Xiao‐Yuan Tao,Xue‐Ying Guan,Gao‐Jie Hong,Yu‐Qing He,Su‐Juan Li,Shou‐Li Feng,Jian Wang,Guang Chen,Fei Xu,Jia‐Wei Wang,Sheng‐Chun Xu
Biotinylated Tn5 transposase‐mediated CUT &Tag efficiently profiles transcription factor‐DNA interactions in plants Xiao‐Yuan Tao et al.
AtSPL9,B‐CUT&Tag,defence,flowering,OsPHR2,Tn5
Summary In contrast to CUT&Tag approaches for profiling bulk histone modifications, current CUT&Tag methods for analysing specific transcription factor (TF)‐DNA interactions remain technically challenging due to TFs having relatively low abundance. Moreover, an efficient CUT&Tag strategy for plant TFs is not yet available. Here, we first applied biotinylated Tn5 transposase‐mediated CUT&Tag (B‐CUT&Tag) to produce high‐quality libraries for interrogating TF‐DNA interactions. B‐CUT&Tag combines streptavidin‐biotin‐based DNA purification with routine CUT&Tag, optimizing the removal of large amounts of intact chromatin not targeted by specific TFs. The biotinylated chromatin fragments are then purified for construction of deep sequencing libraries or qPCR analysis. We applied B‐CUT&Tag to probe genome‐wide DNA targets of Squamosa promoter‐binding‐like protein 9 (SPL9), a well‐established TF in Arabidopsis; the resulting profiles were efficient and consistent in demonstrating its well‐established target genes in juvenile‐adult transition/flowering, trichome development, flavonoid biosynthesis, wax synthesis and branching. Interestingly, our results indicate functions of AtSPL9 in modulating growth‐defence trade‐offs. In addition, we established a method for applying qPCR after CUT&Tag (B‐CUT&Tag‐qPCR) and successfully validated the binding of SPL9 in Arabidopsis and PHR2 in rice. Our study thus provides a convenient and highly efficient CUT&Tag strategy for profiling TF‐chromatin interactions that is widely applicable to the annotation of cis‐regulatory elements for crop improvement.
Biotinylated Tn5 transposase‐mediated CUT &Tag efficiently profiles transcription factor‐DNA interactions in plants Xiao‐Yuan Tao et al.

In contrast to CUT&Tag approaches for profiling bulk histone modifications, current CUT&Tag methods for analysing specific transcription factor (TF)‐DNA interactions remain technically challenging due to TFs having relatively low abundance. Moreover, an efficient CUT&Tag strategy for plant TFs is not yet available. Here, we first applied biotinylated Tn5 transposase‐mediated CUT&Tag (B‐CUT&Tag) to produce high‐quality libraries for interrogating TF‐DNA interactions. B‐CUT&Tag combines streptavidin‐biotin‐based DNA purification with routine CUT&Tag, optimizing the removal of large amounts of intact chromatin not targeted by specific TFs. The biotinylated chromatin fragments are then purified for construction of deep sequencing libraries or qPCR analysis. We applied B‐CUT&Tag to probe genome‐wide DNA targets of Squamosa promoter‐binding‐like protein 9 (SPL9), a well‐established TF in Arabidopsis; the resulting profiles were efficient and consistent in demonstrating its well‐established target genes in juvenile‐adult transition/flowering, trichome development, flavonoid biosynthesis, wax synthesis and branching. Interestingly, our results indicate functions of AtSPL9 in modulating growth‐defence trade‐offs. In addition, we established a method for applying qPCR after CUT&Tag (B‐CUT&Tag‐qPCR) and successfully validated the binding of SPL9 in Arabidopsis and PHR2 in rice. Our study thus provides a convenient and highly efficient CUT&Tag strategy for profiling TF‐chromatin interactions that is widely applicable to the annotation of cis‐regulatory elements for crop improvement.

