remodeling, influencing cell expansion, SCW deposition, and lignification in various plant species. In this study, we performed a comprehensive genome-wide characterization of the XTH gene family in pecan, analyzing their phylogenetic relationships, gene structures, chromosomal locations, and expression patterns. Furthermore, we investigated their potential roles in shell formation by examining expression profiles during shell development. Our findings provide novel insights into the molecular mechanisms underlying pecan shell formation and establish a foundation for future functional studies on XTH genes in woody perennials.

2. Materials and Methods

2.1. Plant Materials

Fruits were collected from pecan cultivars “Caddo” and “Shaoxing” trees, growing in LuFeng city, ChuXiong Yi Autonomous Prefecture, Yunnan Province, China. Samples were collected from 18-year-old trees from June to September 2024. There are three endocarp developmental periods, including the rapid expanding stage at 60 days after pollination (DAP, denoted as KD1 and SX1), the hardening stage at 90 DAP (KD2 and SX2), and the mature stage at 120 DAP (KD3 and SX3). The fruits of “Caddo” were slender and had relatively thin shells and a low degree of lignification, whereas “Shaoxing” had short and round fruits, thick shells, and a relatively high degree of lignification, as shown in Figure S1. Fruits of uniform size and maturity were randomly sampled from at least three healthy trees. The green exocarp and seed tissues were carefully removed, leaving only the shell tissues, which were immediately frozen in liquid nitrogen and stored at −80 °C for further analysis.

2.2. Identification and Characterization of XTH Family Genes in Pecan

The reference genome of pecan was obtained from recently published data [27]. The amino acid sequences of all XTHs in Arabidopsis were retrieved from the TAIR database and used as queries to identify putative XTH genes in the pecan genome via a local BLASTP search with an E-value threshold of <1 × 10−10. To confirm candidate genes, two conserved XTH-related domains—Glycosyl hydrolase family 16 (Glyco_hydro_16, PF00722) and xyloglucan endo-transglycosylase C-terminus (XET_C, PF06955)—were retrieved from the Pfam database (http://pfam-legacy.xfam.org/, accessed on 3 January 2025) and employed for genome-wide searches using HMMER (v3.3.2) [28]. The physicochemical properties of the identified CilXTH proteins, including molecular weight (MW) and isoelectric point (pI), were predicted using ExPASy ProtParam (https://web.expasy.org/, accessed on 3 January 2025) [29]. Subcellular localization was predicted with Plant-mPLoc v2.0 (http://www.csbio.sjtu.edu.cn/bioinf/plant-multi/, accessed on 6 January 2025) [30].

2.3. Multiple Sequence Alignment, Conserved Domain, and Gene Structure Analysis

Multiple sequence alignments of all CilXTH proteins were performed using MAFFT and visualized with ESPript 3.0 (https://espript.ibcp.fr/ESPript/cgi-bin/ESPript.cgi, accessed on 6 January 2025). Conserved domains were identified using the InterPro database (https://www.ebi.ac.uk/interpro/, accessed on 9 January 2025). The gene structures of CilXTH genes were analyzed and illustrated using the Gene Structure Display Server (version 2.0) (http://gsds.cbi.pku.edu.cn/, accessed on 9 January 2025). To further explore protein structure, the three-dimensional structures were modeled using AlphaFold3 (https://alphafoldserver.com/, accessed on 16 January 2025). The PDB files generated were visualized and analyzed using PyMOL 3.0 (https://pymol.org/#download, accessed on 16 January 2025) [31,32].

2.4. Evolutionary Relationships of XTH Genes in Pecan

The amino acid sequences of identified CilXTHs from pecan and AtXTHs from A. thaliana were aligned, and a phylogenetic tree was constructed using the Maximum Likelihood (ML) method with 1000 bootstrap replicates. The resulting tree was visualized using iTOL (https://itol.embl.de/, accessed on 20 January 2025) [33].

2.5. Chromosomal Distribution, Collinearity, and Evolutionary Analysis

The chromosomal locations of CilXTH genes were determined based on the pecan genome assembly and visualized using TBtools v2.303 [34]. Gene duplication events were identified using MCScanX [35]. Tandem and segmental duplication events were classified according to collinear gene pair files extracted from the pecan genome. Collinearity relationships among CilXTH genes were further analyzed and visualized using the jcvi package [36]. The synonymous substitution rate (Ks) of collinear gene pairs was calculated using the WGDI package with the core programs -ks and -kp [37].

2.6. RNA Extraction, Library Construction, Expression Profiling, and Gene Co-Expression Analysis

To investigate the expression patterns of CilXTH genes across various pecan tissues, publicly available RNA-seq data were retrieved and analyzed [38]. Tissue-specific expression patterns were assessed using K-means clustering and visualized with the pheatmap package in R [39]. To further explore the potential roles of CilXTH genes in shell thickness determination, total RNA was extracted from developing shell tissues of the thin-shelled cultivar “Caddo” and the thick-shelled cultivar “Shaoxing” at three different developmental stages (60, 90, and 120 DAP) using the RNAprep Pure Plant Kit (Tiangen, Beijing, China), following the manufacturer’s instructions. The quantity and integrity of the RNA were assessed using a NanoDrop 2000 spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA) and an Agilent 2100 Bioanalyzer (Agilent Technologies, Santa Clara, CA, USA), respectively. Only high-quality RNA samples with an RNA integrity number (RIN) ≥7.0 were used for library preparation. RNA-seq libraries were constructed using the NEBNext Ultra II RNA Library Prep Kit for Illumina (San Diego, CA, USA) according to the manufacturer’s protocol. The libraries were then sequenced on an Illumina NovaSeq 6000 platform, generating 150 bp paired-end reads. Raw sequencing data were subjected to quality control using FastQC (v0.11.9) [40] and Trimmomatic (v0.39) [41] to remove low-quality reads and adapter sequences. Clean reads were mapped to the reference genome of pecan using HISAT2 (v2.2.1) [42], and gene-level expression in fragments per kilobase of transcript per million mapped reads (FPKM) was estimated with StringTie (v2.2.1) [43]. Furthermore, expression profiles were compared between the thin-shelled “Caddo” and thick-shelled “Shaoxing” cultivars during shell development through ClusterGVis (https://github.com/junjunlab/ClusterGVis, accessed on 26 January 2025) with K-means methods. Co-expression networks were constructed for genes involved in shell enlargement to maturation stages using the GENIE3 package, based on previously established gene expression profiles [44]. The resulting gene interaction networks were analyzed and visualized using Cytoscape (v3.9.1) [45].

2.7. Cis-Regulatory Elements Analysis of CilXTH Genes

To identify potential cis-regulatory elements, 2000 bp upstream promoter sequences of CilXTH genes were extracted from the pecan genome. The New PLACE database (https://www.dna.affrc.go.jp/PLACE/?action=newplace, accessed on 9 February 2025) was utilized to predict transcription factor binding sites and associated functional motifs. The identified cis-regulatory elements were visualized and analyzed using TBtools v2.303 [32].

