Auxin inhibits lignin and cellulose biosynthesis in stone cells of pear fruit via the PbrARF13‐PbrNSC‐PbrMYB132 transcriptional regulatory cascade

Summary

Stone cells are often present in pear fruit, and they can seriously affect the fruit quality when present in large numbers. The plant growth regulator NAA, a synthetic auxin, is known to play an active role in fruit development regulation. However, the genetic mechanisms of NAA regulation of stone cell formation are still unclear. Here, we demonstrated that exogenous application of 200 μm NAA reduced stone cell content and also significantly decreased the expression level of PbrNSC encoding a transcriptional regulator. PbrNSC was shown to bind to an auxin response factor, PbrARF13. Overexpression of PbrARF13 decreased stone cell content in pear fruit and secondary cell wall (SCW) thickness in transgenic Arabidopsis plants. In contrast, knocking down PbrARF13 expression using virus‐induced gene silencing had the opposite effect. PbrARF13 was subsequently shown to inhibit PbrNSC expression by directly binding to its promoter, and further to reduce stone cell content. Furthermore, PbrNSC was identified as a positive regulator of PbrMYB132 through analyses of co‐expression network of stone cell formation‐related genes. PbrMYB132 activated the expression of gene encoding cellulose synthase (PbrCESA4b/7a/8a) and lignin laccase (PbrLAC5) binding to their promotors. As expected, overexpression or knockdown of PbrMYB132 increased or decreased stone cell content in pear fruit and SCW thickness in Arabidopsis transgenic plants. In conclusion, our study shows that the ‘PbrARF13‐PbrNSC‐PbrMYB132’ regulatory cascade mediates the biosynthesis of lignin and cellulose in stone cells of pear fruit in response to auxin signals and also provides new insights into plant SCW formation.

Introduction

Pears (Pyrus spp.) are economically important tree crops and popular with consumers because the fruit is juicy and has a sweet and sour flavour. Based on their geographical origins, pears are divided into Asian and European types that differ significantly in the ripening process, skin colour, taste and number of stone cells in the fruit flesh (Hiwasa et al., 2004; Wu et al., 2013; Yao et al., 2017; Zhang et al., 2014). These differences result from the independent domestication processes for Asian and European pears (Wu et al., 2018). The genetics of stone cell formation and the significance of their presence are currently unclear, although previous studies have shown that they are mainly formed during the early stages of pear fruit development by the thickening of secondary cell walls (SCWs) (Smith, 1926; Tao, 2009).

Secondary cell walls form through the deposition of lignin, cellulose and hemicellulose between the primitive cell walls and cell membrane to provide thickened cell walls for stronger mechanical support to plant cells (Hofte and Voxeur, 2017; Meents et al., 2018; Sakamoto et al., 2018; Terrett and Dupree, 2019; Zhang et al., 2021a; Zhong and Ye, 2015). Transcription factors (TFs) of NAC and MYB families play key regulatory roles in SCW formation, and their regulatory functions are relatively conserved among plant species (Johnsson et al., 2019; Kubo et al., 2005; Wang and Dixon, 2012; Zhao, 2016). Similarly, lignin and cellulose are major components of stone cells (Zhang et al., 2021b), which further implies that stone cells are thick‐walled cells with SCW, and that the SCW TFs may involve in the regulation of stone cells. As shown in previous studies, PbrMYB169 is a transcriptional activator of lignin biosynthesis that regulates stone cells lignification by binding to the AC elements in the promoters of genes related to lignin biosynthesis (Xue et al., 2019b); PbKNOX1 is a negative regulator of lignin metabolism that inhibits stone cell lignification (Cheng et al., 2019). Moreover, our research identified an important positive regulator of lignin and cellulose biosynthesis in stone cells, PbrNSC, through eQTL and co‐expression network analyses of lignin and cellulose pathway‐related genes (Wang et al., 2021). Thus, these studies provide guidance for our subsequent stone cells studies.

Many factors can influence the formation of SCW, among which hormones play a prominent role (Didi et al., 2015; Le Gall et al., 2015; Moura et al., 2010). Plant hormones including cytokinin (De Rybel et al., 2014; Delay et al., 2013), abscisic acid (ABA; Ramachandran et al., 2021), gibberellic acid (GA; Huang et al., 2015), ethylene (Pesquet and Tuominen, 2011) and brassinolide (BR; Du et al., 2020) have been reported to affect SCW formation. Auxin can have a various levels of effect on the SCW formation depending on its concentration and plant tissue types. For example, in Arabidopsis and Zinnia, auxin stimulated the differentiation of mesophyll cells into xylem tubular cells (Fukuda, 1997; Minami and Fukuda, 1995) while it inhibited the thickening of the pollen cell wall (Cecchetti et al., 2008). However, the effect of auxin on SCW formation in the stone cells is still unclear and deserves to be explored.

Auxin exerts its functions through auxin response factors (ARFs) (Guilfoyle et al., 1998; Guilfoyle and Hagen, 2007; Korasick et al., 2014). Therefore, ARFs are involved in a wide variety of plant developmental processes (Korasick et al., 2014; Ulmasov et al., 1999a,b) such as plant embryo formation (Mallory et al., 2005; Mao et al., 2020; Zhang et al., 2018), root growth (Attia et al., 2009; Fukaki and Tasaka, 2009; Nagpal et al., 2005; Zhang et al., 2022a), leaf blade extension (Uzair et al., 2021), flower organ development (Nagpal et al., 2005), senescence (Cecchetti et al., 2008; Nagpal et al., 2005) and nutrient uptake (Qi et al., 2012). Also, ARFs play important roles in regulating fruit quality in horticultural crops. For example, SlARF6 regulates chlorophyll biosynthesis, photosynthesis, sugar accumulation and fruit development in tomatoes (Yuan et al., 2019); in apples, MdARF5 binds to the promoters of MdACS3a, MdACS1 and MdACO1 to promote ethylene biosynthesis in fruit (Yue et al., 2020). MdARF13 interacts with MdMYB10 to inhibit anthocyanin accumulation (Wang et al., 2018). However, the possible involvement of AFRs in stone cells formation in pear fruit has not been reported.

In this study, we investigated the inhibition of stone cell formation by exogenous NAA application. Strikingly, we found that NAA is involved in the inhibition of lignin and cellulose biosynthesis in the stone cells by inducing the expression of the auxin response factor PbrARF13. We showed that PbrARF13 directly binds to the promoter of PbrNSC, a key gene for lignin and cellulose biosynthesis in stone cells (Wang et al., 2021), to inhibit its expression, thereby reducing stone cell content. We also provide evidence that PbrNSC positively regulates the expression of PbrMYB132, which encodes a protein that can bind to the promoters of genes encoding cellulose synthase (PbrCESA4a/7b/8c) and laccase (PbrLAC5) to promote cellulose and lignin biosynthesis in stone cells. Taken together, our study highlights the important role of the ‘PbrARF13‐PbrNSC‐PbrMYB132’ signalling cascade in the inhibition of lignin and cellulose biosynthesis in stone cells and provides novel insights for studying the roles of phytohormones on stone cells formation in pear fruit and SCW formation in plants.

Results

NAA treatments reduce stone cell content and decrease the expression levels of genes related to lignin and cellulose biosynthesis in stone cells

Several concentrations of NAA (0, 100, 200 and 300 μm) were sprayed on ‘Dangshansuli’ fruit at 20–21 days after full bloom (DAFB) and the fruit were sampled at 28, 35 and 49 DAFB for multiple analyses. The results showed that NAA treatments significantly reduced fruit stone cell content and that the number of stone cells was negatively correlated with NAA concentration (Figure 1a,b). Moreover, the content of lignin and cellulose in fruit flesh also showed a negative correlation with NAA concentration when they were tested at 35 DAFB (the key stage for stone cells formation) (Figure 1d,e). However, the effects of NAA concentration on fruit growth varied. The lower concentrations of exogenous NAA (100 and 200 μm) used in the experiment increased fruit weight, while 300 μm NAA was less effective in promoting fruit development than 200 μm (Figure 1a,c). To achieve a balanced effect between stone cell content and fruit growth, we selected the 200 μm NAA treatment for further experiments. Then, we sprayed 200 μm NAA on the fruit of six Chinese pear cultivars and found that fruit treated with 200 μm NAA had higher fruit weight and lower stone cell content at 35 DAFB and maturity (Figure S1). This result indicates that the 200 μm NAA treatment may be potentially useful for improving pear fruit production and quality.