The study of protein‐chromatin interactions characterizes the dynamic binding of transcription factors or chromatin architecture factors to cis‐regulatory elements of the genome, which account for the spatiotemporal control of gene expression (Spitz and Furlong, 2012). For decades, the traditional assay for determining the occupancy of individual TFs has been chromatin immunoprecipitation (ChIP) coupled with either deep sequencing (ChIP‐seq) or target‐specific quantitative PCR (ChIP‐qPCR). In these immunoprecipitation‐based methods, a protein of interest is enriched along with its associated nascent DNA fragments (Furey, 2012). However, ChIP experiments suffer from need for high sample input, low signal‐to‐noise ratio and inconsistency of signals (Bartosovic et al., 2021). Cleavage Under Targets and Tagmentation (CUT&Tag) is an alternative strategy for epigenomic profiling, in which the target chromatin protein is bound in situ by the specific antibody, which then tethers the protein A/G‐Tn5 transposase fusion protein. The pA/G‐Tn5 transposase is activated in the presence of Mg2+ and acts to ‘cut‐and‐paste’ the target chromatin, resulting as the fragmented target DNA tagging with adaptor sequences, that is tagmentation reaction (Kaya‐Okur et al., 2019). In CUT&Tag for animal cells, the cell membrane is permeabilized with digitonin (a non‐ionic detergent) to allow the antibodies and pA‐Tn5 to diffuse into the cell. While for plant CUT&Tag, as the presence of cell wall in plant cells cannot be permeabilized effectively by digitonin, the plant intact nuclei need to be isolated for reaction. CUT&Tag‐seq outperforms ChIP in its simplicity, low dependence on devices/kits, low input requirement and high signal‐to‐noise ratio (Kaya‐Okur et al., 2020). While CUT&Tag approaches for profiling histone modifications have seen good success (Bartosovic et al., 2021; Kaya‐Okur et al., 2020; Tao et al., 2020; Wu et al., 2021), analysing specific transcription factor (TF)‐DNA interactions with CUT&Tag remains technically challenging due to the relatively low abundances of TFs. To date, only a few general chromatin architecture factors present in high abundance (CTCF and Rad21) and limited number of TFs (Olig2, Pax6) have been successfully profiled using CUT&Tag in mammalian cells (Bartosovic et al., 2021; Kaya‐Okur et al., 2019; Qiao et al., 2022). An efficient CUT&Tag strategy with successful application to plant TFs is not currently available. In addition, in this enzyme‐tethering method, target chromatin sites are fragmented by the in situ ‘cut‐and‐paste’ action of Tn5 transposase (Reznikoff, 2003), which means that intact chromatin not recognized by the antibody remains present after reaction. Thus, it is not currently possible to apply qPCR after standard CUT&Tag. Here in this work, we provide for the first time a biotinylated Tn5 transposase‐mediated CUT&Tag (B‐CUT&Tag) strategies to produce a high‐quality library for TF‐DNA interactions. We focused on Squamosa promoter‐binding‐like protein 9 (SPL9) in Arabidopsis, which functions in juvenile‐adult transition/flowering (Wang et al., 2009), trichome development (Yu et al., 2010), flavonoid biosynthesis (Gou et al., 2011), wax synthesis (Li et al., 2019), branching (Xie et al., 2020) and gibberellin (GA) and jasmonate (JA) responses (Mao et al., 2017; Zhang et al., 2020). We were able to profile AtSPL9 occupancy in Arabidopsis and to confirm the method's accuracy and consistency by its good recapitulation of the well‐established target genes of AtSPL9, which were previously confirmed by multiple in vitro and in vivo assays including reporter gene activity assays involving mutation of the core ‘GTAC’ motif in promoters (Gou et al., 2011; Yu et al., 2010), EMSA assays (Li et al., 2019) and ChIP‐qPCR (Gou et al., 2011; Yu et al., 2010). We further combined B‐CUT&Tag and mRNA‐seq data to obtain multiple high‐confidence candidate target genes of AtSPL9 with involvement in plant defence responses. Finally, we applied an optimized B‐CUT&Tag‐qPCR procedure to confirm binding of TFs including AtSPL9 and OsPHR2 to their specific targets. Taken together, this study provides a method to effectively study the epigenetic regulation of TFs in plants.

Our B‐CUT&Tag approach builds on biotinylated Tn5 transposase‐mediated tagmentation and biotin‐streptavidin‐based purification. In principle, two Tn5 transposase monomers were complexed with adaptors containing the 19 bp Tn5 mosaic end (ME) transposon sequence (Reznikoff, 2003) and the 14/15 bp 5′ adaptor sequence compatible with Illumina paired‐end sequencing (adaptor AB: adaptor AC at 1:1 ratio, refer to Methods for details) to form the Tn5 transposase homo‐dimer (Xu et al., 2021), in which Primer B in adaptor AB were 5′ biotin‐triethylene glycol spacer (TEG) modified (Table S1). The intact nuclei were isolated and applied for plant B‐CUT&Tag. After antibody binding (including a primary antibody and a secondary antibody, refer to Method part for details), pA/G‐Tn5 tethering and tagmentation reaction, the biotinylated adaptor was incorporated into the target chromatin sites, giving rise to biotinylated chromatin fragments. The total DNA was then extracted and the biotinylated DNA fragments were purified using a biotin‐streptavidin (biotin‐SA) magnetic beads purification system, resulting in the removal of large amounts of intact chromatin not targeted by the selected TF. The resultant biotinylated DNA fragments were used for library construction and deep sequencing (B‐CUT&Tag‐seq) or subjected to quantitative PCR (B‐CUT&Tag‐qPCR) (Figure 1, refer to Methods for details).