3. Results

3.1. The Identification and Structural Features of CiXTH Genes from the Pecan Genome

To comprehensively identify XTH genes in the pecan genome, we employed both BLASTP and HMMER searches. Initially, 38 putative XTH genes were identified through BLAST v2.9.0 analysis (Figure S2). Among them, 33 genes possessed both Glycosyl hydrolase family 16 (Glyco_hydro_16, PF00722) and the xyloglucan endo-transglycosylase C-terminal (XET_C, PF06955) domains, while 4 genes contained only the Glyco_hydro_16 domain, and 1 gene contained only the XET_C domain (Figure S2).
Multiple sequence alignment confirmed that nearly all CilXTH proteins contained the highly conserved ExDxE catalytic motif within the Glyco_hydro_16 domain, along with potential N-linked glycosylation sites (N-X-T/S) (Figure 1). The only exception was Cil_01G_00640V2, which lacked the Glyco_hydro_16 domain but retained the XET_C domain, suggesting it still harbors XET activity (Figure 2). Therefore, we consider all identified CilXTH genes to be potential functional candidates and have sequentially named CilXTH1 to CilXTH38 according to their chromosomal locations (Table 1). The length of CilXTH genes ranged from 297 bp (CilXTH2) to 1779 bp (CilXTH38), and these genes encoded proteins that ranged from 98 to 592 amino acids (aa), with molecular weights between 11.51 and 66.15 kDa. The isoelectric points (pI) of 20 CilXTH proteins were greater than 7.0, indicating that these proteins are predominantly basic in nature. Proteins with high pI values typically contain higher proportions of basic amino acids such as lysine (K), arginine (R), and histidine (H), which contribute positively charged side chains. To further support this, we analyzed the basic amino acid composition of these XTH proteins and found that the proportion of basic residues ranged from 9.41% to 19.39% (Table 1), reinforcing the classification of these proteins as basic. Subcellular localization (Sub-Loc) analysis suggested that all CilXTH genes were predominantly localized in the cell wall, with 14 genes also found in the cytoplasm and two in the chloroplast (Table 1).
Based on the sequence characteristics and gene structures, all CilXTH genes were divided into four groups (Group a–Group d) (Figure 1). The length of the Glyco_hydro_16 domain ranged from 85 to 299 aa, and the XET_C domain ranged from 300 to 351 aa (Figure 2). The gene structures and organization analysis indicated that 10 out of 12 members in Group a contained 4 or 5 exons, while CilXTH15 contained 8 exons, and CilXTH36 contained 10 exons (Figure 2). Meanwhile, 13 out of 17 members in Group b contained 3 or 4 exons, but CilXTH38 contained 10 exons. For nine genes in Group c, six genes contained four exons, one gene contained five exons, and two genes contained six exons. These genes with longer sequences or more exons always contained other domains, except for the Glyco_hydro_16 and XET_C domains. For example, CilXTH35 contained an Rps17e-like domain, and CilXTH38 contained a 56 kDa selenium-binding protein (SBP56) domain (Figure 1).
In this study, a total of 15 β strands and four α helices were detected among these XTH genes, with catalytic sites mainly positioned between the β7 and β8 strands. The Glyco_hydro_16 domain was located from α1 to α2 (85–299 aa), and the XET_C domain was located from η6 to α2 (300–351 aa) (Figure 1). Multiple sequence alignment revealed that the amino acid residues were relatively conserved at the start and end positions of α helices or β strands, or on their connected positions. For example, within the Glyco_hydro_16 domain, the amino acid residue F85 at the start position of α1 was highly conserved, and the amino acid residues F89 and W/F93 at the start and end positions of β1 were also highly conserved. Within β4-β6, the multiple conserved amino acid residues were identified: sites G116, F117, and S119 in β4; sites Y123, G126, I/M130, I132, K133, and L134 in β5; and VTAFY (143–147) and L/M148 in β6. In contrast, within the XET_C domain, only some sites in β15, η8, and α4 were relatively conserved (Figure 1). In total, the conservation of the Glyco_hydro_16 domain was higher than in the XET_C domain, and most conserved amino acids were located at the start or end of the structural transformation region.
However, the loop structures within the Glyco_hydro_16 domain were highly varied among CilXTH proteins, particularly Loop2. Secondary and tertiary structure predictions using ESPript and AlphaFold revealed that CilXTH proteins share a conserved sandwich structure (Figure 1, Figure 3, Figures S3 and S4). As shown in Figure 1, an insertion containing six amino acids exists in Loop1 among members in both Groups IIIA and IIIB, while this insertion contains only three amino acids in most Group I and II members (Figure 1). Interestingly, a longer Loop2 region consisting of seven amino acid residues was observed in the Group IIIA proteins, while the corresponding insertion consisted of only four amino acid residues in Group IIIB members, and only two amino acid residues (GN/D) in Group I and II members. Furthermore, two conserved residues, S250 and D255, in Loop3 of Group IIIA were different from Groups I, II, and IIIB (Figure 1). Overall, these findings provide compelling evidence that Group IIIA genes primarily exhibit transglycosylation (XET) activity, whereas Group IIIB and ancestral groups (Groups I and II) may function as strict or predominant XETs. The Loop2 region likely plays a crucial role in determining XEH versus XET activity.

3.2. Phylogenetic Analysis of XTH Genes in Pecan

A phylogenetic tree was constructed using 33 Arabidopsis and 38 pecan XTH genes based on previously established classification criteria [12]. These results were categorized into four groups: I, II, IIIA, and IIIB, containing 12, 17, 4, and 5 members, respectively, which correspond to Clusters a, b, c, and d (Figure 4). Notably, three pecan genes in Group I (CilXTH13, CilXTH35, and CilXTH36) clustered within ancestral XTH genes from Arabidopsis, including AtXTH1, AtXTH2, AtXTH3, and AtXTH11, indicating a potential conservation of an ancient function (Figure 4). The remaining Group I members exhibited clear orthologous relationships between Arabidopsis and pecan, mostly in one-to-one or one-to-two correspondence. A similar pattern of orthology was observed in Group IIIB (Figure 4). In contrast, Group II genes showed species-specific clustering, whereby Arabidopsis and pecan members formed distinct, separate clades, suggesting possible lineage-specific expansion or functional divergence. Groups I and II are generally associated with XET activity or dual enzymatic functions, acting as both XET and XEH enzymes [46]. Furthermore, two well-characterized Group IIIA genes from Arabidopsis, which encode predominant XEH activity, were grouped with four pecan genes (CilXTH8, CilXTH11, CilXTH26, and CilXTH37). Collectively, the phylogenetic classification of XTH genes appears to be closely associated with their enzymatic preferences (XET or XEH). These findings support the hypothesis that XEH activity may have evolved from XET function, highlighting a possible functional diversification within the XTH gene family.

3.3. Chromosome Distribution and Collinearity Analysis of CilXTH Genes

The 38 CilXTH genes were unevenly distributed across 15 chromosomes and one scaffold, with each chromosome containing 1–8 CilXTH genes. The expansion of the CilXTH genes was primarily driven by both tandem and segmental duplications. Specifically, 11 genes formed clusters on Chromosomes 12, 15, and 16, likely resulting from tandem duplication (Figure 5A). For example, CilXTH19, CilXTH20, and CilXTH21 were clustered and located on chr12, and they all belong to Group II. In contrast, a total of 29 CilXTH genes arose from segmental duplication, with most pairs belonging to the same phylogenetic group, except for CilXTH5/CilXTH17 (belonging to Groups I and IIIB, respectively) and CilXTH8/CilXTH24 (Groups IIIA and I) (Figure 5B; Table S1).
To further investigate the evolutionary relationships, synonymous substitution rates (Ks) were calculated for these collinear CilXTH gene pairs. A total of 16 pairs of CilXTH genes were detected, including 11 pairs of segmentally duplicated genes and 5 pairs of tandem duplicates. Among these segmental duplicated genes, four out of seven gene pairs in Group I and two out of three gene pairs in Group II exhibited higher Ks values than those of the Group IIIB genes, suggesting that these duplication events likely occurred earlier than those in Group IIIB. Among the Group I genes, segmental duplicates exhibited higher Ks values than tandem duplicates (Table S2), consistent with the idea that segmental duplications often result from ancient whole-genome or large-scale duplications, whereas tandem duplications are generally more recent and localized [47,48]. These patterns support the hypothesis that Group I and II genes are evolutionarily older. The Ka/Ks values of all XTH gene pairs ranged from 0.05 to 0.86 and were consistently below 1, indicating that these genes have undergone purifying selection. This suggests that non-synonymous mutations are selectively constrained, likely due to functional importance. The XTH gene family plays a critical role in cell wall remodeling, and the observed purifying selection likely reflects the evolutionary pressure to maintain essential biological functions across gene duplicates [47,48].