Figure 1.

NAA treatment of ‘Dangshansuli’ fruit reduced the content of stone cells, lignin and cellulose. (a) Images show whole and sectioned pear fruit at three developmental stages (28, 35 and 49 DAFB, days after full bloom) under four treatments with different NAA concentrations (0, 100, 200 and 300 μm). The fruit sections were stained red with phloroglucinol‐HCl to show the presence of lignin. (b‐e) Fruit samples similar to those shown in (a) were used to analyse single fruit weight (c) and the content of the stone cells (b), lignin (d) and cellulose (e) in fruit flesh tissues (n ≥ 10 biological replicates). Different letters above the bars indicate a significant difference (P < 0.05) obtained by the one‐way ANOVA test. (f) Heatmap shows downregulation of two transcription factor (TF) genes, three lignin and three cellulose biosynthesis genes in fruit treated with 200 μm NAA compared to those treated with 0 μm NAA, at 35 DAFB. The legend of the heatmap was subjected to Min‐max normalization. (g) Relative expression levels of eight genes related to secondary cell wall (SCW) formation in stone cells were determined in ‘Dangshansuli’ fruit at 35 DAFB under 0 or 200 μΜ NAA treatment, using qRT‐PCR analysis. The expression level of each gene was standardized against the reference gene (PbrGAPDH). Statistical significance was determined by a one‐tailed paired t‐test. *P < 0.05. Values are mean ± SD of three technical replicates (three biological replicates have similar expression trends, we selected one for presentation).

Next, to understand the genetic mechanism of NAA inhibition of lignin and cellulose biosynthesis in stone cells, RNA‐seq analyses were performed using fruit treated with 0 and 200 μm NAA at 35 DAFB. The results showed that NAA treatment reduced the expression level of PbrNSC (Figure 1f), the key gene that positively regulates lignin and cellulose biosynthesis in stone cells (Figure S2a; Wang et al., 2021). In addition, the analyses also showed reduced expression of the TF gene PbrMYB169; the lignin biosynthesis genes Pbr4CL4, PbrLAC4, PbrLAC5; and the cellulose biosynthesis genes PbrCESA4a, PbrCESA7a and PbrCESA8b (Figure 1f). These genes, which are the target of PbrNSC, are known to be involved in the lignin and cellulose biosynthesis in stone cells (Figure S2b; Wang et al., 2021), and the reduced expression of these genes after NAA treatment was confirmed by quantitative real‐time polymerase chain reaction (qRT‐PCR) analyses (Figure 1g). The phenotypic and gene expression data together indicate that exogenous NAA treatment reduces stone cell content in fruit by inhibiting the expression of genes related to lignin and cellulose biosynthesis.

PbrARF13 binds to the PbrNSC promoter and negatively regulates its expression

Because PbrNSC is considered to be a ‘switch’ for lignin and cellulose biosynthesis in stone cells (Figure S2; Wang et al., 2021), we were interested in understanding how NAA inhibits the expression of PbrNSC. We hypothesized that PbrNSC may be connected to auxin through PbrARFs, because auxin function depends on ARFs to regulate gene expression. To test this hypothesis, all 31 PbrARFs identified in the ‘Dangshansuli’ pear genome (Table S1; Figure S3) were analysed for their ability to regulate PbrNSC expression using dual luciferase assays with the ‘35S::PbrARF_1–31‐GFP’ effector vector and the ‘PbrNSC (promoter)‐LUC’ reporter vector (Figure 2a). The results showed that PbrARF13 significantly inhibited PbrNSC promoter activity (Figure 2b), implying that PbrARF13 may negatively regulate the expression of PbrNSC in vivo.

Figure 2.

Auxin response factor PbrARF13 inhibited PbrNSC expression. (a) Schematic diagrams show effector and reporter vectors used for dual luciferase assays. (b) The effect of 31 PbrARFs on the activity of the PbrNSC promoter was tested using dual luciferase assays that showed a significant reduction of the promotor activity by PbrARF13. Statistical significance was determined by a one‐tailed paired t‐test. *P < 0.05. (c and d) Heatmap shows 200 μm NAA treatment had no significant effect on the expression level of PbrARF13 at 35 days after full bloom (DAFB) (c). At 28 DAFB, 200 μm NAA treatment increased PbrARF13 while decreasing PbrNSC expression levels (d). The legend of the heatmap was subjected to Min‐max normalization. (e and f) The expression level of PbrNSC (e) and PbrARF13 (f) and stone cells content in ‘Dangshansuli’ fruit was analysed at eight developmental stages between 21 and 160 DAFB after the treatment with 0 μm or 200 μm NAA. The expression of each gene was normalized to the reference gene PbrGAPDH. Values are mean ± SD of three technical replicates (three biological replicates have similar expression trends, we selected one for presentation).

At this point, if exogenous administration of NAA can induce the expression of PbrARF13, then PbrARF13 may be an important gene in the inhibition of lignin and cellulose biosynthesis in stone cells by NAA. We first examined the expression trends of PbrARF13 in the 35 DAFB‐RNA‐seq dataset, and the results showed that NAA did not have an apparent effect on PbrARF13 expression (Figure 2c). Considering the time‐sensitive nature of hormone action, we then performed RNA‐seq analysis using 28 DAFB fruit. Unexpectedly, 200 μm NAA treatment significantly increased the level of PbrARF13‐specific mRNA at 28 DAFB, while it reduced the relative expression of PbrNSC (Figure 2d). This result indicated that the effect of NAA treatment on PbrARF13 transcription occurred before 35 DAFB and that the gene response to the NAA signal was rapid.

To further analyse the relationship between PbrARF13 and PbrNSC, we examined the dynamic expression pattern (pear fruit development from 21 to 160 DAFB) of the two genes using qRT‐PCR, at the key stage of stone cells formation, PbrNSC mRNA levels were high, and 200 μm NAA treatment reduced PbrNSC expression levels (Figure 2e). In contrast, PbrARF13 mRNA levels were low at this key stage, but 200 μm NAA treatment increased the expression level of PbrARF13 (Figure 2f). Based on the qRT‐PCR expression levels, a dynamic fitting curve analysis showed that the expression levels of PbrNSC and PbrARF13 were negatively correlated in the absence of exogenous NAA treatment (Figure S4a and  S4b). This negative correlation relationship was enhanced by NAA treatment (Figure S4c). In addition, the negative correlation between the expression levels of PbrARF13 and PbrNSC was also confirmed by qRT‐PCR analysis in the other six pear cultivars (Figure S4d). Based on the above results, we postulate that in response to NAA, PbrARF13 may be involved in the PbrNSC‐mediated lignin and cellulose biosynthesis in stone cells.

Further analysis revealed that the PbrARF13 protein localizes to the nucleus and has conserved DBD and ARF structural domains. The ARF structural domain is rich in serine, glycine and proline (Figure S5), implying that PbrARF13 may act as a transcriptional repressor by negatively regulating the expression of downstream genes (Guilfoyle and Hagen, 2007; Ulmasov et al., 1999a,b).