TFs display transient, dynamic and unstable properties when bound to their target DNA (Swift and Coruzzi, 2017). Thus, the unfixed/native conditions employed in CUT&Tag for abundant histone markers are not suitable when assaying TFs. To overcome this problem, chromatin cross‐linking is included as a mandatory step to preserve the protein‐DNA interactions. We first asked whether formaldehyde cross‐linked nuclei can be used for CUT&Tag and compared native CUT&Tag and cross‐linked CUT&Tag (X‐CUT&Tag) at profiling H3K27me3 landscape using nuclei isolated from leaves of 30‐day‐old cotton seedlings. X‐CUT&Tag was performed using isolated nuclei pre‐treated with 1% formaldehyde for 5 min. The results showed that X‐CUT&Tag generated H3K27me3 signals consistent with native CUT&Tag (Figure S1). H3K4me3 profiling using the same nuclei from cotton leaves is provided for comparison. From the results, a trend was discovered that chromatin with higher degrees of H3K27me3 modification showed lower H3K4me3 levels (e.g. genes in cluster 1) and vice versa (genes in cluster 2) (Figure S1), indicating that CUT&Tag is applicable not only for detecting activating chromatin markers but also silencing chromatin markers. A lowered FRiP value in H3K27me3 X‐CUT&Tag sample was found compared with native CUT&Tag (Figure S1), which may be resulted from the overcross‐linking of the nuclei. About 75% peaks were shared between X‐CUT&Tag and native CUT&Tag (Figure S1), indicating that the anti‐H3K27me3 antibody and Tn5 transposase can enter the cross‐linked nuclei efficiently. Thus, formaldehyde cross‐linked nuclei are compatible with CUT&Tag.

We first determined whether B‐CUT&Tag could generate consistent chromatin landscapes in comparison to conventional CUT&Tag. CUT&Tag, B‐CUT&Tag without biotin‐SA magnetic bead purification (B‐CUT&Tag Puri.‐), and B‐CUT&Tag (B‐CUT&Tag) for H3K4me3 (a histone marker for active chromatin) were performed using low input of nuclei (equals ~1 μg chromatin for each reaction) isolated from shoots of 21‐day‐old wild‐type (Col‐0 background) Arabidopsis plants (Figure 2a). The bulk of the DNA fragments from above H3K4me3 CUT&Tag, B‐CUT&Tag Puri.‐ and B‐CUT&Tag libraries ranged from 250 to 750 base pairs (bp) after PCR amplification (Figure 2b). Analysis of H3K4me3 signals near protein‐coding genes in three different CUT&Tag samples showed high correlation and consistency (r ≥ 0.99) (Figure 2c). Similar numbers of peaks were called in CUT&Tag, B‐CUT&Tag Puri.‐ and B‐CUT&Tag samples (17 076, 18 632 and 17 309, respectively; Figure 2d). We summarized the number of H3K4me3 peaks and the fractions of reads in peaks (FRiP) obtained using different amounts of clean reads (in millions, M); FRiP values of up to ~0.6 were obtained using 9 m clean reads from three CUT&Tag samples and remained stable as the number of sampled clean reads was increased (Figure 2e). In summary, B‐CUT&Tag generates consistent profiles, including in terms of signal intensity, signal‐to‐noise ratio and number of peaks, compared with conventional CUT&Tag when low input of nuclei was applied for abundant H3K4me3 signals.

To verify whether B‐CUT&Tag can be used to efficiently identify target genes of plant TFs, we conducted genome‐wide profiling of the targets of Squamosa promoter‐binding‐like protein 9 (SPL9) in Arabidopsis. For this, we used pSPL9::3xflag‐rSPL9 plants (Wang et al., 2009) in which the 3xFLAG‐tagged miR‐156‐resistant form (rSPL9) was expressed from its native promoter. Relative to wild type (WT, Columbia‐0), pSPL9::3xflag‐rSPL9 plants exhibited early flowering (Figure 2f). We performed B‐CUT&Tag using nuclei isolated from cross‐linked inflorescences of pSPL9::3xflag‐rSPL9 plants, in which AtSPL9 is highly expressed (Yu et al., 2010). B‐CUT&Tag and routine CUT&Tag assays for AtSPL9 were performed in parallel using high‐input of nuclei (equals ~5 μg chromatin/reaction, refer to Methods part for details). In AtSPL9 B‐CUT&Tag, the intact chromatin not tagmentated by pA‐Tn5 that unbound to SA magnetic beads were removed (Figure S2). The purified biotinylated target fragments were subjected to on‐beads PCR for the library construction, the bulk of the DNA fragments from the AtSPL9 B‐CUT&Tag libraries ranged from 250 to 750 base pairs (bp) after 18 cycles of PCR (Figure 2h), which is the regular product sizes for CUT&Tag assay. In comparison, without biotin‐SA based purification in conventional CUT&Tag for AtSPL9, the large amount of intact chromatin DNA in the PCR reaction affected the PCR efficiency, the resulting DNA fragments from the AtSPL9 CUT&Tag libraries showed a long smear from 250 bp up to more than 5000 bp after 18 cycles of PCR (Figure 2g) and failed to pass the Quality Control (QC) for NGS. These results indicated that B‐CUT&Tag efficiently profiles plant TF‐DNA interactions once high‐input of nuclei was used in the assay.

