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. 2023 Jan 6;191(3):1734–1750. doi: 10.1093/plphys/kiad002

The transcription factor PbrbZIP52 positively affects pear pollen tube longevity by promoting callose synthesis

Zhongheng Xia 1,#, Binxu Wen 2,#, Jing Shao 3,#, Tianci Zhang 4,#, Mengmeng Hu 5, Lin Lin 6, Yiping Zheng 7, Zhixin Shi 8, Xinlin Dong 9, Juanjuan Song 10, Yuanshan Li 11, Yongjie Wu 12, Yafang Yuan 13, Juyou Wu 14,✉,3,4,5, Qingxi Chen 15,✉, Jianqing Chen 16,✉
PMCID: PMC10022607  PMID: 36617219

Abstract

In pear (Pyrus bretschneideri), pollen tube growth is critical for the double fertilization associated with seed setting, which in turn affects fruit yield. The normal deposition of callose mediates the polar growth of pollen tubes. However, the mechanism regulating callose synthesis in pollen tubes remains relatively uncharacterized. In this study, we revealed that the typical pear pollen tube lifecycle has a semi-growth duration (GD50) of 16.16 h under in vitro culture conditions. Moreover, callose plugs were deposited throughout the pollen tube lifecycle. The formation of callose plugs was inhibited by 2-deoxy-D-glucose, which also accelerated the senescence of pear pollen tubes. Additionally, PbrCalS1B.1, which encodes a plasma membrane-localized callose synthase, was expressed specifically in pollen tubes and restored the fertility of the Arabidopsis (Arabidopsis thaliana) cals5 mutant, in which callose synthesis is inhibited. However, this restoration of fertility was impaired by the transient silencing of PbrCalS1B.1, which restricts callose plug formation and shortens the pear pollen tube lifecycle. More specifically, PbrbZIP52 regulated PbrCalS1B.1 transcription by binding to promoter A-box elements to maintain the periodic formation of callose plugs and normal pollen tube growth, ultimately leading to double fertilization. This study confirmed that PbrbZIP52 positively affects pear pollen tube longevity by promoting callose synthesis. This finding may be useful for breeding high-yielding pear cultivars and stabilizing fruit setting in commercial orchards.

Introduction

The success of sexual reproduction resulting in high fruit yields depends on healthy pollen tube growth. After adhering to stigma papilla cells and hydrating, pollen grains germinate and produce a tube that grows over a relatively long distance to reach the ovule for the subsequent double fertilization (Higashiyama and Takeuchi, 2015). During this process, vesicles containing cell wall matter are transported by the actin cytoskeleton to the apical region via highly dynamic cytoplasmic flow, after which they fuse with the plasma membrane (i.e. exocytosis) to facilitate the continuous synthesis of the cell wall, ultimately leading to the morphological changes that enable the rapid elongation of pollen tubes (Guan et al., 2013; Hepler et al., 2013). Callose is a structural component of the cell wall that is widely distributed in reproductive tissues.

Consistent callose metabolism, during which callose is deposited and degraded, is an indispensable dynamic process (Dong et al., 2005; Zhang et al., 2007). Callose is initially deposited between the plasma membrane and the primary wall of microsporocytes during the first meiotic division, after which it gradually extends toward the center of the cell and then forms the callose wall in the equatorial plate (Zhang et al., 2007). The callose wall separates the microsporocyte into a tetrad, thereby preventing the fusion and premature development of microspores; it is subsequently hydrolyzed by β-1,3-glucanases, leading to the release of the microspores (Lu et al., 2014). A vegetative cell and a germ cell, which are produced by the asymmetric division of uninucleate microspores during the first mitosis, are separated by a new callose wall. To facilitate the insertion of the germ cell into the cytoplasm of the vegetative cell to form a mature pollen grain, the callose wall is eventually degraded (Oh et al., 2021). The mature pollen grain germinates and produces a tube in which callose is deposited substantially more in the inner layer of the lateral wall than at the tip. This particular spatial distribution provides the lateral walls with sufficient mechanical strength to resist the effects of inflationary pressure, but it also increases the ductility at the tip to maintain polar pollen tube growth (Parre and Geitmann, 2005; Chebli et al., 2012). Intriguingly, the periodic formation of callose plugs is a unique phenomenon in pollen tubes. This results in sperm cells surrounded by the pollen tube cytoplasm at the tip, which maintains a relatively stable internal environment conducive to the rapid elongation of the pollen tube that eventually releases sperm cells in time for the double fertilization (Guan et al., 2013). However, the precise molecular mechanisms mediating the effects of the callose plug on pollen tube growth and fertility need to be further clarified.

Callose synthase (CalS), which is also known as glucan synthase-like (GSL), is a plasma membrane-localized enzyme that uses UDP-glucose as a substrate to synthesize callose. Twelve CalS-encoding genes (AtCalS1 to AtCalS12) have been identified in the Arabidopsis (Arabidopsis thaliana) genome (Hong et al., 2001), several of which are actively expressed during the pollen development stage. For example, callose wall formation in microspores is mediated by AtCalS11 and AtCalS12 in the tetrad stage (Enns et al., 2005). The silencing of AtCalS9 and AtCalS10 results in abnormal microspore division and the eventual death of male gametophytes during the first mitosis (Töller et al., 2008). There is increasing evidence that only AtCalS5 regulates callose synthesis from the pollen development stage to the pollen tube elongation stage (Dong et al., 2005; Nishikawa et al., 2005; Abercrombie et al., 2011; Mizuta and Higashiyama, 2014). Another study indicated that AtCalS5 controls the development of the peripheral callose wall to prevent the early degeneration of microspores (Dong et al., 2005; Nishikawa et al., 2005). Compared with pollen grains, callose represents a larger proportion of the structural components of pollen tubes, which is critical for pollen tube growth. Specifically, callose forms the inner layer of the pollen tube cell wall, but it is also present as a distinct structure (i.e. callose plug). However, the specific CalS gene encoding the regulator of callose synthesis in pollen tubes remains relatively uncharacterized.

Callose synthesis mediated by CalS is regulated at the transcriptional level. In A. thaliana, auxin response factor 17 controls the transcription of AtCalS5 by binding to the auxin-responsive element, leading to the sustained formation of the pollen wall during the binuclear pollen stage (Yang et al., 2013). A recent study revealed that the corresponding transcriptional regulatory pathway is more complex in cotton (Gossypium hirsutum), in which a pollen-specific protein (i.e. PSP231) inhibits the binding of RNA-binding protein-like 1 to GhWRKY15 mRNA, which is a transcriptional repressor, thereby promoting the transcription of CalS4 and CalS8 and activating callose biosynthesis to facilitate pollen maturation (Li et al., 2020b). In terms of the transcriptional modification pathway, cyclin-dependent kinase G1 modulates the pollen wall pattern formation by regulating the splicing of AtCalS5 pre-mRNA in A. thaliana (Huang et al., 2013). Despite these earlier findings, the delicate mechanism regulating CalS gene expression to promote callose deposition, especially the formation of callose plug, in pollen tubes lifecycle has yet to be comprehensively explored.

This study was conducted to answer the following three key questions: (1) How does the deposition of callose plugs affect the fertility-related function of pear (Pyrus bretschneideri) pollen tubes? (2) Which CalS gene regulates callose plug synthesis in pear pollen tubes? (3) Which mechanisms regulate callose plug synthesis in pear pollen tubes? The data presented herein revealed a molecular regulatory mechanism that PbrbZIP52 maintains pollen tube longevity in pear by regulating PbrCalS1B.1-mediated callose plug periodically formation. These findings bridge the gap in the delicate regulation mechanisms of callose plug formation in pollen tube. It provides distinctive insights into pear pollen tube growth and the seed setting rate, which may be relevant for breeding high-yielding pear cultivars.