PbrARF13 represses the activity of PbrNSC by binding to the auxin‐responsive element (AuxRE) in the promoter

To further understand the molecular mechanism by which PbrARF13 negatively regulates PbrNSC, we analysed the promoter sequence of PbrNSC and identified two potential ARF protein binding sites known as auxin‐responsive elements (AuxREs) (Figures 3a and S13a). A dual luciferase activation assay showed that PbrARF13 represses the promotor activities of the two PbrNSC promoter fragments (−1 to −1500 and −1 to −2000 bp) that contain AuxRE‐2 but not the other fragment (−1 to −1000 bp) without AuxRE‐2 (Figure 3b). This result was confirmed using the GUS reporter (Figure 3c). An electrophoretic mobility shift assay (EMSA) showed that PbrARF13 (containing a His tag) binds directly to the biotin‐labelled AuxRE‐2 probe (5′ biotin‐TGTCTC‐3′) to form a protein–DNA complex that has reduced electrophoretic mobility on a polyacrylamide gel. The relative intensity of this mobility‐shifted band was reduced as increasing amounts of unlabelled 5’‐TGTCTC‐3′ probes were added to the reaction, indicating that there was a competitive relationship between the labelled and unlabelled probe sequences. When the 5′ biotin‐TGTCTC‐3′ sequence was mutated to 5′ biotin‐GAGAGA‐3′, PbrARF13‐His was unable to bind to the mutated probe (Figure 3d). The results of unlabelled sequence competition and mutated probe analysis indicated that the binding of PbrARF13 to AuxRE‐2 is sequence specific. The binding of PbrARF13 to the PbrNSC promoter was further verified using a yeast one‐hybrid (Y1H). Yeast cells co‐expressing pGADT7‐PbrARF13 and pHIS2‐PbrNSC‐Promoter were able to grow on the selective medium (SD/−Leu/ −Trp /−His medium supplemented with 20 mM 3‐Amino‐1,2,4‐triazole). In contrast, the growth of yeast cells transformed with the empty control vector was poor (Figure 3e). The results of in vivo and in vitro experiments demonstrated that PbrARF13 binds to the 5’‐TGTCTC‐3′ sequence in the PbrNSC promoter to repress PbrNSC expression.

Figure 3.

PbrARF13 regulation of PbrNSC expression was dependent on the AuxRE‐2 (TGTCTC) element in the promoter. (a) Schematic diagrams show effector and reporter vectors used for dual luciferase and GUS activity assays. (b) Dual luciferase assay and (c) GUS activity was performed to analyse the activation effect of PbrARF13 on PbrNSC promoter segmentation. The promoter of PbrNSC was divided into three segments according to the distribution of AuxRE (Cis‐acting elements of auxin‐responsive genes). (d) Electrophoretic mobility shift assay (EMSA) showed that PbrARF13 binds directly to the AuxRE‐2 (TGTCTC, −1350–−1355) sequence in the PbrNSC promoter. Hot‐P and Mut‐P represent biotin‐labelled WT and mutation probes respectively; competitor‐P represents the unlabelled WT sequence. The black triangle represents the rates of competitors (10×, 50× and 100×). (e) Yeast one‐hybrid (Y1H) validates the ability of PbrARF13 to bind to the PbrNSC promoter. Yeast cells were co‐transformed with a bait vector (containing a PbrNSC promoter fragment fused to the HIS2 reporter gene) and a prey vector (containing PbrARF13 fused to the GAL4 activation domain). The yeast strains were grown on SD/‐Leu/‐Trp and SD/‐Leu/‐Trp/‐His with 20 mM 3‐AT (3‐amino‐1, 2, 4‐triazole) media for 3 d. 20 mM 3‐AT (3‐amino‐1,2,4‐triazole) was used to inhibit background growth and test the strength of the interaction. Statistical significance was determined by a one‐tailed paired t‐test. *P < 0.05. Values are mean ± SD of three technical replicates (three biological replicates have similar expression trends, we selected one for presentation).

PbrARF13 acts as a negative regulator to inhibit lignin and cellulose biosynthesis in stone cells of ‘Dangshansuli’ fruit and reduces the thickness of SCW in transgenic Arabidopsis plants

The molecular biology and biochemical experiments described above showed that PbrARF13 inhibits the expression of PbrNSC, and we verified this function of PbrARF13 by transiently overexpressing or silencing PbrARF13 in ‘Dangshansuli’ fruit at 35 DAFB (Figure 4a). Transient overexpression of PbrARF13 reduced the content of stone cells (Figure 4b), lignin (Figure 4c) and cellulose (Figure 4d) in the fleshy fruit tissues around the infiltration sites. In addition, overexpression of PbrARF13 decreased the expression levels of stone cell‐formation related genes as shown by qRT‐PCR (Figure S6a,b). In contrast, silencing of PbrARF13 in fruit through the virus‐induced gene silencing (VIGS) technique produces the opposite phenotype (Figures 4b–d and S6c,d). These results show that PbrARF13 acts as a negative regulator to inhibit lignin and cellulose biosynthesis in the stone cells of the ‘Dangshansuli’ fruit.

Figure 4.

Functional validation and genetic analysis of PbrARF13. ‘Dangshansuli’ fruit at 35 days after full bloom (DAFB) was used for injection of Agrobacterium cells containing PbrARF13 construct. (a) Schematic diagram of Agrobacterium injections containing different vectors. 35S::PbrARF13 overexpression vector mediated by 35S strong promoter (35S::GFP empty vector as control) and virus‐mediated VIGS‐PbrARF13 silencing vector (pTRV2‐PbrARF13 and pTRV1 co‐injected, pTRV2 and pTRV1 co‐injected as control), respectively. (b‐d) After 7 d (d, days after injection), fruit samples similar to those shown in (a) were used to analyse the content of the stone cells (b) (n ≥ 20 biological replicates), lignin (c) and cellulose (d) in the fleshy tissue around the infiltration sites (n ≥ 10 biological replicates). Statistical significance was determined by a one‐tailed paired t‐test. *P < 0.05. (e–g) Paraffin‐stained sections of stems of 8‐week‐old Col‐0 and transgenic Arabidopsis plants overexpressing PbrARF13 were observed using fluorescence microscopy at 4X and 40X fields of view. (e) Toluidine blue staining, (f) UV‐excited lignin autofluorescence and (g) Congo red staining. (h–j) Determination of secondary cell‐wall‐related physiological indicators in 8‐week‐old Col‐0 and PbrARF13‐overexpressing Arabidopsis. (h) Interfascicular fibres (IF) thickness (n ≥ 100 count repeats), lignin (i), and cellulose (j) content in stem plants (n ≥ 3 biological replicates). Statistical significance was determined by a one‐tailed paired t‐test. *P < 0.05.

Three transgenic Arabidopsis plant lines overexpressing 35S::PbrARF13‐GFP (OE‐1#, 6# and 10#) were generated via Agrobacterium‐mediated transformation. These transgenic plants were slightly higher than WT plants (Figure S6e–g), and the expression levels of genes responsible for lignin and cellulose biosynthesis in SCW were also significantly downregulated in their stems (Figure S6h). Paraffin‐embedded thin sections of 8‐week‐old WT and transgenic plants were stained with toluidine blue O to examine changes in the SCWs. It was clear that 35S::PbrARF13‐GFP expression reduced the SCW thickness in the interfascicular fibres (IF) (Figure 4e,h), and decreased the deposition of lignin and cellulose in the cell walls of the IF in the transgenic plants (Figure 4f,g). Moreover, 35S::PbrARF13‐GFP overexpression reduced lignin and cellulose content in the stems of the transgenic Arabidopsis plants at 8 weeks old (Figure 4i,j). Taking all of these results together, PbrARF13 acts as a negative regulator to reduce the thickness of SCWs in the IF by reducing the content of lignin and cellulose.