We compared the procedures for the three‐day protocols for both B‐CUT&Tag and ChIP, B‐CUT&Tag outperforms ChIP as it is less device‐ and kit‐ dependent, it does not need expensive sonication device or the NGS library construction kit (Figure 3a). AtSPL9 B‐CUT&Tag‐seq signals were mainly distributed upstream of transcription start sites (TSSs), that is in promoter regions (Figure 3b). Two technical replicates showed high correlation (Figure 3c, Pearson's r = 0.99), with calling of 9290 and 9188 peaks, respectively (fold enrichment >2, q‐value <0.05). Of those, 6383 peaks (~70%) were overlapping and could be assigned to 4476 protein‐coding genes in the Arabidopsis genome (Figure 3d). We also performed AtSPL9 B‐CUT&Tag using nuclei isolated from shoots of 21‐day‐old pSPL9::3xflag‐rSPL9 plants. In total, 3183 peaks (60%–70%) identified in that assay were shared with inflorescence samples (Figure 3e). In parallel of the AtSPL9 B‐CUT&Tag‐seq, we performed AtSPL9 ChIP‐seq using the same amount of nuclei (equals ~5 μg chromatin/reaction) isolated from 1 g cross‐linked inflorescences of pSPL9::3xflag‐rSPL9 plants, unfortunately, we failed to obtain adequate resultant DNA, lead to low‐quality NGS libraries with low number of peaks (Figures 3d, S3). Alternatively, we provided AtSPL9 ChIP‐seq data using shoots of 14‐day‐old pSPL9::3xflag‐rSPL9 plants, which yielded 5841 peaks corresponding to 4744 protein‐coding genes (fold enrichment >2, q‐value <0.05, data from the Jia‐Wei Wang lab), among which 1713 (36.1%) and 1951 (41.1%) peak‐associated genes overlapped with the target genes revealed by AtSPL9 B‐CUT&Tag‐seq of the inflorescences and shoots of 21‐day‐old plants, respectively (Figure 3e). These data indicate spatial and temporal variation of AtSPL9 regulation during plant growth and development. We further evaluated the efficiency and consistency of B‐CUT&Tag by checking for the presence of multiple well‐established AtSPL9 target genes. This analysis revealed that B‐CUT&Tag extensively covered the well‐documented target genes of AtSPL9 implicated in multiple developmental and physiological pathways, including genes encoding a helix‐turn‐helix TF (LFY) and MADS‐box proteins (AP1, SOC1, FUL and AGL42) in the juvenile‐to‐adult phase transition and flowering (Lian et al., 2021; Smaczniak et al., 2012; Wang et al., 2009), MYB or MYB‐like proteins (TCL1, TRY, CPC and ETC3) in abaxial trichome initiation (Yu et al., 2010), flavanone 3‐hydroxylase (F3H) in flavonoid biosynthesis (Gou et al., 2011), a TCP family TF (BRC1) in shoot branching (Xie et al., 2020) and an alkane‐forming enzyme (CER1) in epidermal wax synthesis (Li et al., 2019) (Figure 3f). Gene ontology (GO) enrichment analysis further revealed that the potential target genes of AtSPL9 were significantly enriched (fold enrichment >2, P value <0.001) in multiple biological processes including meristem initiation, shoot and root development, lipid localization, abiotic/phytohormone stimulus responses and shade avoidance (Figure S4).

mRNA‐seq was performed using inflorescences from WT and pSPL9::3xflag‐rSPL9 plants and revealed 1001 differentially expressed genes (DEGs, fold change >1.5 or <0.67; FDR <0.05). Among them, 233 genes (90 up‐regulated and 143 down‐regulated) overlapped with the list of prospective target genes from B‐CUT&Tag (Figure 4a, Table S3). GO enrichment analysis of the AtSPL9‐targeted DEGs revealed them to be significantly enriched in responses to abiotic stimulus or phytohormones (fold enrichment >2, FDR < 0.05) (Figure 4b), indicating that AtSPL9 regulates pathways partially shared with the response pathways of multiple stimuli. We are interested in the DEGs also represented in B‐CUT&Tag peaks, which are most likely target genes of AtSPL9. Most of them constitute new discoveries as potential targets of AtSPL9, and these genes are broadly involved in growth, development and immunity, secondary metabolism, phytohormone catabolism & homeostasis, phytohormone signalling, abiotic stress responses, shade avoidance, aging and nutrient utilization (Figure 4c, Table S4). It has been reported that DELLA proteins, the key signalling factors in the GA pathway, inhibit AtSPL9 activity through direct interaction at the protein level (Mao et al., 2017). Interestingly, the B‐CUT&Tag results indicated that AtSPL9 binds to the promoter regions of DELLA genes (RGA, GAI and RGL1, 2 and 3) (Figure 3f), and RGL3 was found to be down‐regulated in the inflorescences of pSPL9::3xflag‐rSPL9 plants (Table S4), implying possible feedback between AtSPL9 and DELLA in GA responses (Figure 3f). Figure 4c and Table S4 summarize ten DEGs having mutant or over‐expression lines with documented phenotypes under salt, cold, or drought stress. For example, the salinity‐responsive DWARF AND DELAYED FLOWERING 1 (DDF1) gene was down‐regulated in pSPL9::3xflag‐rSPL9 plants (Figure 4c,d); plants overexpressing this gene exhibit delayed flowering and dwarfism, reduction of GA biosynthesis and increased tolerance to high levels of salt (Magome et al., 2008). Combining the expression profiles of multiple abiotic stress response‐related DEGs (including DDF1, SRO5, ANAC055, AIRP2, MYB2 and COR15B) with their documented phenotypes (Abe et al., 2003; Borsani et al., 2005; Cho et al., 2011; Fu et al., 2018; Magome et al., 2008; Thalhammer and Hincha, 2014) (Table S4) suggests pSPL9::3xflag‐rSPL9 plants to be more sensitive to abiotic stresses, which is consistent with previous reported phenotypes for rSPL9 plants under salt and osmotic stress (Cui et al., 2014).