Results

Callose plug deposition occurs during the lifecycle of pear pollen tubes under in vitro culture conditions

The pollen tube lifecycle starts after the pollen grain germinates. The pollen tube then undergoes highly polarized and rapid tip growth to enable it to extend over relatively long distances to reach the ovule prior to fertilization. In this study, the growth–senescence curve for “Dangshansuli” pear pollen tubes under in vitro culture conditions had an S-shape. More specifically, we detected rapid growth starting at 3 hours post-culture (HPC), after which the growth rate decreased at 15 HPC and then stabilized after 18 HPC (Figure 1, A and C). Because senescent cells were precisely eliminated via programmed cell death (PCD), the terminal-deoxynucleotidyl transferase-mediated nick-end labeling (TUNEL) staining technique was used to assess DNA-strand breaks, which are indicative of the occurrence of PCD in pear pollen tubes (Chen et al., 2018) (Figure 1B). The PCD rate increased from 12 to 21 HPC, implying that the pollen tubes in the late growth stage were gradually senescing as PCD was initiated. Most of the cells were dead at 24 HPC (Figure 1C). These observations suggest that under in vitro culture conditions, the completion of the pear pollen tube lifecycle is mediated by PCD.

Figure 1.

Figure 1

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Callose deposition during the pear pollen tube lifecycle. A, Typical images of pear pollen tubes under in vitro culture conditions at 3 HPC (left) and 15 HPC (right). Bar = 100 μm. B, TUNEL-based detection of PCD in pollen tubes. Typical images of TUNEL signals in pollen tubes at 3 HPC (left) and 15 HPC (right) are presented. The DAPI staining results indicate that the TUNEL-positive signals are localized in the nuclear DNA. “Positive” reveals that a PCD event has occurred in the pollen tube, while “negative” means not. Bar = 10 μm. C, Time-course analysis of the TUNEL-reactive nuclei in pear pollen tubes. The data represent the mean value and standard error of three independent biological replicates, each comprising at least 100 pollen tubes. D, Typical image of a pear pollen tube with multiple callose plugs at 15 HPC under in vitro culture conditions. The arrows indicate the callose plugs in the pollen tube. Bar = 100 μm. E, Number of callose plugs during the pollen tube growth stage. The data represent the mean value and standard error of three independent biological replicates, with an average of at least 500 pollen tubes per replicate. F, Relative callose content during the pear pollen tube lifecycle. Data are presented as the mean value and standard error of three independent biological replicates.

To explore the potential factors affecting the pear pollen tube lifecycle, we investigated the formation of callose plugs in pollen tubes. These plugs, which were first detected at 6 HPC, were widely distributed in pear pollen tubes and continued to grow until 24 HPC. Most of the examined pollen tubes contained more than four callose plugs during their lifecycle (Figure 1, D and E). Furthermore, the change in the relative callose content was consistent with the pollen tube elongation rate (R = 0.991), which decreased substantially during the acceleration of PCD in pear pollen tubes (Figure 1, C and F). These findings imply that the deposition of callose plugs is accompanied by pollen tube elongation and is an integral part of the pollen tube lifecycle under in vitro culture conditions.

Inhibited callose plug formation leads to the premature senescence of pear pollen tubes

To verify the importance of callose plugs, we used 2-deoxy-D-glucose (2DDG) (Riehl and Jaffe, 1984) to disrupt callose biosynthesis in pollen tubes. The 2DDG treatment substantially decreased the number of callose plugs and the relative callose content in pear pollen tubes. The 0.5 mM treatment was sufficient for the optimal inhibitory effects of 2DDG (Figure 2, A, D, and E). Compared with the mock treatment, the 0.5 mM 2DDG treatment substantially inhibited pear pollen tube growth (Figure 2, B and F) and shortened the lifecycle, with the semi-growth duration (GD50) for the PCD decreasing significantly from 16.16 ± 1.22 to 11.39 ± 1.48 h (P < 0.05, n = 3) (Figure 2G). Moreover, the actual density of callose plug from pollen tube whose callose synthesis was inhibited by 0.5 mM 2DDG was significantly reduced by 44.94% in an equal style area which enriches pollen tubes, compared to the Mock (Supplemental Figure 1). Ultimately, pollen tubes with defective callose synthesis by 0.5 mM 2DDG grew to only 53.6% of the distance to the stigma position, resulting in a failure to complete fertilization. However, normal fertilizations were observed for the mock-treated dried pollen grains and pollen grains cultured in medium (Figure 2, H and I). These results indicate that the continuous formation of callose plugs is essential for a normal pear pollen tube lifecycle and fertility.

Figure 2.

Figure 2

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2DDG inhibits callose synthesis to shorten the pear pollen tube lifecycle. A, Typical images of the mock-treated pollen tubes (top) and the pollen tubes treated with 0.5 mM 2DDG (bottom) with callose plugs at 15 HPC under in vitro culture conditions. The arrow indicates the callose plug in the pollen tube. Bar = 100 μm. B, C, Images of the typical growth (B) and TUNEL signals (C) in the mock-treated pollen tubes (left) and the pollen tubes treated with 0.5 mM 2DDG (right) at 15 HPC. Bars = 50 and 10 μm, respectively. D–G, Pollen incubated in liquid medium containing different concentrations of 2DDG at different time-points. The number of callose plugs (D), relative callose content (E), length (F), and semi-growth duration (GD50) (G) for the pear pollen tubes were investigated. The chart in the top right corner of (D) is a magnified figure of statistical number of pollen tube with “4” and “≧5” callose plugs. At least 500 measurements were performed for the callose plug experiments. The data for the pollen tube length (F) and GD50 (G) were calculated on the basis of the length and PCD at 1, 3, 6, 9, 12, 15, 18, 21, and 24 HPC, with at least 90 measurements per time-point. H, I, Dried (UT), mock-treated, and 2DDG-treated “Dangshansuli” pollen grains were germinated on the stigma of “Cuiguan” flowers. Typical images (H) and pollen tube lengths relative to the pistil length (I) are presented. The arrow indicates the longest pollen tube in the pistil. Bar = 100 mm. The data are presented as the mean value and standard error of three replicates. Different letters indicate significant differences as determined by ANOVA followed by Tukey’s multiple comparison test (P < 0.05).

PbrCalS1B.1 expresses in the pear pollen tube

To investigate the specific CalS genes involved in the regulation of callose synthesis in pear pollen tubes, we systematically identified 100 CalS genes in nine Rosaceae species, including pear (Figure 3A). These genes along with A. thaliana AtCalS genes were used to construct a phylogenetic tree according to the maximum likelihood method. The analysis of phylogenetic relationships revealed that the CalS genes from Rosaceae species were divided into nine clusters in three groups (Figure 3B). This classification was supported by bootstrap values exceeding 74% for the branches as well as the uniformity of the conserved domains and gene structures (Figure 3; Supplemental Figure 2). Clearly, the CalS genes in Rosaceae species were homologous to the A. thaliana genes, with no additional clusters, possibly reflecting the conserved functions among the homologous CalS genes.

Figure 3.

Figure 3

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Identification of the CalS gene family members in Rosaceae. A, Distribution and the members of each CalS cluster in Rosaceae. The pentagrams represent genome-wide replication events. B, Phylogenetic relationships among CalS genes from A. thaliana and Rosaceae species. The CalS protein sequences were compared by a multiple sequence alignment using the MAFFT software. A maximum likelihood phylogenetic tree was constructed using IQ-TREE2, with 1,000 replications. Scale bar, 20 substitutions/100 bases. The sequence alignment of CalS protein used for the phylogenetic analysis is listed in Supplemental File 1.