3.4. Expression Profiles of CilXTH Genes in Different Tissues and Pecan Varieties

To explore the spatial and varietal expression patterns of CilXTH genes, transcriptomic data from five distinct pecan tissues—embryo, pericarp, peel, stem, and leaves—were analyzed [27,37]. Among the 31 CilXTH genes with detectable expression levels (FPKM > 1), 10, 13, and 8 genes belong to Groups I, II, and III, respectively. K-means clustering further grouped these genes into six distinct expression clusters (A1–A6) based on their tissue-specific expression patterns (Figure 6A). Cluster A1 contained seven genes predominantly expressed in the stem, all of which were classified into Group I (Figure 6A and Table S3). Clusters A2 and A3 included seven and three genes, respectively, exhibiting high expression in developing pericarp, whereas Cluster A4 comprised six genes with preferential expression in the peel (Figure 6A and Table S6). Cluster A5 encompassed five genes specifically expressed in leaves, while three genes in Cluster A6 were predominantly expressed in the embryo (Figure 6A).
To further investigate expression divergence between pecan varieties, transcriptomic profiles of “KD” (thin-shelled) and “SX” (thick-shelled) cultivars were examined across three stages of shell development. Genes with FPKM values below 1 across all samples were excluded from this analysis, resulting in 31 genes retained for clustering. These genes were grouped into six temporal expression clusters (B1–B6) (Figure 6B). Cluster B1 included five genes with high expression during the middle and late stages of shell development in “SX” (SX2 and SX3), while B6 comprised five genes primarily expressed in the early stage of “SX” shell development (SX1). Clusters B4 and B2 contained six and five genes, respectively, that were highly expressed in the middle (KD2) and late (KD3) developmental stages of the “KD” shell. Cluster B5 included seven genes showing high expression at the early developmental stage in both varieties (KD1 and SX1). Notably, three genes in B3 exhibited dual specificity, being highly expressed in the middle stage of “KD” (KD2) and the early stage of “SX” (SX1), suggesting the possible conserved regulatory roles of CilXTH genes during shell cell division and expansion (Figure 6B).
A comparative analysis of tissue- and variety-specific expression clusters revealed additional insights. Six genes from the stem-specific Cluster A1 were also highly expressed in KD2 (Figure S5), and they were Group II members. This overlap implies that these genes may contribute to rapid cell elongation or expansion in both the stem and developing shells of the “KD” variety. Among the ten genes grouped in the pericarp-dominant Clusters A2 and A3, four genes (CilXTH2, CilXTH27, CilXTH34, and CilXTH38) were specifically expressed in SX2 and SX3 (Cluster B1), suggesting their roles in late-stage shell thickening of “SX”. Three genes (CilXTH6, CilXTH10, and CilXTH22) were preferentially expressed in KD3 (Cluster B2). Two genes (CilXTH4 and CilXTH18) showed early expression in both “KD” and “SX” (Cluster B5). One gene, CilXTH35, was uniquely expressed in the early shell development stage of “SX” (SX1) (Figure 6B and Figure S6). Collectively, these findings highlight the differential and dynamic expression of the CilXTH genes in relation to tissue types and cultivar-specific shell development stages, suggesting their potential roles in modulating shell structure through tissue-specific cell wall remodeling.

3.5. Gene Regulatory Network of CilXTH Genes in Pecan Shell Development

Co-expression network analysis was carried out for all expressed genes during pecan shell development. By combining the co-expression network analysis results with our unpublished results, we proposed two regulatory networks of CilXTH genes during pecan shell development. In total, 5 CilXTH genes were co-expressed with 2321 genes and transcription factors, predominantly expressed in KD (thin-shelled pecan) during the early development stage (Figure 7A). GO enrichment analysis suggested that these genes were mainly involved in RNA modification (GO: 0009451), hydrolase activity (GO: 0016787), and the cytoplasm (GO: 0005737) (Figure S7). As shown in Figure 7A, CilXTH14/18/37 were co-expressed with several cell expansion and growth-related genes, such as FLA17 (fasciclin-like arabinogalactan protein 17), MAP70, CESA1/3, and LTP (lipid-transfer protein). In addition, the expression of CilXTH14/18/37 was regulated by several transcription factors, such as ERF12, NAC110, bHLH071, MYB3, and WD40 (Figure 7A). Meanwhile, several plant hormone-related genes may also regulate the expression of CilXTH14/18/37, such as RGL2, AUX1, ARF3, and ABF2 (Figure 7A). Conversely, CilXTH2/27/34 were highly expressed in SX (thick-shelled pecan) during the middle and late development stage (Figure 7B). A total of 3590 genes were involved in this process, and GO enrichment analysis revealed these genes were involved in transferase activity (GO: 0016740), catalytic complex (GO: 1902494), and plant-type SCW biogenesis (GO: 0009834) (Figure S8). The network showed that several SCW biosynthesis-related genes, such as IRX3, C4H, CAD, and 4CL1, and plant hormone-related genes, such as PYL4, SCL1, GA2ox1, and IAA10, were co-expressed in this cluster (Figure 7B). In addition, the expression of CilXTH2/27/34/36 genes was regulated by ANAC080, BLH1, ABF2, MADS57, and WRKY65 (Figure 7B). Moreover, the cis-regulatory element analysis revealed that the expression of all CilXTH genes was induced by auxin, GA, or ABA. Interestingly, the ATHB-2 element was only detected in the promoters of Group I, II, and IIIB genes. While the elements of Dof, bZIP, MYB, and E2F were present in nearly all the CilXTH genes (Figure 7C).

4. Discussion

Xyloglucan endotransglucosylase/hydrolase (XTH) enzymes play a crucial role in modifying plant cell walls, regulating processes such as root elongation, stress responses, fruit maturation, and SCW development [49,50,51]. These enzymes function by cleaving and reorganizing xyloglucan (XyG) polymers, thus modulating the structure and integrity of the cell wall. XTH-mediated remodeling influences cell wall extensibility, structural plasticity, and SCW biosynthesis across diverse plant species [20,52,53].
In this study, we identified 38 XTH genes in Carya illinoinensis, a number comparable to those reported in other plant species, such as Physcomitrella patens (32 genes), Arabidopsis thaliana (33 genes), Oryza sativa (29 genes), and Cucumis sativus (29 genes) [12,13,54,55]. Phylogenetic analysis categorized these CilXTH genes into four distinct groups: I, II, IIIA, and IIIB (Figure 4). Groups I, II, and IIIB generally possess both xyloglucan endotransglucosylase (XET) and xyloglucan endohydrolase (XEH) activities, whereas Group IIIA is primarily associated with XEH activity (Figure 1 and Figure 4). This classification is consistent with phylogenetic groupings observed in other plant species, such as strawberry, melon, and Selaginella kraussiana [46,56,57,58]. Moreover, synonymous substitution (Ks) analysis among these CilXTH gene pairs supports an evolutionary trajectory in which Group I represents a more ancestral clade.
Functionally, XET activity primarily facilitates the integration of newly synthesized XyG into cell wall polysaccharides or the depolymerization of cell walls through XyG cleavage [10]. XET is considered a key agent in regulating wall expansion, wall strengthening, and many other aspects of cell wall biosynthesis [59]. In contrast, XEH activity hydrolyzes β−1,4 glycosidic bonds within XyG, promoting the irreversible degradation of XyG molecules involved in processes such as cell wall expansion, remodeling, and repair [10,60]. Despite the conservation of catalytic domains across XTHs, the divergence between XET and XEH functionalities is largely determined by their three-dimensional structural features (Figure 3, Figures S3 and S4). Specifically, sequence variations in Loop2 have been extensively investigated, with findings indicating that the length and amino acid composition of Loop2 are highly conserved across different species, including Arabidopsis, rice, nasturtium (Tropaeolum majus), and some other species [61].
Importantly, XyG is now recognized not only as a flexible load-bearing polysaccharide but also as a key hotspot and crosslinking agent in the plant cell wall [62]. XyG molecules bind tightly to cellulose microfibrils and serve as molecular tethers that regulate cell wall mechanical properties [63]. These “hotspots”—defined as localized regions of concentrated XyG–cellulose interactions—are essential sites where XTHs mediate cell wall plasticity through transglycosylation or hydrolytic cleavage [64]. XTH enzymes facilitate dynamic crosslinking between cellulose-bound XyG chains, thereby reinforcing or loosening the wall depending on developmental or environmental cues. Several studies have demonstrated their roles in cell wall modification, including maize XTH1, soybean BRU1, and Arabidopsis AtXTH31/32 [65,66,67]. Furthermore, XTHs contribute to SCW formation, maintaining cell wall thickness and integrity [18]. In poplar, the expression pattern of XET16A suggests that XET enzymes primarily function in primary wall restructuring and SCW deposition [59].
Our expression analyses revealed that CilXTH genes were involved in both primary wall expansion and SCW deposition. Several CilXTH genes exhibited cultivar-specific expression patterns: Cluster B1 and B6 genes were predominantly expressed in the thick shells of the “SX” cultivar, while B2 and B4 genes were dominant in the thin shells of the “KD” cultivar (Figure 6). Integrative analyses of shell phenotype, gene expression, and co-expression networks identified core XTH genes with potential functional relevance. XTH2, XTH27, XTH34, and XTH36 were likely involved in regulating SCW thickening through crosslinking activity and cellulose reinforcement, possibly acting at xyloglucan hotspots within the developing shell (Figure 5). Meanwhile, XTH14, XTH18, and XTH37 were more closely associated with fruit elongation, likely mediating XET activity in regions of active cell wall expansion (Figure 7).
Future research should focus on elucidating the transcriptional and post-transcriptional regulatory mechanisms governing XTH gene function. This includes exploring the roles of upstream transcription factors such as NAC, MYB, and ARF, as well as signaling pathways like auxin and GA-mediated regulation. Additionally, studies should investigate the functional redundancy and species-specific regulatory networks that underpin XTH-mediated cell wall plasticity, particularly in response to environmental factors such as mechanical stress and temperature fluctuations. Collectively, these findings underscore the functional diversification of CilXTH genes and highlight their key regulatory roles in pecan fruit development, particularly through modulating xyloglucan-mediated crosslinking and cell wall remodeling at critical structural domains.