PbrNSC is upstream of PbrMYB132 in the co‐expression network regulating stone cells

Lignin and cellulose are the main components of stone cells. In previous studies, the mechanism of PbrNSC regulation of lignin biosynthesis was characterized, but how it regulates cellulose is not clear (Figures S2 and S7). To identify the downstream targets of PbrNSC, we constructed the ‘PbrNSC‐TF’ regulatory module (Figure 5a‐I) using co‐expression networks of stone cell‐formation genes that were established using RNA‐seq data from developing fruit of 206 pear cultivars (Table S2). In the ‘PbrNSC‐TF’ regulatory module (Figure 5a‐I,b), PbrMYB132 was shown to be the strongest target of PbrNSC. Also, published transcriptome databases on the dynamics of fruit development in different pear varieties visually confirmed the high correlation between PbrNSC and PbrMYB132 (Figure S8a). Further analysis showed that PbrMYB132 is an R2‐R3 MYB TF that localizes to the nucleus (Figure S8b,c,e). In addition, the relative expression of PbrMYB132 was correlated with stone cell content in the different pear cultivars (Figure S8d). Those suggest that PbrMYB132 may be involved in lignin and cellulose biosynthesis in stone cells, functioning as a positive regulator.

Figure 5.

Identification of PbrMYB132 as a key factor of stone cell cellulose biosynthesis. (a) Co‐expression network related to stone cell formation genes was constructed using RNA‐Seq data of developing fruit of 206 pear cultivars. (I) represents transcription factor (TF) genes co‐expressed with PbrNSC; (II) represents stone cell formation‐related genes co‐expressed with PbrMYB132. Gene nodes are indicated by green dots; key genes are highlighted with different colours, such as yellow for PbrNSC and PbrMYB132, red for PbrMYB169, and orange for cellulose synthase genes. The width of the line between the dots represents the weight value between them. The red line has the largest weight value, while the weight values decrease by rotating clockwise along the red line. (b, c) The weight values in (a, I) and (a, II) are further clearly displayed in (b) and (c), respectively. (d and e) Relative expression levels of PbrMYB132, PbrCESA8b, PbrCESA4a, PbrCESA4b and PbrCESA7a in PbrNSC overexpressed (d) or silenced (e) fruit fleshy tissue were analysed using qRT‐PCR and the reference gene PbrGAPDH. Statistical significance was determined using a one‐tailed paired t‐test. *P < 0.05. Values are mean ± SD of three technical replicates (three biological replicates have similar expression trends, we selected one for presentation).

Interestingly, PbrMYB132 expression was highly correlated with PbrCESA8b/4a/4b/7a in the co‐expression network of PbrMYB132 and downstream structural genes (Figure 5a‐II,c; Table S3). This may imply that PbrMYB132 may be involved in the biosynthesis of cellulose in stone cells. Furthermore, the expression levels of PbrMYB132 and the PbrCESA genes were increased or decreased when PbrNSC was overexpressed or silenced in the ‘Dangshansuli’ fruit (Figure 5d,e). Therefore, we suggest that PbrMYB132 may be involved in the regulation of stone cell cellulose deposition as a target of PbrNSC.

PbrNSC directly regulates the expression of PbrMYB132 by binding to the SNBE in the promoter

To investigate how PbrNSC regulates PbrMYB132 expression, a SCW NAC binding element (SNBE) was found between −1237 and −1255 bp upstream of the PbrMYB132 start codon (Figures 6a and S13b). The activities of three PbrMYB132 promoter fragments (−1 to −1000 bp, −1 to −1500 bp and −1 to −2000 bp) fused to a reporter gene were tested together with the 35S::PbrNSC‐GFP effector construct in N. benthamiana leaves (Figure 6a). PbrNSC activated the two promoter fragments containing the SNBE but not a fragment without the SNBE based on the activities of two reporter genes, LUC (Figure 6b) and GUS (Figure 6c). The binding specificity of PbrNSC to the SNBE was subsequently confirmed by EMSA (Figure 6d) and Y1H (Figure 6e) assays.

Figure 6.

PbrNSC promotes PbrMYB132 expression by directly binding secondary cell wall (SCW) NAC binding element (SNBE) element in the PbrMYB132 promoter. (a) Schematic diagrams show effector and reporter vectors used for dual luciferase and GUS activity assays. (b and c) The activation effect of PbrNSC on three PbrMYB132 promoter fragments was tested using dual luciferase (b) and GUS (c) assay. Statistical significance was determined by a one‐tailed paired t‐test. *P < 0.05. (d) Electrophoretic mobility shift assay (EMSA) showed PbrNSC directly binding to the SNBE (−1237–−1255) on the PbrMYB132 promoter. Biotin‐P and Mut‐P represent biotin‐labelled WT and mutation probes, respectively; competitor‐P represents the unlabelled WT sequence. The black triangle represents the rates of competitors (10×, 50× and 100×). (e) Yeast one‐hybrid (Y1H) assay showed that PbrNSC binds the promoter of PbrMYB132. Yeast cells were co‐transformed with a decoy vector (containing a PbrMYB132 promoter fragment fused to the HIS2 reporter gene) and a prey vector (containing PbrNSC fused to the GAL4 activation domain). 10 mM, 40 mM and 60 mM 3‐AT (3‐amino‐1,2,4‐triazole) were used to inhibit background growth and test the strength of the interaction. (f) Expression analysis of PbrMYB132 in ‘Dangshansuli’ fruit at different developmental stages under 0 μm or 200 μm NAA treatment. The expression level of each gene was standardized against the reference gene (PbrGAPDH). Statistical significance was determined using a one‐tailed paired t‐test. *P < 0.05. (g) Heatmap shows 200 μm NAA treatment decreasing PbrMYB132 RPKM value at 35 days after full bloom (DAFB). (h) PbrMYB132 expression level in PbrARF13 overexpressed or silenced fruit fleshy tissues was analysed using qRT‐PCR and reference gene PbrGAPDH. Statistical significance was determined using a one‐tailed paired t‐test. *P < 0.05. Values are mean ± SD of three technical replicates (three biological replicates have similar expression trends, we selected one for presentation).

In addition, the PbrMYB132 expression level was reduced by NAA treatment as shown by qRT‐PCR (Figure 6f) and RNA‐seq data analyses (Figure 6g). The PbrMYB132 expression levels were also reduced and increased by overexpression and silencing PbrARF13 in pear fruit, respectively (Figure 6h). This suggests that PbrMYB132 is involved in the NAA‐PbrARF13‐mediated inhibition of lignin and cellulose biosynthesis in the stone cells.

PbrMYB132 promotes SCW formation by activating cellulose and lignin biosynthesis genes

Transient overexpression PbrMYB132 increased the numbers of stone cells and the lignin and cellulose content in fruit tissues around the infiltration sites. PbrMYB132 silencing produced opposite results (Figures 7a–d and S9a,b). In two stable Arabidopsis transgenic lines (OE‐2# and 12#) (Figure S9c,d,e), 35S::PbrMYB132‐GFP overexpression increased the thickness of the SCWs in IF (Figure 7e,h). While the expression levels of genes associated with SCW formation were significantly upregulated in the stems of the transgenic plants (Figure S9f). Moreover, the transgenic Arabidopsis lines at 8 weeks old contained more lignin and cellulose (Figure 7f,g,i,j). These results strongly suggest that PbrMYB132 acts as a positive regulator of SCW formation in stone cells of pear.

Figure 7.

Functional validation and genetic analysis of PbrMYB132. ‘Dangshansuli’ fruit of 35 days after full bloom (DAFB) was used for Agrobacterium‐mediated transient injection of PbrMYB132. (a) Schematic diagram of Agrobacterium injections containing different vectors. 35S::PbrMYB132 overexpression vector mediated by 35S strong promoter (35S::GFP empty vector as control) and virus‐mediated VIGS‐MYB132 silencing vector (pTRV2‐PbrMYB132 and pTRV1 co‐injected, pTRV2 and pTRV1 co‐injected as control), respectively. (b‐d) After 7 d, fruit samples similar to those shown in (a) were used to analyse the content of the stone cells (b) (n ≥ 20 biological replicates), lignin (c) and cellulose (d) in the fleshy tissue around the infiltration sites (n ≥ 10 biological replicates). Statistical significance was determined by a one‐tailed paired t‐test. *P < 0.05. (e–g) Paraffin‐stained sections of stems of 8‐week‐old Col‐0 and transgenic Arabidopsis plants overexpressing PbrMYB132 were observed using fluorescence microscopy at 4X and 40X fields of view. (e) Toluidine blue staining, (f) UV‐excited lignin autofluorescence and (g) Congo red staining. (h‐j) Determination of secondary cell‐wall‐related physiological indicators in 8‐week‐old Col‐0 and PbrARF13‐overexpressing Arabidopsis. (h) Interfascicular fibres (IF) thickness (n ≥ 100 count repeats), lignin (i), and cellulose (j) content in stem plants (n ≥ 3 biological replicates). Statistical significance was determined by a one‐tailed paired t‐test. *P < 0.05.