To explore the expression of CalS genes in pear pollen, we performed a tissue-specific expression analysis using transcriptomic data for the pollen, pistil, bud, leaf, and fruit. Of the examined CalS genes, only PbrCalS1B.1 was specifically expressed in the pollen tube (Figure 4A). We subsequently generated transgenic A. thaliana plants expressing proPbrCalS1B.1-β-glucuronidase (GUS) fusion proteins. Strong GUS activity was detected in the anthers, pollen grains, and pollen tubes (Figure 4B). A similar pollen-specific expression of Group 1B CalS genes was observed in Malus domestica, Prunus mume, and Prunus avium (Supplemental Figure 3). We further checked the correctness of above inference of PbrCalS1B.1 expression in multiple tissues of pear. It confirmed that PbrCalS1B.1 was merely found to be expressed at high level in flower (Figure 4C). In detail, PbrCalS1B.1 expression was only detected in the anther, mature pollen, and pollen tube, while it was not detected in other floral organs, such as ovary, calyx, petal, style, and filament (Figure 4D). Considered together, these findings suggest that Group 1B CalS members may function in the pollen of Rosaceae species. We next constructed recombinant plasmids for the expression of PbrCalS1B.1 with an N-terminal green fluorescent protein (GFP) for the transformation of A. thaliana mesophyll protoplasts. A strong fluorescent signal was detected in the plasma membrane of protoplasts (Figure 4E). Accordingly, PbrCalS1B.1 appears to be a pear pollen-specific protein located in the plasma membrane.

Figure 4.

Figure 4

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Plasma membrane-localized PbrCalS1B.1 expression in pear pollen. A, Transcriptome data for CalS expression in the pollen, pistil, ovary, sepal, bud, leaf, and fruit. The PbrCalS1B.1 gene was specifically expressed in pollen. RPKM, reads per kilobase of exon model per million mapped reads. White-to-dark blue indicates a gradual increase in expression. B, GUS reporter staining of pPbrCalS1B::GUS transgenic Arabidopsis plants revealed the tissue-specific expression of PbrCalS1B.1. The GUS signal was detected in the anthers, pollen grains, and pollen tubes. Bars = 2 mm, 0.5 mm, 10 μm, and 5 μm for the images of the seedlings, flowers, pollen grains, and pollen tubes, respectively. C, The relative expression level of PbrCalS1B.1 in different tissues of “Dangshansuli” as determined by RT-qPCR. D, The expression pattern of PbrCalS1B.1 in spatial and temporal from individual floral organs during floral development, including S1–S6 stages, mature pollen, and 6 HPC pollen tube of “Dangshansuli”. Bar = 5 mm. The images were digitally extracted for comparison and the scale bar provided is accurate for each image. E, Subcellular localization of PbrCalS1B.1 in A. thaliana protoplasts. Protoplasts containing GFP alone or PbrCalS1B.1-GFP were examined using a confocal microscope at 16 HPC. The experiment was repeated three times, with similar results. Bar = 10 mm. Data in (C) and (D) presented as the mean value and standard error of five replicates.

PbrCalS1B.1 regulates callose plug deposition and affects pear pollen fertility

In A. thaliana, AtCalS5 affects pollen fertility and pollen tube growth by controlling callose synthesis (Mizuta and Higashiyama, 2014). In the current study, we revealed the decreased fertility of the atcals5 mutant (Salk_009234). There were no observable differences between the wild-type control and the atcals5-1 mutant during the vegetative stage (Figure 5A). However, compared with the wild-type control, the pod length decreased significantly by 36.5% (P < 0.001, n = 400; Figure 5, B and F), the number of seeds decreased significantly from 64.92 ± 6.47 to 37.61 ± 8.62 (P < 0.001, n = 39; Figure 5, C and G), and the seed loss rate increased significantly (up to 15.93%; P < 0.001, n = 39; Figure 5H) in the atcals5-1 mutant line. Additionally, the surface of the mutant pollen grains had an irregular reticulate pattern. Moreover, the cell wall deficiency rate was up to 62.9% ± 10.1%, which was significantly higher than the corresponding rate in the wild-type control (P < 0.001, n = 14; Figure 5, D and I). More importantly, the callose plug loss rate was significantly higher for the atcals5-1 mutant pollen tubes (49.3%) than for the wild-type pollen tubes at 6 HPC (P < 0.001, n = 15; Figure 5, E and J). The PbrCalS1B.1 gene is a homolog of AtCalS5 and shares the same conserved functional domain (Figure 3; Supplemental Figure 4). Therefore, we restored the fertility of the atcals5-1 mutant by constructing atcals5-1/PbrCalS1B.1-overexpressing transgenic lines. A total of 12 atcals5-1/PbrCalS1B.1-overexpressing transgenic lines were obtained in the T3 generation, of which lines #3 and #4 had the highest expression levels and were selected for the subsequent analysis (Supplemental Figure 5). As expected, in both restorer lines, fertility was recovered to almost wild-type levels (Figure 5). These results indicate that PbrCalS1B.1 regulates callose synthesis to mediate pollen tube development and pollen fertility.

Figure 5.

Figure 5

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Restoration of the fertility of the A. thaliana atcals5-1 mutant by PbrCalS1B.1. A–E, Images of the wild-type, atcals5-1, atcals5-1/PbrCalS1B.1 #3, and atcals5-1/PbrCalS1B.1 #4 (left to right) A. thaliana samples. A, Three-week-old A. thaliana plants during nutritional growth. Bar = 30 mm. B, Inflorescence stems. Bar = 20 mm. C, Full-grown siliques, with red spots representing the aborted seeds. Bar = 1 mm. D, Pollen grain structure. Bar = 5 μm. E, Callose plugs in pollen tubes stained with aniline blue. The yellow and white arrows indicate the positions of the callus plug and the pollen tube tip, respectively. Bar = 10 μm. The images were digitally extracted for comparison and the scale bar provided is accurate for each image in (A–E). F–J, Phenotypic analysis of the wild-type, atcals5-1, atcals5-1/PbrCalS1B.1 #3, and atcals5-1/PbrCalS1B.1 #4 A. thaliana samples. F, Pod length. G, Number of seeds. H, Seed loss rate. I, Cell wall deficiency rate in pollen grains. At least 95 pollen grains were examined per biological replicate. J, Callose plug loss rate in pollen tubes at 6 HPC. For each treatment, at least 95 pollen grains or pollen tubes were examined per biological replicate in (I) and (J). In each box–whisker plot (F–G), interquartile range including middle line (50% of data), bottom line (25% of data), and top line (75% of data), the upper whisker represents extends to maximum data point within 1.5 box heights from the top of box, the lower whisker represents extends to minimum data point within 1.5 box heights from the bottom of box, outlier represents observation beyond the upper or lower whisker. Different letters indicate significant differences above each box plot as determined by ANOVA followed by Tukey’s multiple comparison test (P < 0.05).