5. Conclusions

This study presents a comprehensive genome-wide analysis of the XTH gene family in pecan, revealing their evolutionary divergence, structural features, and functional implications. Expression and co-expression analyses suggest that specific CilXTH genes regulate pecan shell development by modulating cell wall remodeling and hormonal pathways. These findings provide new insights into the molecular mechanisms underlying pecan shell thickness and lay a foundation for future functional validation studies. Further investigations into transcriptional regulation and environmental interactions will be essential for fully elucidating the roles of CilXTH genes in pecan shell development and potential applications in breeding programs.

Supplementary Materials

The following supporting information can be downloaded at https://www.mdpi.com/article/10.3390/horticulturae11060609/s1, Figure S1: Comparison of fruit morphology and cross-sectional staining between KD and SX; Figure S2: The overlapped candidate XTH genes identified by both an HMMER and BLASTP searches for pecan; Figure S3: Tertiary structure predictions of XTH proteins from Groups a–d in pecan; Figure S4: The model confidence of CilXTH14, CilXTH22, and CilXTH37; Figure S5: The overlapped genes between Clusters A1, B2, and B4; Figure S6: The overlapped genes between B1, B2, B5, and A2+A3; Figure S7: GO enrichment analysis of 2321 genes and transcription factors in pecan; Figure S8: GO enrichment analysis of 3590 genes in pecan; Table S1: Gene duplication status of XTH genes in pecan; Table S2: Ka and Ks analysis of CilXTH gene family; Table S3: Clustering and Tissue-Specific Expression of A1-A6 cluster genes in pecan; Table S4: Clustering and Expression Profiles of CilXTH genes in KD and SX during seed development.

Author Contributions

Methodology, M.W.; Data curation, M.W. and Z.Z.; Writing—original draft, Z.Z. and A.Y.; Visualization, J.S. and F.M.; Provide materials, X.X.; Funding acquisition, A.Y. and A.L.; Writing—review and editing, A.Y. and A.L. All authors have read and agreed to the published version of the manuscript.

Funding

This work was supported by the Yunnan Fundamental Research Projects (202301AT070642, 202201AU070205), the National Natural Science Foundation of China (NSFC, 32360475, 32261143461, 32372135), the Forestry Innovation Programs of Southwest Forestry University (LXXK-2023Z02), and the Fund of Yunnan Key Laboratory of Crop Wild Relatives Omics (CWR-2024-05). We thank all the individuals who have helped us in this study. No conflicts of interest are declared.

Data Availability Statement

The original contributions presented in this study are included in the article; further inquiries can be directed to the corresponding authors.

Acknowledgments

We thank Lihong Xiao (Zhejiang A and F University) for kindly supplying the pecan reference genome used in this research.

Conflicts of Interest

The authors declare no conflicts of interest.

Abbreviations

The following abbreviations are used in this manuscript:
XTHXyloglucan endotransglucosylases/hydrolases
SCWSecondary cell wall
XyGXyloglucan
XETXyloglucan endotransglucosylase
XEHXyloglucan hydrolase
GH16Glycoside hydrolase family 16
KDCaddo
SXShaoxing
DAPDays after pollination
MWMolecular weight
pIIsoelectric point
MLMaximum Likelihood
kDakiloDalton
aaAmino acids
CDSCoding DNA Sequence
CWCell wall
CCytoplasm
CPChloroplast
SBPSelenium-binding protein
MbMegabases
GAGibberellic Acid
ABAAbscisic Acid