To determine whether B‐CUT&Tag could likewise be combined with qPCR for validation of the known binding sites, we established the B‐CUT&Tag‐qPCR procedure by validating H3K4me3 profiles for representative sites in the cotton genome. We take advantage of the previous B‐CUT&Tag‐seq data for H3K4me3 modification in cotton plants (Tao et al., 2020) and checked the H3K4me3 levels at specific gene loci as a reference for comparison. We first test the primer pairs that can be used in the qPCR for B‐CUT&Tag samples. Theoretically, there was three different assembly ways of pA‐Tn5 transposome and resulted as four scenarios for adaptor ‘pasting’ after Tn5 tagmentation (Figure S5a). To roughly estimate the proportion of different tagmentation scenarios, we performed Tn5 tagmentation of genomic DNA followed with qPCR to identify the relative content of products from different scenarios, and results showed that the scenario 1 and scenario 2 products accounted for ~99.97% of the tagmentation (Figures 5a, S5b), suggesting that the pA‐Tn5 transposome homo‐dimer harbours both adaptor AB and adaptor AC is the major form after transposome assembly. As an example, we check the H3K4me3 modification levels at GB_D10G1774 gene locus. The fragments from scenario 1 and scenario 2 were quantified using the primer pairs P1/GSPr_GB_D10G1774 and P1/GSPf_GB_D10G1774, respectively (Figure 5a), in which primer P1 harbours the 15 bp 5′ adaptor sequence for Illumina sequencing from adaptor Primer C but not the Tn5 mosaic end (ME) sequence (refer to Table S1 for details), while GSPf_GB_D10G1774 and GSPr_GB_D10G1774 are gene‐specific forward (GSPf) and reverse primers (GSPr) for GB_D10G1774 (Table S1). Unfortunately, we found non‐specific amplification by the single P1 primer, this is more likely resulted from the annealing and extension of single‐strand DNAs (ssDNAs) from the same target site recovered from scenario 1 and scenario 2, respectively (Figure S6a,b). Adding an on‐beads extension step using DNA polymerase before DNA retrieval step significantly reduce non‐specific priming by primer P1 (ΔCt = 5.25) (Figure S6c,d). However, considering the background non‐specific priming by primer P1, qPCR using the primer pairs of P1/GSPf or P1/GSPr is not applied. We then evaluated gene‐specific primer pairs (GSPf/GSPr) in the qPCR reaction. For the gene GB_D10G1774, CUT&Tag‐seq results indicated that H3K4me3 signals were enriched in the gene body and weak in the 3′ downstream region (Figure 5b). B‐CUT&Tag‐qPCR targeted to both the high signal region (Region 1) and the low signal region (Region 2) was performed, consistently, results indicated that the target fragments covering Region 1 having 10.2‐fold enrichment in the H3K4me3 assay group normalized to IgG control, while the target fragments covering Region 2 showed no significant enrichment after normalization (Figure 5c). In an alternative normalization method, we used each DNA sample from before biotin‐SA‐based purification as the reference for normalization of qPCR data. With this normalization, 36.60% and 3.59% of respective fragments in the H3K4me3 assay group and IgG control group covered Region 1 after biotin‐SA‐based purification, while 2.45% and 2.34% of fragments covered Region 2 (Figure 5d). Consistent results was also observed at other gene loci, including GB_D03G0642 and GB_A11G13949, as established by qPCR using gene‐specific primer pairs (Figure S7). Thus, B‐CUT&Tag‐qPCR using target‐specific primer pairs is applicable for validation of histone modifications at specific gene loci, and the cycle threshold (Ct) values from both an IgG control and input DNA can be used for normalization.