An antisense oligodeoxynucleotide (as-ODN) method was employed to knock down the expression of PbrCalS1B.1 in pear pollen tubes to confirm callose plugs have critical functions in pollen tubes. The final concentration of 50 μ M as/s-ODN was employed without ODN's toxins effect and optimal repression in growth was ensured (Supplemental Figure 6). Compared with the control, the PbrCalS1B.1 expression level decreased by 68.8% after the as-ODN-PbrCalS1B.1 treatment, whereas the sense oligodeoxynucleotide (s-ODN) treatment did not inhibit the expression of the target gene (Supplemental Figure 7, A and B). We observed that the as-ODN-PbrCalS1B.1 treatment substantially decreased the number of callose plugs and the relative callose content compared with the effects of the mock treatment (Figure 6, A and B). It also arrested pollen tube growth (Figure 6C) and accelerated the onset of PCD according to the TUNEL staining of pollen tubes, which indicated that the GD50 for the PCD decreased significantly from 16.18 ± 1.43 to 11.4 ± 1.83 h (P < 0.05, n = 3; Figure 6D; Supplemental Figure 8A). A similar result was obtained using fluorescein diacetate, which is another pollen viability marker used for detecting death (Supplemental Figure 8B). Notably, the as-ODN-PbrCalS1B.1 pollen had senescent phenotypes similar to the pollen treated with 2DDG (Figure 2). However, these inhibitory effects on the callose-mediated fertility of pear pollen were not detected after the s-ODN treatment. Hence, PbrCalS1B.1 can regulate callose plug deposition to modulate the pear pollen tube lifecycle and pollen fertility.

Figure 6.

Figure 6

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Silencing of PbrCalS1B.1 expression inhibits callose synthesis in pear pollen tubes and promotes senescence. Pollen was incubated in liquid medium for the mock, s-ODN-PbrCalS1B.1, and as-ODN-PbrCalS1B.1 treatments and then analyzed at different time-points. A, Typical images of the mock-treated pollen tubes and the pollen tubes treated by as-ODN-PbrCalS1B.1 with callose plugs and the distribution of the number of callose plugs in pear pollen tubes at 12 HPC in vitro culture conditions. The arrows indicate the callose plugs in the pollen tube. Bar = 100 μm. Each experiment included at least 500 measurements. B, Relative callose contents in growing pear pollen tubes. Data are presented as the mean value of three replicates. C, D, Pear pollen tube length (C) and PCD (D). Experiments were repeated three times, with each experiment including at least 95 measurements. *P < 0.05, **P < 0.01, and ***P < 0.001 by ANOVA followed by Tukey’s multiple comparison test. Error bars indicate the standard error.

PbrbZIP52 regulates PbrCalS1B.1 transcription to maintain the pear pollen tube lifecycle

To clarify the mechanism regulating PbrCalS1B.1 expression in pollen tubes, PbrCalS1B.1 expression in pollen tubes was activated substantially after 6 HPC, reflecting the potential transcriptional activation of PbrCalS1B.1 in pear pollen tubes. The expression of a basic leucine zipper (bZIP) transcription factor gene (PbrbZIP52) whose divided into cluster 6B of bZIP gene family with AtbZIP52 (Supplemental Figure 9) was strongly correlated with the PbrCalS1B.1 expression pattern (R = 0.975) during the pear pollen tube lifecycle (Figure 7A). It is worth concerning that the tissue expression pattern of PbrbZIP52 (Supplemental Figure 10) also indicates high similarity to that of PbrCalS1B.1 (Figure 4), implying their potential functional relevance. We then inhibited PbrbZIP52 expression using as-ODN (Supplemental Figure 7, C and D), which significantly decreased the PbrCalS1B.1 expression level compared with the mock treatment (Figure 7B). Notably, the pollen tube lifecycle-related phenotypes following the as-ODN-PbrbZIP52 treatment were similar to those after the as-ODN-PbrCalS1B.1 treatment (Figures 6 and 7). For example, compared with the effects of the mock treatment, the number of callose plugs (Figure 7C) and the relative callose content (Figure 7D) decreased substantially, pollen tube growth was restricted, and the GD50 for the PCD decreased significantly from 15.0 ± 1.61 to 9.91 ± 1.09 h (P < 0.05, n = 3; Figure 7, E and F) in response to the as-ODN-PbrCalS1B.1 treatment. In contrast, these fertility-related defects were undetectable following the s-ODN-PbrbZIP52 treatment (Figure 7). These results imply that PbrbZIP52 may mediate the periodic formation of callose plugs by regulating PbrCalS1B.1 transcription to maintain normal pear pollen tube growth.

Figure 7.

Figure 7

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PbrbZIP52 regulates PbrCalS1B.1 transcription to control callose synthesis and the pear pollen tube lifecycle. A, RT-qPCR analysis of the PbrCalS1B.1 and PbrbZIP52 expression patterns. The R-value reflects the correlation between expression models. Twenty milligrams of pollen was incubated in liquid medium for the mock, s-ODN-PbrbZIP52, and as-ODN-PbrbZIP52 treatments and then analyzed at different time-points. B, RT-qPCR analysis of the PbrCalS1B.1 expression pattern in developing pollen tubes. Data are presented as the mean value and standard error of five replicates in (A) and (B). C–F, The typical images and number of callose plugs (C), relative callose content (D), length (E), and semi-growth duration (GD50) (F) of pear pollen tubes are presented. The arrows indicate the callose plugs in the pollen tube. Bar = 100 μm. At least 500 measurements were performed for the callose plug experiments. The data for the pollen tube length (E) and GD50 (F) were calculated on the basis of the length and PCD at 1, 3, 6, 9, 12, 15, 18, 21, and 24 HPC, with at least 95 measurements per time-point. Data are presented as the mean value and standard error of three replicates. Different letters indicate significant differences and *P < 0.05, **P < 0.01, and ***P < 0.001 by ANOVA followed by Tukey’s multiple comparison test.

The bZIP transcription factors regulate diverse biological processes by binding to core cis-acting elements containing ACGT, including ABRE [(C/T/G)ACGT(G/T)], A-box (TACGTA), G-box (CACGTG), and C-box (GACGTC), in the promoter region of target genes (Izawa et al., 1994; Fujita et al., 2005). Two A-box elements located in the −219 to −291 bp promoter region of PbrCalS1B.1 were detected, but there were no other ACGT-containing elements (Figure 8A). We subsequently determined whether PbrbZIP52 can bind to the PbrCalS1B.1 promoter on the basis of a yeast one-hybrid (Y1H) analysis. The observed auto-activation activities of the PbrCalS1B.1 promoter containing two A-box elements (pCalS1B.1WT) were inhibited by 200 ng mL−1 aureobasidin A (AbA200) (Figure 8B). After the Y1H-Gold yeast strain cells were co-transformed with pGADT7-PbrbZIP52 and pAbAi-PbrCalS1B.1WT, the binding of PbrbZIP52 to the pCalS1B.1WT region was revealed by the abundant growth on the SD/−Leu/−Ura/+AbA200 medium (Figure 8B). In the electrophoretic mobility shift assay (EMSA), the detected protein–DNA complex comprised the recombinant GST-PbrbZIP52 fusion protein bound directly to pCalS1B.1WT. The strength of this binding decreased gradually in the competition experiments involving five- and 25-fold molar excesses of unlabeled probes (Figure 8C). This specific binding was further confirmed in a transient transcriptional activity assay. When the effector vector pCaMV35S-PbrbZIP52 was co-expressed with the reporter vector pCalS1B.1WT-LUC (i.e. luciferase), with pCaMV35S-GUS as an internal reference control, the LUC:GUS ratio increased significantly (i.e. 2.7-fold higher than the control; P < 0.05, n = 5; Figure 8D). Thus, PbrbZIP52 can interact with the PbrCalS1B.1 promoter in the pear pollen tube.

Figure 8.