To understand how PbrMYB132 promotes SCW formation, we analysed the relationship between PbrMYB132 and stone cell cellulose biosynthesis genes PbrCESA4a/4b/7a/7b/7c/8a/8b in pear (Figure S10). The expression levels of these genes were positively correlated with the expression level of PbrMYB132 as shown by qRT‐PCR (Figure 8a). PbrMYB132 could activate the promoters of PbrCESA4b/7a/8a, with the strongest activation on PbrCESA8a, as shown by a dual luciferase assay (Figure 8b,c). Subsequent EMSA evidence suggests that PbrMYB132 binds directly to the ‘SMRE‐I’ element of the PbrCESA8a promoter (Figures 8d and S13c).

Figure 8.

PbrMYB132 promotes PbrCESA8a expression through binding to the SMRE element in the PbrCESA8a promoter. (a) Expression levels of PbrCESA4a, PbrCESA4b, PbrCESA7a, PbrCESA7b, PbrCESA7c, PbrCESA8a and PbrCESA8b, important to stone cell cellulose biosynthesis, were measured in PbrMYB132 overexpressed or silenced fruit fleshy tissues using qRT‐PCR. Statistical significance was determined by a one‐tailed paired t‐test. *P < 0.05. (b) The activating effects of PbrNSC or PbrMYB132 on the promoters of the genes shown in (a) were examined using dual luciferase (The promoter sequence of PbrCESA4a could not be cloned successfully). Statistical significance was determined by a one‐tailed paired t‐test. *P < 0.05. (c) The promoter of PbrCESA8a was divided into two parts (−1–−1000 and −1–−2000) according to the distribution of SMRE positions on the PbrCESA8a promoter, and the activation effect of PbrMYB132 on each part was detected separately using dual luciferase. Statistical significance was determined by a one‐tailed paired t‐test. *P < 0.05. (d) Electrophoretic mobility shift assay (EMSA) analysis confirmed that PbrMYB132‐HIS binds to the SMRE‐I element in the PbrCESA8a promoter. Biotin‐P and Mut‐P represent biotin‐labelled WT and mutation probes, respectively; competitor‐P represents the unlabelled WT Sequence. The black triangle represents the rates of competitors (10×, 50× and 100×). Values are mean ± SD of three technical replicates (three biological replicates have similar expression trends, we selected one for presentation).

Because Pbr4CL4, PbrLAC4 and PbrLAC5 are involved in lignin biosynthesis in stone cells (Figures 1f,g and S2) (Wang et al., 2021), we were interested in determining whether PbrMYB132 can regulate the expression of these genes. As shown in Figure 9a, the expression of the Pbr4CL4, PbrLAC4 and PbrLAC5 genes was enhanced by PbrMYB132 overexpression, but it was reduced when PbrMYB132 was silenced. In a dual luciferase assay experiment, PbrMYB132 activated the PbrLAC5 promoter but not the Pbr4CL4 or PbrLAC4 promoters. We also found that PbrNSC activated the Pbr4CL4 and PbrLAC4 promoters, but not the PbrLAC5 promoter (Figure 9b,c).

Figure 9.

PbrMYB132 promotes lignin biosynthesis through binding to SMRE‐III/IV in the PbrLAC5 promoter. (a) Expression levels of Pbr4CL4, PbrLAC4 and PbrLAC5 have been measured in fruit fleshy tissues with PbrMYB132 overexpressed or silenced after infiltration using qRT‐PCR. Statistical significance was determined by a one‐tailed paired t‐test. *P < 0.05. (b) Schematic diagram of the construction of effector and reporter vectors for dual luciferase assays. (c) The activation effect of PbrNSC or PbrMYB132 on the promoters of Pbr4CL4, PbrLAC4 and PbrLAC5 was detected by dual luciferase. Statistical significance was determined by a one‐tailed paired t‐test. *P < 0.05. (d) According to the distribution of SMRE on the PbrLAC5 promoter, the promoter of PbrLAC5 was divided into three parts (−1–−1000, −1–−1500, and −1–−2000), and the activation effect of PbrMYB132 on each part was detected by dual luciferase assay. Statistical significance was determined by a one‐tailed paired t‐test. *P < 0.05. (e) Electrophoretic mobility shift assay (EMSA) showed that PbrMYB132 binds directly to the SMRE within −1–−1500 bp of the PbrLAC5 promoter. Hot‐P stands for biotin‐labelled probes with SMRE fragments. (f and g) The binding ability of PbrMYB132‐His to SMRE‐III/IV was detected by EMSA. The results showed PbrMYB132 directly binds the SMRE‐III (f) and SMRE‐IV (g) elements of the PbrLAC5 promoter. Hot‐P and Mut‐P represent biotin‐labelled WT and mutation probes, respectively; competitor‐P represents the unlabelled WT sequence. The black triangle represents the rates of competitors (10×, 50× and 100×). Values are mean ± SD of three technical replicates (three biological replicates have similar expression trends, we selected one for presentation).

In additional experiments, PbrMYB132 activated two PbrLAC5 promoter fragments from −1 to −1000 bp and −1 to −1500 bp (Figure 9d). Four SMRE elements (SMRE‐I, II, III and IV) were identified in the region between −1 and −1500 bp upstream of PbrLAC5 (Figure S13d). EMSA showed that PbrMYB132 binding to the SMRE‐III and IV sequences was specific as shown in a further experiment using unlabelled SMRE‐III and IV probes and mutated probe sequences (Figure 9e–g). Taken together, the results of these experiments suggest that PbrMYB132 binds directly to SMRE‐III and IV in the PbrLAC5 promoter to activate the expression of PbrLAC5.

Discussion

Exogenous NAA application inhibits stone cell formation in pear fruit

Plant hormones have a wide range of applications in fruit tree production. For example, spraying with GA can reduce the effects of grape diseases and promote unisexual fruiting for the production of seedless grapes (Acheampong et al., 2017); exogenous application of ABA accelerates the translocation of nitrate from apple roots to the shoots (Liu et al., 2021); BR treatment inhibits ethylene production and delays pear fruit ripening (Ji et al., 2021); and jasmonic acid activates expression of the TF MdMYC2 to promote ethylene biosynthesis during apple fruit ripening (Li et al., 2017). Auxin (NAA) is commonly used as a flower‐ and fruit‐thinning agent in pear cultivation (Bangerth, 2000). In our study, exogenous sprays of 100–300 μm NAA reduced the content of stone cells, with 200 μm NAA having a significant promotion effect on fruit weight (Figure 1a‐e). This may be related to the nature of the auxin, after all, the effect of the auxin is different and relies on the tissue, developmental stages and the auxin concentration (Zhao, 2010, 2018). More importantly, treatment with 200 μm NAA is suitable for the main pear varieties as a useful agronomic measure to reduce the content of stone cells (Figure S1), although it may require further optimization when it is used in large‐scale fruit production systems.