References

  1. Le Gall, H.; Philippe, F.; Domon, J.M.; Gillet, F.; Pelloux, J.; Rayon, C. Cell Wall Metabolism in Response to Abiotic Stress. Plants 2015, 4, 112–166. [Google Scholar] [CrossRef] [PubMed]
  2. Cosgrove, D.J. Growth of the Plant Cell Wall. Nat. Rev. Mol. Cell Biol. 2005, 6, 850–861. [Google Scholar] [CrossRef]
  3. Xu, H.; Giannetti, A.; Sugiyama, Y.; Zheng, W.; Schneider, R.; Watanabe, Y.; Oda, Y.; Persson, S. Secondary Cell Wall Patterning-Connecting the Dots, Pits and Helices. Open Biol. 2022, 12, 210208. [Google Scholar] [CrossRef]
  4. Cosgrove, D.J.; Jarvis, M.C. Comparative Structure and Biomechanics of Plant Primary and Secondary Cell Walls. Front. Plant Sci. 2012, 3, 204. [Google Scholar] [CrossRef]
  5. von Schantz, L.; Gullfot, F.; Scheer, S.; Filonova, L.; Cicortas Gunnarsson, L.; Flint, J.E.; Daniel, G.; Nordberg-Karlsson, E.; Brumer, H.; Ohlin, M. Affinity Maturation Generates Greatly Improved Xyloglucan-Specific Carbohydrate Binding Modules. BMC Biotechnol. 2009, 9, 92. [Google Scholar] [CrossRef]
  6. Takahashi, D.; Johnson, K.L.; Hao, P.; Tuong, T.; Erban, A.; Sampathkumar, A.; Bacic, A.; Livingston, D.P., 3rd; Kopka, J.; Kuroha, T.; et al. Cell Wall Modification by the Xyloglucan Endotransglucosylase/Hydrolase XTH19 Influences Freezing Tolerance after Cold and Sub-Zero Acclimation. Plant Cell Environ. 2021, 44, 915–930. [Google Scholar] [CrossRef]
  7. Li, Y.; Hua, J.; Hou, X.; Qi, N.; Li, C.; Wang, C.; Yao, Y.; Huang, D.; Zhang, H.; Liao, W. Brassinosteroids Is Involved in Methane-Induced Adventitious Root Formation Via Inducing Cell Wall Relaxation in Marigold. BMC Plant Biol. 2023, 23, 2. [Google Scholar] [CrossRef] [PubMed]
  8. Hrmova, M.; Stratilová, B.; Stratilová, E. Broad Specific Xyloglucan:Xyloglucosyl Transferases Are Formidable Players in the Re-Modelling of Plant Cell Wall Structures. Int. J. Mol. Sci. 2022, 23, 1656. [Google Scholar] [CrossRef]
  9. Holland, C.; Simmons, T.J.; Meulewaeter, F.; Hudson, A.; Fry, S.C. Three Highly Acidic Equisetum XTHs Differ from Hetero-Trans-β-Glucanase in Donor Substrate Specificity and Are Predominantly Xyloglucan Homo-Transglucosylases. J. Plant Physiol. 2020, 251, 153210. [Google Scholar] [CrossRef] [PubMed]
  10. Eklöf, J.M.; Brumer, H. The Xth Gene Family: An Update on Enzyme Structure, Function, and Phylogeny in Xyloglucan Remodeling. Plant Physiol. 2010, 153, 456–466. [Google Scholar] [CrossRef]
  11. Viborg, A.H.; Terrapon, N.; Lombard, V.; Michel, G.; Czjzek, M.; Henrissat, B.; Brumer, H. A Subfamily Roadmap of the Evolutionarily Diverse Glycoside Hydrolase Family 16 (GH16). J. Biol. Chem. 2019, 294, 15973–15986. [Google Scholar] [CrossRef] [PubMed]
  12. Yokoyama, R.; Nishitani, K. A Comprehensive Expression Analysis of All Members of a Gene Family Encoding Cell-Wall Enzymes Allowed Us to Predict Cis-Regulatory Regions Involved in Cell-Wall Construction in Specific Organs of Arabidopsis. Plant Cell Physiol. 2001, 42, 1025–1033. [Google Scholar] [CrossRef] [PubMed]
  13. Uozu, S.; Tanaka-Ueguchi, M.; Kitano, H.; Hattori, K.; Matsuoka, M. Characterization of XET-Related Genes of Rice. Plant Physiol. 2000, 122, 853–859. [Google Scholar] [CrossRef] [PubMed]
  14. Cheng, Z.; Zhang, X.; Yao, W.; Gao, Y.; Zhao, K.; Guo, Q.; Zhou, B.; Jiang, T. Genome-Wide Identification and Expression Analysis of the Xyloglucan Endotransglucosylase/Hydrolase Gene Family in Poplar. BMC Genomics 2021, 22, 804. [Google Scholar] [CrossRef]
  15. Dhar, S.; Kim, J.; Yoon, E.K.; Jang, S.; Ko, K.; Lim, J. Short-Root Controls Cell Elongation in the Etiolated Arabidopsis hypocotyl. Mol. Cells 2022, 45, 243–256. [Google Scholar] [CrossRef]
  16. Hyodo, H.; Yamakawa, S.; Takeda, Y.; Tsuduki, M.; Yokota, A.; Nishitani, K.; Kohchi, T. Active Gene Expression of a Xyloglucan Endotransglucosylase/Hydrolase Gene, XTH9, in Inflorescence Apices Is Related to Cell Elongation in Arabidopsis thaliana. Plant Mol. Biol. 2003, 52, 473–482. [Google Scholar] [CrossRef]
  17. Li, Y.; Zheng, X.; Wang, C.; Hou, D.; Li, T.; Li, D.; Ma, C.; Sun, Z.; Tian, Y. Pear Xyloglucan Endotransglucosylase/Hydrolases Pcbru1 Promotes Stem Growth through Regulating Cell Wall Elongation. Plant Sci. 2021, 312, 111026. [Google Scholar] [CrossRef]
  18. Kushwah, S.; Banasiak, A.; Nishikubo, N.; Derba-Maceluch, M.; Majda, M.; Endo, S.; Kumar, V.; Gomez, L.; Gorzsas, A.; McQueen-Mason, S.; et al. Arabidopsis XTH4 and XTH9 Contribute to Wood Cell Expansion and Secondary Wall Formation. Plant Physiol. 2020, 182, 1946–1965. [Google Scholar] [CrossRef]
  19. Vissenberg, K.; Oyama, M.; Osato, Y.; Yokoyama, R.; Verbelen, J.P.; Nishitani, K. Differential Expression of AtXTH17, AtXTH18, AtXTH19 and AtXTH20 Genes in Arabidopsis Roots. Physiological Roles in Specification in Cell Wall Construction. Plant Cell Physiol. 2005, 46, 192–200. [Google Scholar] [CrossRef]
  20. Matsui, A.; Yokoyama, R.; Seki, M.; Ito, T.; Shinozaki, K.; Takahashi, T.; Komeda, Y.; Nishitani, K. AtXTH27 Plays an Essential Role in Cell Wall Modification During the Development of Tracheary Elements. Plant J. 2005, 42, 525–534. [Google Scholar] [CrossRef]
  21. Nishikubo, N.; Takahashi, J.; Roos, A.A.; Derba-Maceluch, M.; Piens, K.; Brumer, H.; Teeri, T.T.; Stålbrand, H.; Mellerowicz, E.J. Xyloglucan Endo-Transglycosylase-Mediated Xyloglucan Rearrangements in Developing Wood of Hybrid Aspen. Plant Physiol. 2011, 155, 399–413. [Google Scholar] [CrossRef] [PubMed]
  22. Nakaune, M.; Hanada, A.; Yin, Y.G.; Matsukura, C.; Yamaguchi, S.; Ezura, H. Molecular and Physiological Dissection of Enhanced Seed Germination Using Short-Term Low-Concentration Salt Seed Priming in Tomato. Plant Physiol. Biochem. 2012, 52, 28–37. [Google Scholar] [CrossRef] [PubMed]
  23. Fu, M.M.; Cao, F.; Qiu, C.W.; Liu, C.; Tong, T.; Feng, X.; Cai, S.; Chen, Z.H.; Wu, F. Xyloglucan Endotransglucosylase-Hydrolase 1 Is a Negative Regulator of Drought Tolerance in Barley Via Modulating Lignin Biosynthesis and Stomatal Closure. Plant Physiol. Biochem. 2024, 216, 109171. [Google Scholar] [CrossRef]
  24. Badenes, M.L.; Byrne, D.H. Fruit Breeding; Springer Science & Business Media: Berlin/Heidelberg, Germany, 2012; Volume 8. [Google Scholar]
  25. Toro-Vazquez, J.F.; Charó-Alonso, M.A.; Pérez-Briceño, F. Fatty Acid Composition and Its Relationship with Physicochemical Properties of Pecan (Carya illinoensis) Oil. J. Am. Oil Chem. Soc. 1999, 76, 957–965. [Google Scholar] [CrossRef]
  26. Liu, J.; Xue, T.; Ren, L.; Cui, M.; Jiang, T.; Yang, X. Study on Pecan Seed Germination Influenced by Seed Endocarp. Open Life Sci. 2022, 17, 851–855. [Google Scholar] [CrossRef]
  27. Xiao, L.; Yu, M.; Zhang, Y.; Hu, J.; Zhang, R.; Wang, J.; Guo, H.; Zhang, H.; Guo, X.; Deng, T.; et al. Chromosome-Scale Assembly Reveals Asymmetric Paleo-Subgenome Evolution and Targets for the Acceleration of Fungal Resistance Breeding in the Nut Crop, Pecan. Plant Commun. 2021, 2, 100247. [Google Scholar] [CrossRef] [PubMed]
  28. Potter, S.C.; Luciani, A.; Eddy, S.R.; Park, Y.; Lopez, R.; Finn, R.D. Hmmer Web Server: 2018 Update. Nucleic Acids Res. 2018, 46, W200–W204. [Google Scholar] [CrossRef]
  29. Gasteiger, E.; Gattiker, A.; Hoogland, C.; Ivanyi, I.; Appel, R.D.; Bairoch, A. Expasy: The Proteomics Server for in-Depth Protein Knowledge and Analysis. Nucleic Acids Res. 2003, 31, 3784–3788. [Google Scholar] [CrossRef]
  30. Chou, K.C.; Shen, H.B. Plant-Mploc: A Top-Down Strategy to Augment the Power for Predicting Plant Protein Subcellular Localization. PLoS ONE 2010, 5, e11335. [Google Scholar] [CrossRef]
  31. Høie, M.H.; Kiehl, E.N.; Petersen, B.; Nielsen, M.; Winther, O.; Nielsen, H.; Hallgren, J.; Marcatili, P. Netsurfp-3.0: Accurate and Fast Prediction of Protein Structural Features by Protein Language Models and Deep Learning. Nucleic Acids Res. 2022, 50, W510–W515. [Google Scholar] [CrossRef]
  32. Jumper, J.; Evans, R.; Pritzel, A.; Green, T.; Figurnov, M.; Ronneberger, O.; Tunyasuvunakool, K.; Bates, R.; Žídek, A.; Potapenko, A.; et al. Highly Accurate Protein Structure Prediction with Alphafold. Nature 2021, 596, 583–589. [Google Scholar] [CrossRef] [PubMed]
  33. Letunic, I.; Bork, P. Interactive Tree of Life (Itol) V5: An Online Tool for Phylogenetic Tree Display and Annotation. Nucleic Acids Res. 2021, 49, W293–W296. [Google Scholar] [CrossRef] [PubMed]
  34. 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] [PubMed]
  35. Wang, Y.; Tang, H.; Debarry, J.D.; Tan, X.; Li, J.; Wang, X.; Lee, T.H.; Jin, H.; Marler, B.; Guo, H.; et al. Mcscanx: A Toolkit for Detection and Evolutionary Analysis of Gene Synteny and Collinearity. Nucleic Acids Res. 2012, 7, e49. [Google Scholar] [CrossRef]
  36. Sleator, R.D. Jcvi-Syn3.0—A Synthetic Genome Stripped Bare! Bioengineered 2016, 7, 53–56. [Google Scholar] [CrossRef]
  37. Sun, P.; Jiao, B.; Yang, Y.; Shan, L.; Li, T.; Li, X.; Xi, Z.; Wang, X.; Liu, J. Wgdi: A User-Friendly Toolkit for Evolutionary Analyses of Whole-Genome Duplications and Ancestral Karyotypes. Mol. Plant 2022, 15, 1841–1851. [Google Scholar] [CrossRef] [PubMed]
  38. Huang, Y.; Xiao, L.; Zhang, Z.; Zhang, R.; Wang, Z.; Huang, C.; Huang, R.; Luan, Y.; Fan, T.; Wang, J.; et al. The Genomes of Pecan and Chinese Hickory Provide Insights into Carya Evolution and Nut Nutrition. Gigascience 2019, 8, giz036. [Google Scholar] [CrossRef]