Figure 8

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The A-box motif of the PbrCalS1B.1 promoter is crucial for the interaction with PbrbZIP52. A, Schematic diagram of the PbrCalS1B.1 promoter. The A-box element is indicated by a line. m1, m2, and m1&m2 represent the mutated A-box sequences. B, The interaction between PbrbZIP52 and the PbrCalS1B.1 promoter region (−2,000 bp) was analyzed in a yeast one-hybrid assay. The auto-activation of the PbrCalS1B.1 promoter and the inhibitory effects of AbA were assessed on −Ura selection medium (top). The co-transformed strains were incubated on −Ura/−Leu medium supplemented with 200 ng mL−1 AbA and X-gal (below). C, EMSA-based analysis of the binding between PbrbZIP52 and the PbrCalS1B.1 promoter. The biotin-labeled probes and unlabeled competitors were incubated with the GST-PbrbZIP52 fusion protein. The presence or absence of specific probes is indicated by “+” or “−”. The arrowheads indicate the shifted probe (top) and the free probe (bottom). D, Binding of PbrbZIP52 to the PbrCalS1B.1 promoter determined on the basis of transcriptional activity assays. A schematic diagram of the various vectors (top) and the results of the transcriptional activation analysis (bottom) are presented. The LUC:GUS ratio for proPbrCalS1B.1WT-LUC/pCaMV35S-GUS was used as the calibrator (set as 1). Different letters indicate significant differences as determined by ANOVA followed by Tukey’s multiple comparison test (P < 0.05, n = 5). Error bars indicate the standard error.

To further verify the interaction, the A-box elements in the PbrCalS1B.1 promoter were mutated (ACGT converted to GATC). More specifically, we mutated the proximal A-box-binding site (m1), the distal A-box-binding site (m2), and both A-box-binding sites (m1 & m2) (Figure 8A). The Y1H, EMSA, and transient transcriptional activity assay results confirmed that the binding of PbrbZIP52 to the PbrCalS1B.1 promoter region was substantially weakened by the single mutations, while the interaction was almost completely inhibited by the double mutation (Figure 8). These observations indicate that the binding of PbrbZIP52 to the PbrCalS1B.1 promoter is dependent on the A-box elements.

Discussion

Pollen tubes transmit sperm cells to the ovule via the style for the double fertilization in angiosperms (Higashiyama and Takeuchi, 2015). During this process, the formation of callose plugs causes the cytosol and sperm cells to concentrate at the pollen tube tip to continuously form the matter required for rapid growth, thereby ensuring sperm cells are released appropriately (Guan et al., 2013). The precise regulation of callose synthesis is essential for successful fertilization. Regrettably, the elaborate mechanism regulating callose synthesis in pollen tubes has not been fully characterized. The results of this study indicate that callose plugs accumulate throughout the pear pollen tube lifecycle under in vitro culture conditions. The inhibition of callose synthesis substantially impeded pear pollen tube elongation and led to premature senescence. Additionally, PbrCalS1B.1, which encodes a plasma membrane protein, was revealed to be the only CalS gene specifically expressed in pear pollen tubes. The transient silencing of PbrCalS1B.1 can decrease the relative callose content and the number of callose plugs, thereby shortening the pear pollen tube lifecycle. Furthermore, the PbrbZIP52 transcription factor was observed to bind to the A-box element in the PbrCalS1B.1 promoter to influence CalS-mediated pollen fertility. Thus, we propose that PbrbZIP52 regulates PbrCalS1B.1 transcription to modulate callose synthesis and maintain healthy pear pollen tube growth (Figure 9).

Figure 9.

Figure 9

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Model of the bZIP52–CalS1B.1 signaling that regulates the synthesis of callose plugs necessary for maintaining the pear pollen tube lifecycle. In growing pollen tubes, PbrCalS1B.1 expression is regulated by the binding of PbrbZIP52 to the A-box, ultimately resulting in the production of PbrCalS1B.1 localized in the cytoplasmic membrane, wherein it mediates the synthesis of callose in pollen tubes. The callose forms plugs or the callose wall to maintain the normal pollen tube lifecycle.

Callose plugs are crucial for the healthy pollen tube growth necessary for double fertilization

The rapid extension of the pollen tube is closely linked to the production of callose plugs in angiosperms (Williams, 2008). The number of callose plugs reportedly can serve as an indicator of the pollen tube growth rate in Hibiscus moscheutos (Snow and Spira, 1991). In contrast, in some gymnosperms, including Cycas and Pinus species, the pollen tubes have no or only a few callose plugs and grow slowly (Fernando et al., 2010). The results of the current study provide evidence of the positive association between the periodic formation of callose plugs and the pear pollen tube growth rate (Figure 1). Consistent with this finding, fast-growing pollen tubes with a considerable abundance of callose plugs have been detected in A. thaliana and Solanum lycopersicum (Qin et al., 2012).

Callose plugs divide pollen tubes into the highly vacuolated distal region and the apical cytoplasmic flow region. This spatial distribution allows the cytoplasm to concentrate at the tip for the synthesis of cell wall materials needed for growth (Guan et al., 2013; Li et al., 2020b). Moreover, the periodic formation of callose plugs can “reset” the length of the apical cytoplasmic flow region to maintain the continuous turgor pressure at the tip for the polar growth of the pollen tube (Dumais, 2021). Furthermore, after callose plugs form, the plasma membrane H+-ATPase mediates the development of a pH gradient in the apical cytoplasmic flow region that results in the tip being exposed to a slightly acidic environment; this stimulates actin polymerization to enhance the efficient delivery of vesicles that provide the cell wall material needed for pollen tube growth (Certal et al., 2008; Hepler et al., 2013). A decrease in the number of callose plugs due to the down-regulated expression of AtCalS5 and SHY (encoding a leucine-rich repeat protein) severely inhibits pollen tube growth in A. thaliana (Mizuta and Higashiyama, 2014) and Petunia hybrida (Guyon et al., 2004), respectively. A similar phenotype was observed following the suppression of callose synthesis by the 2DDG or as-ODN-PbrCalS1B.1 treatments of pear pollen tubes. The lack of callose plugs substantially delayed the growth and shortened the lifecycle of pear pollen tubes (Figures 2 and 6). These findings imply that the callose plug function related to the maintenance of healthy pollen tube growth may be conserved in the plant kingdom.

The members of the CalS1B cluster may be conserved genes that regulate callose synthesis in pollen tubes

The CalS gene family in A. thaliana comprises 12 members (AtCalS1–AtCalS12), of which five genes (AtCalS5, AtCalS9, AtCalS10, AtCalS11, and AtCalS12) are involved in pollen development during reproduction (Dong et al., 2005; Enns et al., 2005; Töller et al., 2008). However, AtCalS5 is the only CalS gene that has been reported to regulate callose synthesis during the period from pollen development to pollen tube elongation (Abercrombie et al., 2011). Previous research demonstrated that the disruption of AtCalS5 expression in A. thaliana decreases callose synthesis and restricts pollen tube growth, ultimately leading to decreased fertility (Dong et al., 2005; Mizuta and Higashiyama, 2014). Interestingly, the CalS1B gene cluster in Rosaceae was revealed to be orthologous to AtCalS5 in the present study (Figure 3; Supplemental Figure 4). Additionally, all of the available transcriptome data for the pollen of Rosaceae species indicated that the CalS1B genes in P. bretschneideri, M. domestica, P. mume, and P. avium were all highly expressed in pollen, with expression patterns similar to that of AtCalS5 (Figure 4; Supplemental Figure 3). Thus, by inserting the overexpression vector pro35S::PbrCalS1B.1 into the atcals5-1 A. thaliana mutant, we determined that PbrCalS1B.1 can reverse the reproduction-related defects, including the decreases in the pod length and number of seeds, the absence of callose plugs, and the deformation of the pollen cell wall (Figure 5). These results imply that the biological functions of PbrCalS1B.1 and perhaps other members of the CalS1B cluster in Rosaceae are similar to that of AtCalS5; the encoded proteins maintain pollen fertility and pollen tube growth by controlling callose synthesis. Notably, among other AtCalS5 homologs, NaGSL1 (Brownfield et al., 2008) and CcCalS5 (Abercrombie et al., 2011) are expressed at high levels in the pollen tubes of Nicotiana tabacum and Cabomba caroliniana, respectively. This implies that the functions of the CalS1B gene cluster associated with pollen fertility may be broadly conserved in many plant species.