The ‘PbrARF13‐PbrNSC’ cascade regulates SCW formation in stone cells in response to NAA signalling

NAA treatment reduces the expression level of PbrNSC, the lignin and cellulose biosynthesis ‘switch’ in stone cells (Figure 1f,g). Therefore, the identification of genes that act upstream of PbrNSC became a key goal of this study, and we hypothesized that these upstream genes are induced by NAA, as the ARFs are known to regulate SCW formation (Lee et al., 2019; Preston et al., 2004; Tang et al., 2020). In Arabidopsis, auxin regulates endothecium lignification via the ‘AtARF8.4‐AtMYB26’ pathway (Ghelli et al., 2018). AtARF17 regulates endothecium lignification through direct activation of the AtMYB108 during anther dehiscence (Xu et al., 2019b). In poplar, the ‘PtoARF5‐PtoHB7/8’ signalling cascade mediates auxin‐triggered xylem cell differentiation during early wood development (Xu et al., 2019a); and PtrARF2.1, a homologue of AtARF2, affects lignin biosynthesis (Fu et al., 2019). In rice, OsARF6/17 controls the flag leaf angle by regulating SCW biosynthesis in lamina joints (Huang et al., 2021a). In our study, the exogenous NAA treatment‐induced ‘PbrARF13‐PbrNSC’ signalling cascade was found to inhibit SCW formation in stone cells (Figures 2, 3, 4).

Also, the NAA‐induced expression of PbrARF13 is of interest to us. In general, auxin does not directly increase the expression levels of ARFs that are related to the transduction pathway of auxin (Rutten et al., 2022). Under low auxin concentrations, ARFs normally bind to the repressor protein AUX/IAAs. Conversely, at higher auxin concentrations, the ARF/AUX/IAA complex can segregate, allowing ARFs to play their role in auxin signalling (Chapman and Estelle, 2009). In this process, ARF proteins rely on the PB1 structural domain (A large majority of ARF proteins contain three conserved domains: DNA‐binding domain (DBD), middle region (MR), and Phox and Bem 1 (PB1)) to form heterodimers with Aux/IAA proteins (Szemenyei et al., 2008). However, PbrARF13 contains only the DBD and MR and lacks the PB1 domain (Figure S5b), which suggests that PbrARF13 may not interact directly with Aux/IAA proteins, meaning that the response of PbrARF13 to NAA may be independent of the classical model of auxin signalling transduction. In our study, 200 μm NAA treatment significantly increased the expression level of PbrARF13 at 28 DAFB (7 days after NAA spraying), but did not affect its expression at 35 DAFB (14 days after NAA spraying). Such results suggest that NAA treatment may induce transcription of PbrARF13 and that such induction has a temporal effect. Similarly, Arabidopsis AtARF3 also lacks the PB1 domain but its expression is induced by auxin, peaks at 4 h of auxin treatment and then gradually decreases (Cheng et al., 2013; Zhang et al., 2018). Furthermore, the expression of AtARF3 is inhibited by auxin inhibitors yucasin and NPA (Zhang et al., 2022b).

Based on the above information, we generally consider that the application of exogenous NAA may have induced the expression of PbrARF13. However, it is not clear how NAA induces the expression of PbrARF13. Based on the data shown in Figure 1a,c that 200 μm NAA treatment promoted fruit enlargement, the effect of NAA on cell division and differentiation may indirectly affect the expression of PbrARF13, and the induced PbrARF13 is subsequently involved in the regulation of lignin and cellulose biosynthesis in stone cells through the ‘PbrARF13‐PbrNSC‐PbrMYB132’ cascade reaction. In conclusion, it is equally interesting to understand how NAA induces the expression of PbrARF13.

Other than that, we are more interested in the effect of endogenous auxin on stone cell formation. With the help of previous studies of the trends in endogenous auxin content during the dynamics of fruit development in ‘Dangshansuli’ pear (Chen, 2019), we showed that endogenous auxin concentration gradually decreased during the period of rapid stone cells formation (10–30 DAFB). From 30 to 50 DAFB, when the rate of stone cell formation is relatively slow, the concentration of endogenous auxin gradually increased. This also suggests that the endogenous auxin concentration may be negatively correlated with stone cell formation.

The ‘PbrARF13‐PbrNSC‐PbrMYB132’ cascade is involved in the regulation of lignin and cellulose

Lignin and cellulose are the main components of stone cells (Zhang et al., 2021b) and SCW (Hofte and Voxeur, 2017; Meents et al., 2018). Lignin biosynthesis is mainly dependent on the regulation of the phenylalanine metabolic pathway by the ‘NAC‐MYB’ TF cascade (Du and Groover, 2010; Humphreys and Chapple, 2002; Kubo et al., 2005; Zhong et al., 2006). Our previous study also clarified that ‘PbrNSC‐PbrMYB169’ promotes lignin biosynthesis in the stone cells of the pear (Wang et al., 2021).

Cellulose biosynthesis is relatively straightforward and depends mainly on the catalytic action of cellulose synthase on polysaccharide substrates (Endler and Persson, 2011; McFarlane et al., 2014). However, the understanding of the regulatory network of cellulose biosynthesis in stone cells is limited. NACs often regulate SCW cellulose synthase genes through a regulatory cascade. For example, the ‘OsNAC29/31‐OsMYB61‐OsCESA’ (Huang et al., 2015) and ‘GhFSN1‐GhMYB7‐GhCESA4’ (Huang et al., 2021b) cascades rely on MYB TFs as ‘intermediate factors’ to help NAC proteins regulate cellulose biosynthesis. In this study, we identified PbrMYB132 as a potential ‘intermediate factor’ linking PbrNSC and PbrCESA genes by analysing a co‐expression network of stone cell formation‐related genes. Biochemical experiments and functional analysis established that PbrMYB132, a positive regulator of lignin and cellulose biosynthesis in stone cells, participates in the regulation of cellulose biosynthesis in stone cells through the reactions of the ‘PbrNSC‐PbrMY132‐PbrCESA’ cascade (Figures 5, 6, 7, 8). In addition, NAA treatment reduced the expression level of PbrMYB132 (Figure 6f,g,h), suggesting that the inhibitory effect of NAA on cellulose may be achieved through the ‘PbrARF13‐PbrNSC‐PbrMYB132‐PbrCESAs’ cascade.

Functional analyses revealed that PbrMYB132 not only promotes cellulose biosynthesis, but also lignin biosynthesis (Figure 7). This implies that PbrMYB132 plays a dual role in the regulation of lignin and cellulose biosynthesis in stone cells. In SCW regulation, AtMYB103 acts as a regulator of cellulose biosynthesis and also binds to the promoter of FH5 to promote lignin biosynthesis (Ohman et al., 2013). PtoMYB6 not only promotes flavonoid accumulation in poplar, but also interacts with PtoKNAT7 to inhibit lignin biosynthesis (Wang et al., 2019). However, genes and molecular mechanisms that can regulate both lignin and cellulose biosynthesis have not been reported in stone cell studies. Our subsequent molecular biology experiments showed that PbrMYB132 binds to two SMRE sequences in the PbrLAC5 promoter to participate in the regulation of lignin biosynthesis. More interestingly, PbrNSC by itself has no activation effect on PbrLAC5 (Figure 9a); therefore, PbrMYB132, as the target gene of PbrNSC, further completes the PbrNSC regulatory cascade.

In addition, we also validated the dependence of stone cell formation regulation on the ‘PbrNSC‐PbrMYB132’ module (Figure S11). When we transiently expressed PbrNSC and silenced PbrMYB132 in pear fruit (Figure S11a), the stone cell content was reduced compared with fruit in which PbrNSC was overexpressed and PbrMYB132 expression was normal. However, the stone cell content was still significantly higher than in the control (Figure S11b–d). Therefore, we hypothesized that PbrNSC contributes to the formation of stone cells in ways other than via the ‘PbrNSC‐PbrMYB132’ pathway. We also found that NAA treatment reduced the expression level of PbrNSC and PbrMYB169 (Figure 1f,g), suggesting that ‘PbrNSC‐PbrMYB169’ is also activated by NAA and might be involved in the inhibition of lignin biosynthesis.