  39. Xanthopoulou, A.; Moysiadis, T.; Bazakos, C.; Karagiannis, E.; Karamichali, I.; Stamatakis, G.; Samiotaki, M.; Manioudaki, M.; Michailidis, M.; Madesis, P.; et al. The Perennial Fruit Tree Proteogenomics Atlas: A Spatial Map of the Sweet Cherry Proteome and Transcriptome. Plant J. 2022, 109, 1319–1336. [Google Scholar] [CrossRef]
  40. Brandine, G.S.; Smith, A.D. Falco: High-Speed Fastqc Emulation for Quality Control of Sequencing Data. F1000Research 2019, 8, 1874. [Google Scholar] [CrossRef]
  41. Bolger, A.M.; Lohse, M.; Usadel, B. Trimmomatic: A Flexible Trimmer for Illumina Sequence Data. Bioinformatics 2014, 30, 2114–2120. [Google Scholar] [CrossRef]
  42. Kim, D.; Paggi, J.M.; Park, C.; Bennett, C.; Salzberg, S.L. Graph-Based Genome Alignment and Genotyping with Hisat2 and Hisat-Genotype. Nat. Biotechnol. 2019, 37, 907–915. [Google Scholar] [CrossRef] [PubMed]
  43. Pertea, M.; Pertea, G.M.; Antonescu, C.M.; Chang, T.C.; Mendell, J.T.; Salzberg, S.L. Stringtie Enables Improved Reconstruction of a Transcriptome from Rna-Seq Reads. Nat. Biotechnol. 2015, 33, 290–295. [Google Scholar] [CrossRef] [PubMed]
  44. Huynh-Thu, V.A.; Irrthum, A.; Wehenkel, L.; Geurts, P. Inferring Regulatory Networks from Expression Data Using Tree-Based Methods. PLoS ONE 2010, 5, e12776. [Google Scholar] [CrossRef]
  45. Shannon, P.; Markiel, A.; Ozier, O.; Baliga, N.S.; Wang, J.T.; Ramage, D.; Amin, N.; Schwikowski, B.; Ideker, T. Cytoscape: A Software Environment for Integrated Models of Biomolecular Interaction Networks. Genome Res. 2003, 13, 2498–2504. [Google Scholar] [CrossRef]
  46. Stratilová, B.; Šesták, S.; Stratilová, E.; Vadinová, K.; Kozmon, S.; Hrmova, M. Engineering of Substrate Specificity in a Plant Cell-Wall Modifying Enzyme through Alterations of Carboxyl-Terminal Amino Acid Residues. Plant J. 2023, 116, 1529–1544. [Google Scholar] [CrossRef] [PubMed]
  47. Blanc, G.; Wolfe, K.H. Widespread Paleopolyploidy in Model Plant Species Inferred from Age Distributions of Duplicate Genes. Plant Cell 2004, 16, 1667–1678. [Google Scholar] [CrossRef]
  48. Lynch, M.; Conery, J.S. The Evolutionary Fate and Consequences of Duplicate Genes. Science 2000, 290, 1151–1155. [Google Scholar] [CrossRef]
  49. Cao, J.; Lv, Y.; Li, X. Interspaced Repeat Sequences Confer the Regulatory Functions of AtXTH10, Important for Root Growth in Arabidopsis. Plants 2019, 8, 130. [Google Scholar] [CrossRef]
  50. Gao, Y.; Wang, L.; Li, D.; Qi, D.; Fang, F.; Luo, Y.; Zhang, H.; Zhang, S. Genome-Wide Characterization of the Xyloglucan Endotransglucosylase/Hydrolase Family Genes and Their Response to Plant Hormone in Sugar Beet. Plant Physiol. Biochem. 2024, 206, 108239. [Google Scholar] [CrossRef]
  51. Zhang, Z.; Wang, N.; Jiang, S.; Xu, H.; Wang, Y.; Wang, C.; Li, M.; Liu, J.; Qu, C.; Liu, W.; et al. Analysis of the Xyloglucan Endotransglucosylase/Hydrolase Gene Family During Apple Fruit Ripening and Softening. J. Agric. Food Chem. 2017, 65, 429–434. [Google Scholar] [CrossRef]
  52. Van Sandt, V.S.; Suslov, D.; Verbelen, J.P.; Vissenberg, K. Xyloglucan Endotransglucosylase Activity Loosens a Plant Cell Wall. Ann. Bot. 2007, 100, 1467–1473. [Google Scholar] [CrossRef]
  53. Derba-Maceluch, M.; Awano, T.; Takahashi, J.; Lucenius, J.; Ratke, C.; Kontro, I.; Busse-Wicher, M.; Kosik, O.; Tanaka, R.; Winzéll, A.; et al. Suppression of Xylan Endotransglycosylase PtxtXyn10A Affects Cellulose Microfibril Angle in Secondary Wall in Aspen Wood. New Phytol. 2015, 205, 666–681. [Google Scholar] [CrossRef] [PubMed]
  54. Yokoyama, R.; Uwagaki, Y.; Sasaki, H.; Harada, T.; Hiwatashi, Y.; Hasebe, M.; Nishitani, K. Biological Implications of the Occurrence of 32 Members of the XTH (Xyloglucan Endotransglucosylase/Hydrolase) Family of Proteins in the Bryophyte Physcomitrella patens. Plant J. 2010, 64, 645–656. [Google Scholar] [CrossRef] [PubMed]
  55. Bhatti, M.H.T.; Han, S.; Sami, A.; Haider, M.Z.; Shafiq, M.; Ali, M.; Ahmad, S.; Ali, Q.; Sabir, I.A.; Manzoor, M.A. Genome-Wide Analysis of XTH Gene Family in Cucumber (Cucumis sativus) against Different Insecticides to Enhance Defense Mechanism. Plant Stress 2024, 13, 100538. [Google Scholar] [CrossRef]
  56. Witasari, L.D.; Huang, F.C.; Hoffmann, T.; Rozhon, W.; Fry, S.C.; Schwab, W. Higher Expression of the Strawberry Xyloglucan Endotransglucosylase/Hydrolase Genes FvXTH9 and FvXTH6 Accelerates Fruit Ripening. Plant J. 2019, 100, 1237–1253. [Google Scholar] [CrossRef]
  57. Zhao, S.; Kang, Y.; Lin, Y.; Zheng, X.; Wu, Y.; Yang, Z. A Genome-Wide Identification and Expression Analysis of the Xyloglucan Endotransglucosylase/Hydrolase Gene Family in Melon (Cucumis melo L.). Horticulturae 2024, 10, 1017. [Google Scholar] [CrossRef]
  58. Van Sandt, V.S.; Guisez, Y.; Verbelen, J.P.; Vissenberg, K. Analysis of a Xyloglucan Endotransglycosylase/Hydrolase (XTH) from the Lycopodiophyte Selaginella Kraussiana Suggests That XTH Sequence Characteristics and Function Are Highly Conserved During the Evolution of Vascular Plants. J. Exp. Bot. 2006, 57, 2909–2922. [Google Scholar] [CrossRef]
  59. Bourquin, V.; Nishikubo, N.; Abe, H.; Brumer, H.; Denman, S.; Eklund, M.; Christiernin, M.; Teeri, T.T.; Sundberg, B.; Mellerowicz, E.J. Xyloglucan Endotransglycosylases Have a Function During the Formation of Secondary Cell Walls of Vascular Tissues. Plant Cell 2002, 14, 3073–3088. [Google Scholar] [CrossRef]
  60. Seale, M. Cell Wall Remodeling During Wood Development. Plant Physiol. 2020, 182, 1800–1801. [Google Scholar] [CrossRef]
  61. Baumann, M.J.; Eklöf, J.M.; Michel, G.; Kallas, A.M.; Teeri, T.T.; Czjzek, M.; Brumer, H., 3rd. Structural Evidence for the Evolution of Xyloglucanase Activity from Xyloglucan Endo-Transglycosylases: Biological Implications for Cell Wall Metabolism. Plant Cell 2007, 19, 1947–1963. [Google Scholar] [CrossRef]
  62. Park, Y.B.; Cosgrove, D.J. Xyloglucan and its Interactions with Other Components of the Growing Cell Wall. Plant Cell Physiol. 2015, 56, 180–194. [Google Scholar] [CrossRef] [PubMed]
  63. Cosgrove, D.J. Wall structure and wall loosening. A look backwards and forwards. Plant Physiol. 2001, 125, 131–134. [Google Scholar] [CrossRef] [PubMed]
  64. Takahisa, H.; Rumi, K. Functions of Xyloglucan in Plant Cells. Mol. Plant 2011, 4, 17–24. [Google Scholar]
  65. Genovesi, V.; Fornalé, S.; Fry, S.C.; Ruel, K.; Ferrer, P.; Encina, A.; Sonbol, F.M.; Bosch, J.; Puigdomènech, P.; Rigau, J.; et al. Zmxth1, a New Xyloglucan Endotransglucosylase/Hydrolase in Maize, Affects Cell Wall Structure and Composition in Arabidopsis thaliana. J. Exp. Bot. 2008, 59, 875–889. [Google Scholar] [CrossRef]
  66. Oh, M.-H.; Romanow, W.G.; Smith, R.C.; Zamski, E.; Sasse, J.; Clouse, S.D. Soybean Bru1 Encodes a Functional Xyloglucan Endotransglycosylase That Is Highly Expressed in Inner Epicotyl Tissues During Brassinosteroid-Promoted Elongation. Plant Cell Physiol. 1998, 39, 124–130. [Google Scholar] [CrossRef]
  67. Kaewthai, N.; Gendre, D.; Eklöf, J.M.; Ibatullin, F.M.; Ezcurra, I.; Bhalerao, R.P.; Brumer, H. Group III-a XTH Genes of Arabidopsis Encode Predominant Xyloglucan Endohydrolases That Are Dispensable for Normal Growth. Plant Physiol. 2013, 161, 440–454. [Google Scholar] [CrossRef]
Horticulturae 11 00609 g001
Figure 1. The multi-sequence alignment of the CilXTH protein family. The multiple sequence alignment reveals conserved residues (highlighted in red) among homologous sequences, with secondary structural elements—such as loops and the ExDxE domain—annotated to delineate functional regions. The diagram indicates α-helices (depicted as spirals), β-sheets (shown as arrows), and N-glycosylation sites. The blocks of different colors on the left side represent different XTH groups.
Figure 1. The multi-sequence alignment of the CilXTH protein family. The multiple sequence alignment reveals conserved residues (highlighted in red) among homologous sequences, with secondary structural elements—such as loops and the ExDxE domain—annotated to delineate functional regions. The diagram indicates α-helices (depicted as spirals), β-sheets (shown as arrows), and N-glycosylation sites. The blocks of different colors on the left side represent different XTH groups.
Horticulturae 11 00609 g001
Horticulturae 11 00609 g002
Figure 2. The phylogenetic relationships, conserved domains, and gene structures of CiXTH genes in pecan. The phylogenetic tree clusters the CiXTH genes into four distinct groups: Group a (light yellow), Group b (light green), Group c (light purple), and Group d (pink). Conserved protein domains are shown in green, including the Glyco_hydro_16 and XET_C domains. The gene structures are displayed on the right, where blue boxes represent exons, and black lines represent introns. Gene lengths are scaled in kilobases (kb) from the 5′ to 3′ direction.
Figure 2. The phylogenetic relationships, conserved domains, and gene structures of CiXTH genes in pecan. The phylogenetic tree clusters the CiXTH genes into four distinct groups: Group a (light yellow), Group b (light green), Group c (light purple), and Group d (pink). Conserved protein domains are shown in green, including the Glyco_hydro_16 and XET_C domains. The gene structures are displayed on the right, where blue boxes represent exons, and black lines represent introns. Gene lengths are scaled in kilobases (kb) from the 5′ to 3′ direction.