To determine whether the B‐CUT&Tag‐qPCR procedure we established for assessing histone modification levels was applicable to validating the occupancy of TFs, we employed this method to determine the binding of AtSPL9. Three well‐established target genes of AtSPL9 were selected: TCL1 and TRY, involved in trichome development (Yu et al., 2010), and FUL, involved in flowering control (Wang et al., 2009). The gene‐specific primer pairs used for B‐CUT&Tag‐qPCR were the same with previously documented for ChIP‐qPCR assays (Wang et al., 2009; Yu et al., 2010). The ACT2 gene was used as control. In parallel, we summarized the read counts from B‐CUT&Tag‐seq data that mapped to the qPCR target region (Figure 6a). The ratios of read counts per million mapped reads (RPM) for the qPCR target regions of TCL1, TRY, FUL and ACT2 were 2.45, 2.25, 3.18 and 1.45, respectively, in the rSPL9 assay group (anti‐FLAG antibody) compared with the IgG control (Figure 6b). Consistent with these data, B‐CUT&Tag‐qPCR results showed 7.42‐, 8.58‐, 7.96‐ and 2.21‐fold enrichment for the indicated regions of TCL1, TRY, FUL and ACT2 in the rSPL9 assay group relative to the IgG control (Figure 6c). In comparison, a parallel ChIP‐qPCR results showed 3.23‐, 2.06‐, 3.08‐ and 1.19‐fold enrichment for the indicated regions of TCL1, TRY, FUL and ACT2 in the rSPL9 assay group relative to the IgG control (Figure 6d). These results indicated that B‐CUT&Tag‐qPCR had comparable sensitivity with ChIP‐qPCR. We further applied B‐CUT&Tag‐qPCR to determine the binding of OsPHR2, an important TF that functions in phosphate homeostasis in rice. Three known target genes were selected for analysis: the low‐affinity phosphate (Pi) transporter OsPT2 (Liu et al., 2010), which is the direct target of OsPHR2 and responsible for most of the OsPHR2‐mediated accumulation of excess shoot Pi; the high‐affinity Pi transporter OsPT8, which is also critical for Pi homeostasis (Jia et al., 2011); and the arbuscular mycorrhizae symbiosis‐associated marker gene OsRAM1, which is regulated by an OsPHR2‐centered network (Shi et al., 2021). The B‐CUT&Tag‐qPCR assay was performed using nuclei isolated from cross‐linked shoots of OsPHR2‐Ov1 rice plants (an transgenic plant overexpressing OsPHR2 with FLAG tagging) (Kong et al., 2021). We identified the PHR1 binding site (P1BS; GNATATNC) (Rubio et al., 2001) in the promoters of OsPT2, OsPT8 and OsRAM1 and designed qPCR primers with target regions covering the indicated P1BS cis‐element (Figure 6e). qPCR results showed that, under the Pi‐sufficient (+P) condition, there was no enrichment of target regions for OsPT8 and OsRAM1 in the OsPHR2 assay group (anti‐FLAG antibody) relative to the IgG control group; in contrast, 2.36‐fold enrichment of the target region was observed for OsPT2, which is consistent with the expression pattern of OsPT2 in OsPHR2‐overexpressing rice plants (Liu et al., 2010). Meanwhile, under a Pi‐deficient (−P) condition, 2.56‐, 2.13‐ and 3.47‐fold enrichment was, respectively, observed for the target regions of OsPT2, OsPT8 and OsRAM1 in the OsPHR2 assay group compared with IgG control (Figure 6f). These results indicate different regulation effects of OsPHR2 on Pi homeostasis and mycorrhizal symbiosis under Pi‐sufficient (+P) and Pi‐deficient (−P) conditions.

To summarize, Figure 7 illustrates the comprehensive methodological strategy for B‐CUT&Tag in plants (refer to Methods part for details). Firstly, for cross‐link, the plant tissues were fixed with 1% formaldehyde and quenched with 0.2 m glycine. Secondly, for nuclei isolation, a pilot lysis using a series of Triton X‐100 concentrations is recommended, with 100 mg liquid nitrogen grinded tissue in each 2 mL nuclei lysis buffer. Taking leaf tissue as an example, successful lysis is indicated by accumulation of white/grey nuclei at the bottom and a dark green supernatant after centrifuging at low speed (<500 g for 4 min). The lysate is then filtered through a 500‐mesh stainless steel sieve to collect the nuclei. Thirdly, the B‐CUT&Tag reaction is performed in an optimized volume with respect to the amount of nuclei, antibody, pA/G‐Tn5. Finally, we demonstrated the biotin‐SA magnetic beads‐based DNA purification and the subsequent on‐beads PCR to be feasible for NGS library construction (B‐CUT&Tag‐seq), along with an optimized qPCR procedure for confirming TF‐DNA binding (B‐CUT&Tag‐qPCR).

Beyond the simple affinity of a TF for its corresponding 6–12 bp‐long cis‐regulatory DNA sequence, TF binding and its impact on gene expression are influenced by many different mechanisms, including chromatin accessibility (nucleosome positioning), cofactor proteins, DNA architecture and more (Spitz and Furlong, 2012). Comparisons of TF occupancy from ChIP‐seq data and differentially expressed genes (DEGs) from mRNA‐seq in yeast (Ucar et al., 2009), Drosophila melanogaster (Jakobsen et al., 2007) and mammalian systems (Vokes et al., 2008) have revealed a relatively small overlap between TF binding and gene expression. In our study, B‐CUT&Tag‐seq data revealed totally 4475 potential AtSPL9‐target genes, while RNA‐seq revealed totally 1001 DEGs, in which 233 genes are overlapping, accounting for 23% (233/1001) of the DEGs. This overlap rate is comparable to previous values reported in mammalian systems and plants (10–25%) (Dong et al., 2019; Vokes et al., 2008; Zhang et al., 2017). AtSPL9 has been revealed to positively regulate gene expression by directly binding to a promoter (Wang et al., 2009) and to negatively regulate gene expression (e.g. of anthocyanin biosynthesis genes F3’H, DFR, ANS and UGT78D2) through indirectly destabilizing the MYB‐bHLH‐WD40 activator complex (Gou et al., 2011). We identified 90 up‐regulated DEGs and 143 down‐regulated DEGs as also having B‐CUT&Tag peaks (Figure 5a), which were significantly enriched for the terms response to abiotic stimulus and response to phytohormone stimulus (Figure 5b). Although further experimental confirmation of these TF‐gene interactions remains needed, our data provide new clues for a role of AtSPL9 in coordinating the growth‐defence trade‐offs in plants. Briefly, plants with high AtSPL9 activity adjust their growth/development and exhibit early flowering and accelerated senescence. In this way, plants prioritize responses that improve their capacity to complete their life cycle quickly (i.e. stress escape) while becoming more susceptible to abiotic stress (Cui et al., 2014; Gou et al., 2011), insect herbivory (Mao et al., 2017) or pathogens.