PbrbZIP52 regulates PbrCalS1B.1 expression to activate callose biosynthesis

Previous reports have described the regulatory effects of bZIP transcription factors on callose deposition, which influence diverse biological processes in Oryza sativa and Manihot esculenta, including root growth (Das et al., 2019) and the response to pathogens (Li et al., 2017). Other research involving A. thaliana detected the substantial down-regulated expression of CalS2, CalS5, CalS7, CalS9, and CalS10 in the mature pollen of bzip34 and bzip18 mutants (Drábková and Honys, 2017), reflecting the positive regulatory effects of these transcription factors. Notably, several bZIP transcription factors in A. thaliana, such as AtbZIP18, AtbZIP34, and AtbZIP61, were referred briefly to express in pollen and interact with AtbZIP52, while the detailed molecular functions have not been studied (Gibalová et al., 2009, 2017). In O. sativa and N. tabacum, other bZIP transcription factors, including OsbZIP71, OsbZIP73, and NtBZI-1, reportedly regulate pollen development by affecting carbohydrate transport (Iven et al., 2010; Liu et al., 2019). On the basis of the systematic study of the CalS gene family, some gene members, such as AtCalS5 and PbrCalS1B.1, encode regulators of callose synthesis that function throughout the pollen development and pollen tube growth stages. Therefore, we speculated that the CalS regulatory function is controlled by bZIP transcription factors. In this study, we revealed that PbrbZIP52 is homologous to AtbZIP52, and its expression pattern is highly associated with PbrCalS1B.1 expression during the lifecycle of pear pollen tubes. The silencing of PbrbZIP52 substantially inhibited PbrCalS1B.1 expression, leading to restricted callose synthesis and the premature senescence of pollen tubes (Figure 7).

The bZIP transcription factors bind to the A-box cis-element containing the ACGT core in the promoters of target genes to affect various cellular processes, including abiotic stress responses (Ji et al., 2015; Fang et al., 2021) and the maintenance of photosystem II (Li et al., 2020a). Currently, there is accumulating evidence, suggesting that the binding of bZIP transcription factors to the A-box elements also modulates reproductive processes. The TGACG motif-binding bZIP transcription factors bind to the A-box element in the ARABIDOPSIS THALIANA HOMEOBOX GENE1 promoter to activate transcription, which is essential for flowering and the normal development of the inflorescence architecture (Wang et al., 2019). In rice, the nucleus-localized bZIP transcription factor OsRE1 binds to the A-box element in the Early heading date 1 promoter, with the resulting change in gene expression adjusting the heading date to improve the adaptability of O. sativa during the reproductive period (Chai et al., 2021). In this study, only two A-box elements were identified in the PbrCalS1B.1 promoter (between positions −219 and −291 bp), whereas other ACGT core cis-elements were not detected. The results of the Y1H assay, EMSA, and transcriptional activity assay confirmed that the specific binding of PbrbZIP52 to the PbrCalS1B.1 promoter requires the targeted recognition of the A-box element (Figure 8). These findings suggest the specific binding of bZIP transcription factors to the A-box element in target gene promoters may occur during the reproduction of additional plants.

Considered together, the data generated in this study suggest that PbrbZIP52 regulates PbrCalS1B.1 transcription by binding to the promoter A-box element to increase the synthesis of callose plugs during the normal lifecycle of pear pollen tubes (Figure 9). The study results presented in this article provide the theoretical basis for breeding high-yielding pear varieties and stabilizing the fruit setting rate of pear and other fruit tree species.

Materials and methods

Plant materials

All samples were harvested from 10-year-old P. bretschneideri “Dangshansuli” trees growing in the orchard of Nanjing Agriculture University, Nanjing, China (latitude 118°50′E; longitude 32°2′N). All pollination experiments in vivo carried out in this study were cross-compatible between “Cuiguan” as female parent and “Dangshansuli” as male parent. Besides, “Dangshansuli” pollen was employed for pollen culture in vitro. Pollen grains were obtained from mature flowers (before pollen dispersal), dried in desiccant-filled bottles, and stored at −20°C until needed. The pollen grains were germinated in liquid medium [0.55 mM Ca(NO3)2, 1.60 mM H3BO3, 1.60 mM MgSO4, 1.00 mM KNO3, 440 mM sucrose, and 5 mM MES, pH 6.0–6.2] and then cultured under in vitro conditions. Treated pollen tubes were examined using the EZ4 HD light microscope (Leica Microsystems, Wetzlar, Germany), and the pollen tube lengths were measured using ImageJ software (https://ij.imjoy.io/). Columbia ecotype A. thaliana (Col-0) plants were selected as the wild-type control. The atcals5-1 mutant plants (Salk_009234; Columbia genetic background) were obtained from the Arabidopsis Biological Resource Center (https://abrc.osu.edu/).

Aniline blue staining for the analysis of the callose content and callose plugs

To analyze the distribution of callose in pollen tubes, a suspension containing germinated pollen tubes was stained with a 0.01% (w/v) aniline blue solution for 5 min, after which the fluorescence signals of the callose plugs were examined using the Fluoview FV1200 confocal microscope (Olympus, Tokyo, Japan).

The germination of pollen grains on the stigma was analyzed by placing the samples in an FAA fixative (95% ethanol, 37% glacial acetic acid, formaldehyde; 18:1:1, v/v) at room temperature for 24 h (Li et al., 2020b). The samples were purified in a solution comprising 95% ethanol:acetic acid (3:1, v/v) and then washed several times in 1 M phosphate buffer (pH 7.0). Alternatively, the pollinated stigmas were treated with 1 M NaOH at 65°C for 2 h and then stained with 0.01% (w/v) aniline blue prepared in 100 mM K3PO4 buffer. The growth and deposition of callose plug of pollen tubes in vivo were examined using the M205FA stereomicroscope (Leica Microsystems) and the Fluoview FV1200 confocal microscope (Olympus, Tokyo, Japan), respectively.

The relative callose content in the pollen tubes was determined as previously described (Köhle et al., 1985). Briefly, pollen samples (100 mg) were treated with 95% (v/v) ethanol for 2 min and then ground in 5 mL of 1 M NaOH. The ground material was centrifuged at 10,000 × g for 15 min at room temperature. A 200 μL aliquot of the supernatant was added to a solution consisting of 800 μL of 0.1% (w/v) aniline blue, 200 μL of 1 M HCl, and 1,800 μL of 1 M glycine/NaOH buffer (pH 9.5), which was then mixed vigorously for 20 min at 50°C. Alternatively, the samples were incubated for 30 min at room temperature, after which the fluorescence intensity was evaluated using the Infinite M200 PRO microplate reader (Tecan, Männedorf, Switzerland). Aniline blue fluorescence was detected at excitation and emission wavelengths of 390 and 420 nm, respectively.

As previously described (Riehl and Jaffe, 1984), 2DDG (Sigma) was used to inhibit CalS activity in pear pollen tubes. Specifically, three 2DDG concentrations (0.1, 0.5, and 1.0 mM) were tested.

Evaluation of PCD in pear pollen tubes

PCD was assayed by TUNEL staining as previously described (Chen et al., 2018). The pollen tubes were fixed in 4% (w/v) paraformaldehyde for 2 h and then transferred to 70% (v/v) ethanol for an overnight incubation at −20°C. The Dead End Fluorometric TUNEL system (Promega, Beijing, China) was used to examine the PCD in the samples. After the samples were washed with citrate buffer (pH 4.1), the pollen tubes were stained with 50 μg mL−1 DAPI for 10 min at room temperature and then analyzed using the Fluoview FV1200 confocal microscope (Olympus). The TUNEL signal was detected at excitation and emission wavelengths of 540 and 620 nm, respectively. The DAPI signal was detected at excitation and emission wavelengths of 360 and 420 nm, respectively. Positive TUNEL staining in the pollen tubes was indicative of PCD.