Horticulturae 11 00609 g002
Horticulturae 11 00609 g003
Figure 3. A structural comparison of CilXTH proteins based on superimposed 3D models. CilXTH14 vs. CilXTH22 (left) and CilXTH22 vs. CilXTH37 (right). The conserved catalytic ExDxE domain is shown in cyan, while three structural loops—Loop1 (yellow), Loop2 (purple), and Loop3 (red)—are highlighted to illustrate conformational differences among the proteins. These structural variations may underlie functional divergence in enzymatic activity or substrate recognition.
Figure 3. A structural comparison of CilXTH proteins based on superimposed 3D models. CilXTH14 vs. CilXTH22 (left) and CilXTH22 vs. CilXTH37 (right). The conserved catalytic ExDxE domain is shown in cyan, while three structural loops—Loop1 (yellow), Loop2 (purple), and Loop3 (red)—are highlighted to illustrate conformational differences among the proteins. These structural variations may underlie functional divergence in enzymatic activity or substrate recognition.
Horticulturae 11 00609 g003
Horticulturae 11 00609 g004
Figure 4. A phylogenetic analysis of XTH genes from pecan and Arabidopsis. The phylogenetic tree was constructed using the Maximum Likelihood method based on the amino acid sequences of XTH genes. Bootstrap values (shown on the branches) represent the confidence levels from 1000 replicates. The XTH genes are classified into four major groups (I, II, IIIA, and IIIB) and highlighted in distinct colors. Group I is shown in blue, Group II in red, Group IIIA in teal, and Group IIIB in orange.
Figure 4. A phylogenetic analysis of XTH genes from pecan and Arabidopsis. The phylogenetic tree was constructed using the Maximum Likelihood method based on the amino acid sequences of XTH genes. Bootstrap values (shown on the branches) represent the confidence levels from 1000 replicates. The XTH genes are classified into four major groups (I, II, IIIA, and IIIB) and highlighted in distinct colors. Group I is shown in blue, Group II in red, Group IIIA in teal, and Group IIIB in orange.
Horticulturae 11 00609 g004
Horticulturae 11 00609 g005
Figure 5. The chromosomal distribution and gene duplication analysis of CilXTH genes in pecan. (A) The chromosomal distribution of CilXTH genes across pecan chromosomes. The CiXTH gene locations are marked in red. Chromosome lengths are scaled in megabases (Mb). (B) A schematic representation of gene duplication events among CiXTH genes. The colored ribbons indicate duplicated gene pairs, while the gray lines in the background denote syntenic blocks across the pecan genome.
Figure 5. The chromosomal distribution and gene duplication analysis of CilXTH genes in pecan. (A) The chromosomal distribution of CilXTH genes across pecan chromosomes. The CiXTH gene locations are marked in red. Chromosome lengths are scaled in megabases (Mb). (B) A schematic representation of gene duplication events among CiXTH genes. The colored ribbons indicate duplicated gene pairs, while the gray lines in the background denote syntenic blocks across the pecan genome.
Horticulturae 11 00609 g005
Horticulturae 11 00609 g006
Figure 6. The expression profiles of the CilXTH genes in pecan. (A) A heatmap showing the expression patterns of CilXTH genes across various pecan tissues, including embryo, pericarp, peel, stem, and leaves. The samples are grouped based on tissue type, and Z-score normalization is applied to indicate relative expression levels. (B) A heatmap illustrating the expression dynamics of the CilXTH genes at three developmental stages in two pecan varieties. KD1, KD2, and KD3 represent three shell development stages of the thin-shelled cultivar. SX1, SX2, and SX3 represent three shell development stages of the thick-shelled cultivar. Differences in expression patterns between developmental stages and varieties are highlighted, providing insights into the potential roles of the CilXTH genes in pecan growth and development.
Figure 6. The expression profiles of the CilXTH genes in pecan. (A) A heatmap showing the expression patterns of CilXTH genes across various pecan tissues, including embryo, pericarp, peel, stem, and leaves. The samples are grouped based on tissue type, and Z-score normalization is applied to indicate relative expression levels. (B) A heatmap illustrating the expression dynamics of the CilXTH genes at three developmental stages in two pecan varieties. KD1, KD2, and KD3 represent three shell development stages of the thin-shelled cultivar. SX1, SX2, and SX3 represent three shell development stages of the thick-shelled cultivar. Differences in expression patterns between developmental stages and varieties are highlighted, providing insights into the potential roles of the CilXTH genes in pecan growth and development.
Horticulturae 11 00609 g006
Horticulturae 11 00609 g007
Figure 7. Co-expression network and cis-regulatory element analysis of CilXTH genes. (A) The co-expression network of the CilXTH genes in the shell of KD (thin-shelled pecan) during the early developmental stage. The blue squares represent genes co-expressed with CilXTH, while the pink squares indicate transcription factors co-expressed with CilXTH. (B) The co-expression network of CilXTH genes in the shell of SX (thick-shelled pecan) during the middle and late developmental stages. The blue squares represent genes co-expressed with CilXTH, while the green squares indicate transcription factors co-expressed with CilXTH. (C) The cis-regulatory element analysis of the CilXTH gene promoters. Each horizontal line represents the promoter region of a CilXTH gene, with differently colored squares indicating the presence of various cis-acting elements associated with plant growth, hormone responses, and environmental stress regulation. The legend on the right identifies the functional categories of these elements.
Figure 7. Co-expression network and cis-regulatory element analysis of CilXTH genes. (A) The co-expression network of the CilXTH genes in the shell of KD (thin-shelled pecan) during the early developmental stage. The blue squares represent genes co-expressed with CilXTH, while the pink squares indicate transcription factors co-expressed with CilXTH. (B) The co-expression network of CilXTH genes in the shell of SX (thick-shelled pecan) during the middle and late developmental stages. The blue squares represent genes co-expressed with CilXTH, while the green squares indicate transcription factors co-expressed with CilXTH. (C) The cis-regulatory element analysis of the CilXTH gene promoters. Each horizontal line represents the promoter region of a CilXTH gene, with differently colored squares indicating the presence of various cis-acting elements associated with plant growth, hormone responses, and environmental stress regulation. The legend on the right identifies the functional categories of these elements.
Horticulturae 11 00609 g007
Table 1. Physiological characteristics of XTH gene family in Carya illinoensis.
Table 1. Physiological characteristics of XTH gene family in Carya illinoensis.
Gene NameGene IDCDS (bp)Protein (aa)ExonpIBasic Proteins (%)MW (Da)Sub-Loc
CilXTH1Cil_01G_00632V2119739869.2613.5745,652.15CW/C
CilXTH2Cil_01G_00640V229798110.0219.3911,506.32CW
CilXTH3Cil_02G_00706V2105635145.6812.5440,021.45CW
CilXTH4Cil_03G_00077V293931258.9214.1036,247.32CW/C
CilXTH5Cil_03G_00411V282827549.0713.4531,919.27CW
CilXTH6Cil_03G_01067V2119139566.1814.6545,167.80 CW
CilXTH7Cil_05G_00828V2101133648.9813.9938,764.06CW
CilXTH8Cil_06G_01204V288229359.3615.7033,792.43CW
CilXTH9Cil_07G_00376V287329038.80 11.3832,412.66CW/C
CilXTH10Cil_07G_01585V291230358.6214.5234,514.17CW
CilXTH11Cil_08G_01020V283427746.2910.4730,791.31CW
CilXTH12Cil_09G_02459V285828539.3612.2832,268.65CW/C
CilXTH13Cil_10G_01144V279526445.1611.7430,541.09CW
CilXTH14Cil_10G_01312V298132646.0012.2737,277.34CW
CilXTH15Cil_11G_00212V2135345089.7815.1150,254.36CW
CilXTH16Cil_11G_00890V285529348.1612.9734,115.55CW/C
CilXTH17Cil_11G_01428V2117639168.9014.8344,026.61CW
CilXTH18Cil_11G_01492V2100533446.8614.6738,381.49CW
CilXTH19Cil_12G_00855V293030935.098.4134,160.98CW/C
CilXTH20Cil_12G_00856V236612136.069.9213,971.90 CW/C
CilXTH21Cil_12G_00857V299333036.079.3936,838.15CW/C
CilXTH22Cil_13G_01077V2104134649.2514.7439,987.48CW
CilXTH23Cil_13G_01231V278626137.7410.3428,932.94CW/C
CilXTH24Cil_14G_00900V287929244.6710.2734,105.96CW
CilXTH25Cil_14G_01287V287629137.1112.7133,539.68CW/C
CilXTH26Cil_14G_01719V285228349.2414.1332,416.48CW
CilXTH27Cil_15G_00375V285828546.7012.2832,546.05CW
CilXTH28Cil_15G_01868V280426744.667.8730,032.17CW/C
CilXTH29Cil_15G_01869V286428748.769.4131,723.85CW
CilXTH30Cil_15G_01870V21218345108.9012.8444,668.86CW
CilXTH31Cil_15G_01871V268122644.608.5325,207.82CW
CilXTH32Cil_15G_01872V288229335.019.4332,463.03CW/C
CilXTH33Cil_15G_01873V289329735.948.9734,074.27CW/C
CilXTH34Cil_15G_01963V287329035.9310.5632,377.34CW/C
CilXTH35Cil_16G_00439V2112837559.3810.3342,085.19CW/CP
CilXTH36Cil_16G_00440V21281426105.417.0848,044.13CW/CP
CilXTH37Cil_16G_00824V290130045.7113.8733,734.39CW
CilXTH38Cil_73S_00004V2177959287.0314.1966,148.33CW
CW: Cell wall; C: Cytoplasm; CP: Chloroplast.
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