We have developed B‐CUT&Tag‐seq and B‐CUT&Tag‐qPCR procedures that are likely to be widely applied in profiling plant TF‐DNA interactions. CUT&Tag provides outstanding performance with low chromatin input even at the single‐cell level (Bartosovic et al., 2021; Ouyang et al., 2021); however, for most scenarios involving plant tissues such as leaves, stems, roots, inflorescences, fruits and seeds, the amount of input sample does not appear to be the limiting factor. We have employed CUT&Tag assays in plants for years and identified the most likely limiting factors for its successful application as the isolation of intact nuclei and the abundance of target TFs. For the isolation of intact nuclei, we provided a detailed methodology in which a pilot assay that determining the appropriate concentration of Triton X‐100 was introduced, and also a 500‐mesh sieve was used to filter the nuclei after lysis (Figure 7, refer to method part for details). Our B‐CUT&Tag procedure was designed for profiling the occupancy of plant TFs using quantities of input nuclei equivalent to or less than those needed for ChIP; we further combined biotinylated Tn5 transposase‐mediated tagmentation with biotin‐SA‐based purification, thus optimizing the purification of DNA fragments that specifically resulting from Tn5 tagmentation. As a result, large amounts of intact chromatin not targeted by specific proteins were excluded (Figure S2), which minimized the negative effects of such chromatin in downstream NGS library construction and sequencing. In principle, solid phase reversible immobilization (SPRI) methodology‐based DNA size selection by purification beads (e.g. AMPure XP) might be applicable to remove larger intact chromatin DNA and recover desired size DNA fragments after traditional CUT&Tag. However, the efficiency, sensitivity and specificity between SPRI‐based selection and biotin‐SA based purification in B‐ CUT&Tag need to be compared in further experiment. Our results indicated that biotin‐SA based purification methodology was easy to perform and efficiently purified target DNA for the following NGS library construction (Figure 2h). Thus, our B‐CUT&Tag procedure is recommended when traditional CUT&Tag procedure failed to obtain qualified libraries, which might be especially suitable for TFs with lower abundance or with lower number of target sites that need higher input of nuclei in the reaction. Importantly, for the TFs that with a relatively lower abundance or lower number of target sites, multiple technical replicates of reactions are recommended; the resultant DNA from multiple technical replicates can be combined to undergo biotin‐SA‐based purification. Thus, B‐CUT&Tag enables flexibility in the amount of input nuclei depending on the abundance of target TFs. Notably, B‐CUT&Tag still outperformed traditional ChIP in terms of its low dependence on devices/kits (Figure 3a) and high consistency between biological replicates (Figure 3c). Our B‐CUT&Tag‐qPCR procedure is applicable for profiling binding of TFs in both monocot (rice) and dicot (cotton and Arabisopsis) plants for the known target sites.

Since its invention, CUT&Tag has provided an alternative method for profiling protein‐DNA interactions in both animals and plants. CUT&Tag for bulk histone modification has been successfully applied to investigate the epigenomic landscapes of monocot rice and dicot rapeseed and cotton (Ouyang et al., 2021; Tao et al., 2020). In addition to the bulk histone modification states that reflect chromatin accessibility, annotation of biologically significant cis‐regulatory elements relating to yield, quality and stress tolerance is of great importance in crop production. We provide for the first time high‐quality plant TF occupancy profile data obtained using CUT&Tag‐seq and also establish a CUT&Tag‐qPCR procedure with which to examine binding of TFs. Our results suggest that this CUT&Tag strategy can be widely applied in plants. Moreover, as it is very difficult and costly to systematically produce reliable antibodies for the specific recognition of individual plant proteins (Lu et al., 2020), when generating the overexpressing transgenic plants, we fused the open reading frame (ORF) of the TF with FLAG tag (DYKDDDDK) and utilized a commercial anti‐FLAG antibody to recognize the target tag‐fused protein. Thus, this B‐CUT&Tag method using an antibody against specific protein tag has great promise for applications examining the binding of chromatin proteins and annotating cis‐regulatory TF‐binding elements for crop improvement.