Identification and phylogenetic analysis of the CalS and bZIP family in Rosaceae

Multiple strategies were used to search for members of the CalS and bZIP family in Rosaceae. (1) Keyword search for annotated protein databases; (2) hidden Markov model (HMM) searches with the conservative domain HMM profile (CalS (PF04652, PF14288 and PF02364), bZIP (PF00170, PF07716 and PF12498) for proteins, and then evaluated all significant hits (HMMER E-value < e−5); (3) A. thaliana CalS and bZIP protein sequences were used as queries in exhaustive BLASTP searches with standard parameters until there was convergence with the protein database. All candidate protein sequences were analyzed using the InterProScan5 database (http://www.ebi.ac.uk/interpro/search/sequence-search) to verify the presence of the necessary domain and remove protein sequences that lack these domains. The protein sequences were compared by a multiple sequence alignment using the MAFFT software. A maximum likelihood phylogenetic tree was constructed using IQ-TREE2, with 1,000 replications. Identifying Rosaceae CalS and bZIP genes are named in Supplemental Table 2. The sequence alignment of CalS and bZIP proteins used for the phylogenetic analysis is listed in Supplemental File 1 and File 2, respectively.

Transcript assembly and quantification for RNA-seq

We mined 40 publicly available transcriptome libraries that include data for multiple organizations in Rosaceae. All libraries data were mapped to the reference genome using the HISAT2 aligner. The percentage of uniquely aligned reads varied among samples above 80% (Supplemental Table 3), indicating that the quality of these RNA-seq data was acceptable. The expression level of each isoform was represented by the Raw read counts and RPKM values, which is calculated based on the length of the fragments and the read count mapped to a specific fragment. The StringTie software was used to calculate the RPKM values. Genes with a higher RPKM value indicated increased expression.

Antisense oligonucleotide-based inhibition of gene expression

We silenced the expression of target genes using as-ODNs as previously described (Chen et al., 2018). Sense and antisense oligonucleotide sequences (s-ODN and as-ODN) targeting PbrCalS1B.1 and PbrbZIP52 were designed using the RNAfold web server (http://rna.tbi.univie.ac.at/cgi-bin/RNAfold.cgi). The candidates were then assessed (i.e. in terms of antisense efficiency) using the Soligo software (http://sfold.wadsworth.org/cgi-bin/soligo.pl). The three as-ODN candidates with the highest scores were synthesized with phosphorothioate modifications at the 5′ and 3′ ends and then purified using a high-performance liquid chromatography system. A mixture comprising ODN/cytofectin complexes was incubated in cultivation medium for 15 min before it was added to the pollen culture for a final concentration of 50 mM ODN. After the addition of 15 mg mL−1 ODN, the pollen was cultivated in this mixture for 6 h. The as-/s-ODN pairs were tested for their effects on the relative expression levels of target genes. The most effective pair (#1 in Supplemental Table 1) was selected for further analyses.

PbrCalS1B.1 expression in A. thaliana

To express PbrCalS1B.1 in transgenic A. thaliana plants, the PbrCalS1B.1 promoter (proPbrCalS1B.1) was inserted into the pGWB633 vector containing the GUS-encoding reporter gene, whereas the full-length PbrCalS1B.1 coding sequence was inserted into the pGWB602 vector to construct an overexpression vector using the Gateway system (Invitrogen). The recombinant overexpression plasmid was inserted into Agrobacterium tumefaciens GV3101 cells for the transformation of A. thaliana plants according to the floral dip method (Clough and Bent, 1998). Positive transgenic plants were identified by RT-PCR, which was performed using specific primers designed on the basis of the target gene and plasmid sequences. The T3 generation transgenic A. thaliana plants were analyzed in this study. Primer sequences are provided in Supplemental Table 1. A. thaliana plants with intact floral organs were stained with GUS staining solution [0.5 M sodium phosphate buffer (pH 7.2), 10% (v/v) Triton-X, and 100 mM potassium ferricyanide; Sigma] at room temperature for 25 min. After adding 2 mM 5-bromo-4-chloro-3-indolyl-β-D-galactoside (X-Gal) and adjusting the pH to 7.0, the samples were stained at 37°C for 12 h. The staining reaction was terminated by adding 0.2 M Na2CO3. The samples were rinsed with 70% ethanol to eliminate autofluorescence and then examined using the M205FA stereomicroscope (Leica Microsystems).

Four-week-old Arabidopsis plants with different genotypes were selected for a phenotypic analysis. The pollen grain structure was analyzed using the JSM-8404 scanning electron microscope (Leica Microsystems) and photographed.

Subcellular localization of PbrCalS1B.1

To determine the subcellular localization of PbrCalS1B.1, the full-length PbrCalS1B.1 coding sequence lacking the terminator was inserted into the XbaI/BamHI-digested pUC19 vector to construct the pUC19-PbrCalS1B.1-GFP expression vector. Protoplasts isolated from 6-week-old wild-type A. thaliana (Col-0) leaves were transformed with the expression vector using PEG-Ca2+ as previously described (Wu et al., 2009). After incubating at room temperature for 12 h, the fluorescence signals of GFP (excitation and emission wavelengths of 488 and 510 nm, respectively) and chlorophyll (excitation and emission wavelengths of 640 and 675 nm, respectively) in the protoplasts were detected using the Fluoview FV1200 confocal microscope (Olympus). Primer sequences are provided in Supplemental Table 1.

PbrCalS1B.1 point mutation and Y1H assay

To assess the ability of PbrbZIP52 to bind to PbrCalS1B.1 promoter fragments, the A-box elements of proPbrCalS1B.1 were mutated (m1, m2, and m1 & m2) using the Fast Mutagenesis System (TransGen Biotech, Beijing, China). The Y1H analysis was performed using the Matchmaker Gold Yeast One-Hybrid Library Screening System (TaKaRa, Dalian, China). The proPbrCalS1B.1 fragments generated by the PCR were cloned into the KpnI and SalI sites of the pABAi vector to construct the bait plasmids (pAbAi-proPbrCalS1B.1WT, pAbAi-proPbrCalS1B.1m1, pAbAi-proPbrCalS1B.1m2, and pAbAi-proPbrCalS1B.1m1&m2). Different AbA concentrations were evaluated in terms of their inhibitory effects on the auto-activation in the Y1H system. The full-length PbrbZIP52 coding sequence was cloned into the BamHI and NdeI sites of the pGADT7 vector to produce the prey plasmid pGADT7-PbrbZIP52, which was then inserted into the Y1H-Gold yeast cells containing the bait plasmids. The transformed yeast cells were added to SD/−Leu/−Ura medium containing X-Gal and 200 ng mL−1 AbA and then incubated for 3 days at 30°C for the final determination of positive interactions. Experiments included pGADT7-p53+pAbAi-p53 as the positive control and pGADT7+pAbAi-PbrCalS1B.1WT as the negative control. Primer sequences are provided in Supplemental Table 1.