Of course, as a cascade reaction involving multiple genes, there may be a ‘leapfrog’ regulatory reaction, and we used a dual luciferase assay to show that PbrARF13 does not directly activate the downstream genes PbrMYB169, PbrMYB132, Pbr4CL4, PbrLAC4 and PbrLAC5 (Figure S12a). Also, the results shown in Figure S12b indicated that there appears to be no feedback regulation between PbrARF13 and PbrNSC, PbrMYB169, and PbrMYB132. Therefore, we speculate that the NAA‐mediated PbrARF13‐PbrNSC‐PbrMYB132 cascade reaction is unidirectionally regulated.

The above biochemical experiments and functional verification allowed us to preliminarily determine that the NAA‐mediated ‘PbrARF13‐PbrNSC‐PbrMYB132’ cascade inhibits lignin and cellulose synthesis in stone cells (Figure 10). The formation of stone cells is a complex process involving multiple genes, proteins and cells. It encompasses not only the division and differentiation of pulp cells, but also deposition of lignin and cellulose in the SCW. Although our study has elucidated the role of the NAA‐induced ‘PbrARF13‐PbrNSC‐PbrMYB132’ cascade in regulating lignin and cellulose in stone cells, whether the effect of NAA on pulp cell division and differentiation affects stone cell formation also deserves further exploration.

Figure 10.

Models of the molecular mechanism of exogenous spraying of 200 μm NAA to inhibition of lignin and cellulose biosynthesis in stone cells of pear fruit. Exogenous application of 200 μm NAA promotes the expression of PbrARF13, and a high level of the PbrARF13 protein inhibits PbrNSC expression by binding to the PbrNSC promoter, thereby reducing the stone cell content. In addition, PbrMYB132 as a target gene of PbrNSC promotes cellulose and lignin biosynthesis in stone cells by activating the expression of the cellulose synthase genes PbrCESA4b/7a/8a and the laccase PbrLAC5. In conclusion, our study identified the ‘PbrARF13‐PbrNSC‐PbrMYB132’ regulatory cascade that mediates lignin and cellulose biosynthesis in stone cells in response to auxin signals in pear fruit.

Materials and methods

Plant materials and growth conditions

The pear trees used in this study were grown in an orchard at the Dashahe farm in XuZhou City, China, and were 15 years old. ‘Dangshansuli’ fruit were sprayed with 0, 100, 200 and 300 μm NAA (α‐naphthaleneacetic acid; Aladdin) at 20 and 21 DAFB. The NAA‐treated fruit was collected at 21, 28, 35, 49, 80, 110, 140 and 160 DAFB for testing.

The 35 DAFB ‘Dangshansuli’ fruit were used for transient expression studies to verify the gene functions. Fruit of six varieties (‘Cuiyu’, ‘Qiuyue’, ‘Sucuiyihao’, ‘Hosui’, ‘Huangguan’ and ‘Cuiguan’) were used to verify the effects of spraying with 200 μm NAA.

Arabidopsis Colombia (Col‐0) plants were used for genetic transformation via the floral dip method (Clough and Bent, 1998). Nicotiana benthamiana plants were used for dual luciferase and GUS assays.

Determination of fruit weight and measurement of the stone cells, lignin and cellulose content in fruit flesh

The average weight of 10 NAA‐treated ‘Dangshansuli’ fruit at 28, 35 and 49 DAFB was tested as single fruit weight. Measurement methods for stone cells, lignin and cellulose content in fruit flesh were the same as described previously (Xue et al., 2019a; Zhang et al., 2021b).

Analysis of Phloroglucinol‐HCl staining of lignin

Ten grams of Phloroglucinol solid dye was dissolved in a 100 mL mixed solution (95% alcohol 1:1 50% HCl). The fruit being tested was cut longitudinally and stained with the phloroglucinol‐HCl solution for 7 min, after which the stained areas were rinsed with deionized water to terminate the staining reaction.

Transient gene expression in pear fruit

The coding sequences (CDS) of PbrARF13, PbrNSC and PbrMYB132 were amplified using the high‐fidelity PCR enzyme (Vazyme, China). The PCR products were cloned into the pCAMBIA1300 (35S::GFP) vector to generate fusion protein constructs using the One‐step Rapid Cloning‐Kit. Suspensions of Agrobacterium strain GV3101 cells carrying the 35S::PbrARF13‐GFP, 35S::PbrNSC‐GFP or 35S::PbrMYB132‐GFP plasmids in induction buffer (10 mM MgCl₂; 200 μm acetosyringone; 10 mM MES) were shaken gently for 3 h. The suspensions were then injected into 35 DAFB ‘Dangshansuli’ fruit using a 1 mL syringe, and the injected fruit was placed at room temperature (RT) for 5–7 days. It is necessary to note that due to individual differences in the fruit tested and the efficiency of Agrobacterium injection, we mixed samples between treatments for determination.

Virus‐induced gene silencing

The cDNA fragments of PbrARF13 (1755–2076 bp) or PbrMYB132 (108–484 bp) were cloned into the pTRV2 vector to construct the pTRV2‐PbrARF13 or pTRV2‐PbrMYB132 recombinant plasmids. The plasmids were introduced into Agrobacterium strain GV3101, and suspensions of Agrobacterium cells containing pTRV2‐PbrARF13 or pTRV2‐PbrMYB132 were mixed in equal amounts with cell suspensions of Agrobacterium GV3101 carrying the pTRV1 plasmid. The mixtures of Agrobacterium cell suspensions were injected into 35 DAFB ‘Dangshansuli’ fruit and the injected fruit was placed at RT for 5–7 days.

Measurement of the lignin and cellulose content and histological analysis of Arabidopsis plants

The main stem bases (10 cm in length) from 8‐week‐old Arabidopsis plants were cut into 2 mm slices for the determination of lignin and cellulose content as described previously. Paraffin sections of Arabidopsis stem, toluidine blue and Congo red staining were performed by Service. Toluidine blue‐stained sections were observed under white light. Autofluorescence of lignin was observed under UV light (excitation wavelength of 355/25 nm). Congo red‐stained sections were observed using an excitation wavelength of 525/50 nm and an emission wavelength of 470/40 nm. Observation of the above sections was done using a BX‐51 fluorescence microscope (Olympus, Japan).

Identification of PbrARF genes

The whole genome sequence of ‘Dangshansuli’ pear together with its GFF3 (General Feature Format) annotation file was obtained from http://peargenome.njau.edu.cn/default.asp?d=6&m=1. The conserved AFR domains (PF06507 and PF02309) were downloaded from the Pfam database. Conserved Pfam structural domains were detected by running Hidden Markov Model software with an E‐value <0.05. In addition, the SMART program and NCBI bulk CD search tool were used to identify conserved ARF structural domains in each protein sequence. All 31 PbrARF proteins were finally obtained and named PbrARF_1–31 as shown in Table S1.

Subcellular localization, protein structural domains and construction of phylogenetic trees

Leaves of 4‐week‐old Nicotiana benthamiana plants were infiltrated with Agrobacterium containing the 35S‐PbrARF13‐GFP or 35S‐MYB132‐GFP constructs and placed in the dark for 12 h, after which they were grown under normal conditions for 48 h. The location of the GFP signals in the N. benthamiana leaf cells was observed using a laser confocal microscope (Zeiss LSM880 Airyscan, Germany).

SMART and CD searches were used to analyse the conserved amino acid structural domains of PbrARF13 and PbrMYB132. Neighbour‐joining phylogenetic trees of PbrMYB132 and other related MYB protein sequences were constructed using MEGA7 software with 1000 bootstrap replicates. The annotation and review of the phylogenetic trees were carried out using iTOL.