Wen, M.; Zhou, Z.; Sun, J.; Meng, F.; Xi, X.; Liu, A.; Yu, A. A Genome-Wide Characterization of the Xyloglucan Endotransglucosylase/Hydrolase Family Genes and Their Functions in the Shell Formation of Pecan. Horticulturae 2025, 11, 609. https://doi.org/10.3390/horticulturae11060609

AMA Style

Wen M, Zhou Z, Sun J, Meng F, Xi X, Liu A, Yu A. A Genome-Wide Characterization of the Xyloglucan Endotransglucosylase/Hydrolase Family Genes and Their Functions in the Shell Formation of Pecan. Horticulturae. 2025; 11(6):609. https://doi.org/10.3390/horticulturae11060609

Chicago/Turabian Style

Wen, Mengyun, Zekun Zhou, Jing Sun, Fanqing Meng, Xueliang Xi, Aizhong Liu, and Anmin Yu. 2025. "A Genome-Wide Characterization of the Xyloglucan Endotransglucosylase/Hydrolase Family Genes and Their Functions in the Shell Formation of Pecan" Horticulturae 11, no. 6: 609. https://doi.org/10.3390/horticulturae11060609

APA Style

Wen, M., Zhou, Z., Sun, J., Meng, F., Xi, X., Liu, A., & Yu, A. (2025). A Genome-Wide Characterization of the Xyloglucan Endotransglucosylase/Hydrolase Family Genes and Their Functions in the Shell Formation of Pecan. Horticulturae, 11(6), 609. https://doi.org/10.3390/horticulturae11060609

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