Electrophoretic mobility shift assay

To purify the PbrbZIP52 protein for the EMSA experiments, the full-length PbrbZIP52 gene was amplified by the PCR and incorporated into the pGEX-4T-1 vector at the SmaI and XhoI sites to generate the GST-PbrbZIP52 construct. The construct was introduced into Escherichia coli BL21 (DE3) cells to express the target protein, which was then purified using glutathione sepharose 4B beads (GE Healthcare, Little Chalfont, UK). The proPbrCalS1B.1 oligonucleotide sequence containing the A-box (ACGT) element was synthesized by Sangon (Shanghai, China). Oligonucleotide probes for the wild-type and point-mutated PbrCalS1B.1 promoters were labeled using the Biotin 3′ End DNA Labeling Kit (Pierce, USA). The in vitro binding of the GST-PbrbZIP52 fusion protein to the biotin-tagged proPbrCalS1B.1 was examined using the LightShift Chemiluminescent EMSA kit (Thermo Fisher Scientific, Waltham, MA, USA). The mixture was incubated in binding buffer for 30 min at room temperature, and the protein–DNA complexes were separated in a 10% native polyacrylamide gel. The labeled probe signal was detected by electroblotting according to the manufacturer’s instructions. Biotin-labeled probes incubated with GST proteins served as the negative control. Moreover, five- and 25-fold molar excesses of unlabeled probes were used for the competitive binding analysis.

Transient transcriptional activity assays

The PbrbZIP52 coding sequence was cloned into the modified pBI221 vector without GUS to produce the effector plasmid. Additionally, proPbrCalS1B.1 and a modified LUC reporter gene obtained from the pGL3-basic plasmid were inserted into the pBI221 vector (CaMV 35S promoter removed) to construct the reporter plasmid. The CaMV35S::GUS construct was used for normalizing data, whereas the empty vector lacking PbrbZIP52 served as the negative control. A. thaliana mesophyll cell protoplasts were prepared and transfected as previously described (Yoo et al., 2007). Specifically, the recombinant vectors were inserted into protoplasts via PEG-mediated transformation. After an overnight incubation in darkness, the LUC activity was measured using the Dual Luciferase Reporter Analysis System (Promega, Madison, WI, USA). The activation of proPbrCalS1B.1 by PbrbZIP52 was evaluated on the basis of the LUC:GUS ratio. Primer sequences are provided in Supplemental Table 1.

Statistical analysis

All experimental data are the averages of at least three independent replicates and are shown as the mean ± Se. The data were analyzed using SPSS software, and statistical differences were compared based on Student’s t test for two groups of samples and ANOVA for multiple samples. ANOVA tables are provided in Supplemental Data Set 1. For multiple comparisons, Tukey’s honestly significant difference test was performed.

Availability of data and materials

Publicly available data for pear (P. bretschneideri) genome and annotated protein sequences were downloaded from the Genome Database for Rosaceae (https://www.rosaceae.org/). A. thaliana data were downloaded from The Arabidopsis Information Resource (https://www.arabidopsis.org/). Gene identity information is provided in Supplemental Table 2.

Accession numbers

The complete PbrCalS1B.1 and PbrbZIP52 sequences were deposited in the GenBank database under accession numbers OP207888 and OP207889, respectively.

Supplementary Material

kiad002_Supplementary_Data
Click here for additional data file. (3.1MB, zip)

Acknowledgments

We thank Liwen Bianji (Edanz) (www.liwenbianji.cn) for editing the English text of a draft of this manuscript.

Contributor Information

Zhongheng Xia, College of Horticulture, Fujian Agriculture and Forestry University, Fuzhou 350002, China.

Binxu Wen, College of Horticulture, Fujian Agriculture and Forestry University, Fuzhou 350002, China.

Jing Shao, Institute of Pomology, Jilin Academy of Agricultural Sciences, Gongzhuling 136100, China.

Tianci Zhang, College of Horticulture, Fujian Agriculture and Forestry University, Fuzhou 350002, China.

Mengmeng Hu, College of Horticulture, Fujian Agriculture and Forestry University, Fuzhou 350002, China.

Lin Lin, Anxi College of Tea Science, Fujian Agriculture and Forestry University, Anxi 362406, China.

Yiping Zheng, Fujian Academy of Agricultural Sciences Biotechnology Institute, Fuzhou 350003, China.

Zhixin Shi, College of Horticulture, Fujian Agriculture and Forestry University, Fuzhou 350002, China.

Xinlin Dong, College of Horticulture, Fujian Agriculture and Forestry University, Fuzhou 350002, China.

Juanjuan Song, College of Horticulture, Fujian Agriculture and Forestry University, Fuzhou 350002, China.

Yuanshan Li, College of Horticulture, Fujian Agriculture and Forestry University, Fuzhou 350002, China.

Yongjie Wu, College of Horticulture, Fujian Agriculture and Forestry University, Fuzhou 350002, China.

Yafang Yuan, Department of Horticulture and Landscape Architecture, Fujian Vocational College of Agriculture, Fuzhou 350119, China.

Juyou Wu, Center of Pear Engineering Technology Research, State Key Laboratory of Crop Genetics and Germplasm Enhancement, College of Horticulture, Nanjing Agricultural University, Nanjing 210095, China.

Qingxi Chen, College of Horticulture, Fujian Agriculture and Forestry University, Fuzhou 350002, China.

Jianqing Chen, College of Horticulture, Fujian Agriculture and Forestry University, Fuzhou 350002, China.

Supplemental data

The following materials are available in the online version of this article.

Supplemental Figure S1. Deposition of callose plugs in pollen tube of “Dangshansuli” stigma.

Supplemental Figure S2. Analysis of CalS gene domains and structures in Rosaceae species and Arabidopsis thaliana.

Supplemental Figure S3. Tissue-specific expression patterns of CalS genes in apple, plum, and cherry.

Supplemental Figure S4. Multiple sequence alignment of CalS proteins.

Supplemental Figure S5. RT-PCR analysis of the flowers from PbrCalS1B.1-overexpressing transgenic Arabidopsis thaliana plants in the T3 generation.

Supplemental Figure S6. As-ODN effects on elongating pollen tubes.

Supplemental Figure S7. Analysis of the inhibitory effect of as-ODN on PbrCalS1B.1 expression.

Supplemental Figure S8. Fluorescein diacetate staining to assess pollen tube viability.

Supplemental Figure S9. Identification of the bZIP gene family in Rosaceae.

Supplemental Figure S10. Analysis of the expression pattern of PbrbZIP52.

Supplemental Table S1. Primers used in this study.

Supplemental Table S2. Information regarding CalS and bZIP family identities.

Supplemental Table S3. Information regarding the samples used for the RNA-seq analysis.

Supplemental Data Set S1. ANOVA tables.

Supplemental File S1. Text file for the sequence alignment of CalS used for the phylogenetic analysis in Figure 3.

Supplemental File S2. Text file for the sequence alignment of bZIP used for the phylogenetic analysis in Supplemental Figure S9.

Funding

This work was supported by the National Natural Science Foundation of China (31801814 and 32172543), the Natural Science Foundation of Fujian Province, China (2022J01137), the Fujian Agriculture and Forestry University Natural Science Funds for Distinguished Young Scholar (xjq21005), the Training Funds for Core Young Scholar of Horticulture College, Fujian Agriculture and Forestry University (722022011), and the earmarked fund for the Earmarked Fund for China Agriculture Research System (CARS-28).

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Associated Data

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

Supplementary Materials

kiad002_Supplementary_Data
Click here for additional data file. (3.1MB, zip)

Data Availability Statement

Publicly available data for pear (P. bretschneideri) genome and annotated protein sequences were downloaded from the Genome Database for Rosaceae (https://www.rosaceae.org/). A. thaliana data were downloaded from The Arabidopsis Information Resource (https://www.arabidopsis.org/). Gene identity information is provided in Supplemental Table 2.

Accession numbers

The complete PbrCalS1B.1 and PbrbZIP52 sequences were deposited in the GenBank database under accession numbers OP207888 and OP207889, respectively.

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