Quantitative real‐time polymerase chain reaction

Extraction and reverse transcription of total RNA extracted from the material were carried out according to the manufacturer's instructions (TransGen, China). qRT‐PCR was performed using a CFX384 Touch Real‐Time PCR Detection System (Bio‐Rad, USA), using the PbrGAPDH or AtActin gene as the internal control. Relative expression levels of each gene were calculated using the 2^−ΔCt method. Meanwhile, dynamic fit curve analysis of PbrNSC and PbrARF13 based on RT‐qPCR was performed with the aid of Excel (2003).

RNA‐seq analysis

We performed RNA‐seq analysis on ‘Dangshansuli’ fruit that had been treated with 0 μm and 200 μm NAA at 28 and 35 DAFB, and three groups of samples were randomly selected as the biological replicates for each treatment (each replicate contained 3–5 fruit) for the extraction of total RNA. Total RNA (20 μg) from each sample was used for the construction of sequencing libraries that were then sequenced by Novogene (Beijing, China) using the HiSeq X (Illumina, San Diego, CA) platform. The sequence reads were then mapped onto the ‘Dangshansuli’ reference genome using StringTie. We used TBtools software to min‐max (0–1) normalization RPKM values (reads per kilobase of exon per million reads mapped) for the genes involved in this study. All genes RPKM treated with 0 μm and 200 μm NAA at 28 and 35 DAFB as shown in Table S5.

Dual luciferase and GUS activity analysis

The coding sequence of the TF as an effector was cloned into the pCAMBIA1300 (35S::GFP) vector and transformed into Agrobacterium strain GV3101. Promoter fragments of different lengths as a reporter were cloned into the pGreenII 0800‐LUC (LUC‐0800) vector and transformed into Agrobacterium strain GV3101‐pSoup. Cultures of the two Agrobacterium strains were induced by gentle shaking in a buffered solution for 3 h, mixed in a 9:1 (effector:reporter) ratio and infiltrated into the leaves of 4‐week‐old N. benthamiana plants. The infiltrated tobacco plants were placed in the dark for 12 h and then grown under normal conditions for 48 h, after which the firefly luciferase (LUC) and Renilla luciferase (REN) activities were assayed using dual‐luciferase assay reagents (Promega, USA). Determination of fluorescence values was performed using a microplate reader.

(PerkinElmer‐VICTOR‐X4, USA). GUS activity analysis as described previously (Yu et al., 2021).

Electromobility shift assay

The CDS of PbrARF13, PbrNSC and PbrMYB132 were cloned into the pET‐32a (+) vector to generate three His fusion protein constructs. Purification of the PbrARF13, PbrNSC and PbrMYB132 fusion proteins was performed using HIS Sefinose resin (Sangon Biotech, China). Mutated probes and unlabelled probes were used for competition in the EMSA experiments. Gene promoter sequences containing putative cis‐element binding sites were labelled at the 5′ end using a biotin DNA labelling kit (Beyotime Biotechnology, China) as directed by the manufacturer. EMSA was performed using a chemiluminescent EMSA kit (Beyotime Biotechnology, China).

Yeast one‐hybrid assays

Promoter sequences containing core cis‐elements were inserted into the pHis2 vector. The recombinant vectors were transferred into yeast strain Y187 cells using the LiAC‐PEG4000 method. The TF‐encoding genes were amplified and inserted into the pGADT7 vector, and the resulting constructs were then transformed into the pHis2 yeast strain containing the promoter plasmid. The yeast transformants were further grown on SD/−Leu/−Trp medium. Finally, positive yeast clones were diluted and cultured in SD/−Leu/−Trp/‐His medium containing 20 mM 3‐AT (3‐amino‐1,2,4‐triazole) (OD₆₀₀: 1, 10⁻¹, 10⁻² and 10⁻³) or SD/−Leu/−Trp/‐His medium with different concentrations of 3‐AT added. The relative growth rates of the transformed yeast strains were observed after 2–3 days.

Construction of a co‐expression network for lignin and cellulose

The weight values between the stone cell formation‐related genes in the co‐expression network were downloaded from https://genomebiology.biomedcentral.com/articles/10.1186/s13059‐021‐02531‐8 (Wang et al., 2021). TF weight values centred on PbrNSC and structural gene weight values centred on PbrMYB132 are given in Tables S2 and S3. All gene number codes involved in this study are shown in the supporting data.

Accession numbers

The gene accession numbers involved in this study are shown in the following. PbrARF13, Pbr016201.1; PbrNSC, Pbr038584.1; PbrMYB132, Pbr038701.2; PbrMYB169, Pbr012624.1; PbrLAC4, Pbr035962.1; PbrLAC5, Pbr012358.1; PbrLAC18, Pbr003857.1; Pbr4CL1, Pbr024635.1; Pbr4CL4, Pbr013445.1; PbrCCR1, Pbr022402.1; PbrCAD1, Pbr026287.1; PbrHCT2, Pbr022425.1; PbrCCOMT2, Pbr034039.1; PbrC3H1, Pbr020890.1; PbrCESA4a, Pbr011043.1; PbrCESA4b, Pbr032894.2; PbrCESA7a, Pbr007376.1; PbrCESA7b, Pbr034218.1; PbrCESA7c, Pbr034219.1; PbrCESA8a, Pbr000518.1; PbrCESA8b, Pbr016107.1. The primer sequences used to construct the different vectors for each gene are shown in Table S4.

Conflict of interest

All authors have no conflict of interest to declare.

Author contributions

Jun Wu designed the study; Shaozhuo Xu performed the experiments; Manyi Sun and Xiuxia Liu carried out data analyses; Yongsong Xue, Guangyan Yang, Rongxiang Zhu, Weitao Jiang and Runze Wang collected experimental materials; Shaozhuo Xu wrote the manuscript; Jia‐long Yao, Zhiquan Mao, Cheng Xue and Jun Wu revised the manuscript. All the authors read and approved the final manuscript.

Statistical analysis

Statistical analyses were performed using one‐way ANOVA with IBM SPSS Statistics 19 software. Differences were tested for statistical significance using a one‐sided paired t‐test with GraphPad Prism 8 software.

Supporting information

Figure S1 Exogenous spraying of 200 μm NAA promoted fruit development and reduced stone cells content in six major Chinese pear cultivars.

Figure S2 The pattern of molecular mechanisms by which PbrNSC and PbrMYB169 regulate the lignin and cellulose biosynthesis in stone cells.

Figure S3 Identification of the auxin response factor PbrARF _1–31 in the ‘Dangshansuli’ genome.

Figure S4 PbrNSC and PbrARF13 were involved in NAA inhibition of lignin and cellulose biosynthesis in stone cells with a negative correlation.

Figure S5 Basic properties of PbrARF13.

Figure S6 Biological function and genetic analysis of PbrARF13.

Figure S7 Molecular mechanisms of PbrNSC regulation of lignin and cellulose biosynthesis in stone cells.

Figure S8 Expression patterns of PbrMYB132 in different pear varieties and basic characteristics of PbrMYB132.

Figure S9 Biological function and genetic analysis of PbrMYB132.

Figure S10 Identification of cellulose synthases (CESA) associated with cellulose biosynthesis in the stone cells.

Figure S11 PbrNSC regulation of stone cell formation is not completely dependent on PbrMYB132.

Figure S12 Validation of the regulatory relationship of the ‘PbrARF13‐PbrNSC‐PbrMYB132’ cascade.

Figure S13 Promoter sequences of PbrNSC, PbrMYB132, PbrCESA8a and PbrLAC5.

Table S1 Primer sequences of PbrARFs for the construction of dual luciferase effector vectors.

Table S2 Transcription factors are co‐expressed with PbrNSC and may be involved in lignin and cellulose biosynthesis in stone cells.

Table S3 Structural genes that are co‐expressed with PbrMYB132 may be involved in lignin and cellulose biosynthesis in stone cells.

Table S4 Accession numbers and primer sequences of the genes studied in this study.

Table S5 Gene RPKM values of ‘Dangshansuli’ fruit at 28 and 35 DAFB with 0 or 200 μm NAA‐treated.