Arabidopsis wild type (WT) and pSPL9::3xflag‐rSPL9 lines were grown in a growth chamber at 22 °C with a 16‐h light/8‐h dark photoperiod. The 21‐day‐old pSPL9::3xflag‐rSPL9 plants or inflorescences from 28‐day‐old pSPL9::3xflag‐rSPL9 plants were used in B‐CUT&Tag. Upland cotton (Gossypium hirsutum) was grown in a growth chamber at 28 °C with a 16‐h light/8‐h dark photoperiod. Leaves from 30‐day‐old seedlings were used in cross‐linking CUT&Tag (X‐CUT&Tag). Rice (Oryza sativa L. cv. Nipponbare) OsPHR2‐Ov1 plants (a transgenic plant overexpressing OsPHR2 with FLAG tagging) (Kong et al., 2021) were grown in a growth chamber with a 14‐h light (30 °C)/8‐h dark (25 °C) photoperiod and thermoperiod. Seven‐day‐old rice seedlings initially grown on Pi‐sufficient medium were subjected to P‐sufficient (+P, 200 μm KH2PO4) and P‐deficient (−P, 0 μm KH2PO4) conditions. In the Pi‐deficient medium, phosphate buffer was replaced by equimolar amounts of KCl (Kong et al., 2021). Three days later, rice shoots were harvested for B‐CUT&Tag.

Routine CUT&Tag for profiling histone H3K4me3 or H3K27me3 was performed as described (Tao et al., 2020). Due to the abundance of histone H3K4me3 and H3K27me3 modification, small input of nuclei (equals ~1 μg chromatin) was used in each reaction, and only the primary antibodies including normal mouse IgG (12–371; Millpore, Burlington, MA, USA) for control, anti‐H3K4me3 antibody (ab8580; Abcam, Cambridge, Cambridgeshire, UK) and anti‐ H3K27me3 antibody (ab6002; Abcam) were used.

The B‐CUT&Tag protocol was developed from a previous method with modifications (Tao et al., 2020). All buffers used were the same as described (Tao et al., 2020). Sequences of primers A, B and C for generating adaptor AB and adaptor AC are provided in Table S1. Primer B was 5′‐biotin‐TEG modified. Primer A, B and C was diluted in annealing buffer (10 mm Tris pH 8.0, 50 mm NaCl, 1 mm EDTA) to make a concentration of 100 μm. In two PCR tubes, set up the following reactions by mixing 10 μL primer A with 10 μL primer B (tube 1 for adaptor AB); and 10 μL primer A with 10 μL primers C (tube 2 for adaptor AC). The adaptors were generated using the program in the PCR machine (heat lid, 75°C for 15 min, 60 °C for 10 min, 50 °C for 10 min, 40 °C for 10 min, 25 °C for 30 min). The adaptor AB and adaptor AC were mixed at 1:1 ratio, designated as ‘adaptor mix’ (50 μm). Five microlitre of commercially available pA‐Tn5 (7.5 pmol/μL) for example Vazyme, or Abclonal was complexed with 0.875 μL adaptor mix and 3.5 μL of coupling buffer at 37°C for 60 min to assemble the Tn5 transposase. For B‐CUT&Tag for TF, cross‐linking was first performed, in which each 1 gram of tissue was fixed using 20 mL fixation buffer (10 mm Tris pH 8.0, 10 mm KCl, 0.5 mm spermidine, 1 mm EDTA, β‐Mercaptoethanol, 1% formaldehyde) for 10 min under vacuum; the cross‐linking was subsequently stopped by adding 2 mL of 2 m glycine. The cross‐linked tissue were washed using 50 mL ddH2O for three times. After washing, the tissue were transferred to paper towels to absorb excess water. For nuclei isolation, (optional) a pilot assay was performed to determine the appropriate concentration of Triton X‐100 in the lysis buffer: Half a gram cross‐linked tissue was ground into fine power in liquid nitrogen, then transferred to 50 mL falcon tubes. Ten milliliter of ice‐cold nuclei lysis buffer A (10 mm Tris pH 8.0, 10 mm KCl, 0.5 mm spermidine, 1 mm EDTA, β‐Mercaptoethanol) was added and the material resuspended evenly. The suspension (~10 mL) was transferred to five 2‐mL tubes followed by centrifugation at 400 g at 4°C for 5 min. The supernatant was then decanted and the pellet (~100 mg/tube) was resuspended by 1 mL nuclear isolation buffer B (10 mm Tris pH 8.0, 10 mm KCl, 0.5 mm spermidine, 1 mm EDTA, X% Triton X‐100, 0.1% protease inhibitor cocktail) with 0.1%, 0.25%, 0.5%, 1% and 2% Triton X‐100, respectively, for each tube. Take plant leaves as an example, the appropriate concentration of Triton X‐100 for lysis is indicated by accumulation of white/grey nuclei at the bottom and a dark green supernatant after centrifugation (400 g for 5 min). For cross‐linked inflorescences or shoots of Arabidopsis for AtSPL9 assay, and cross‐linked shoot of seven‐day‐old rice seedlings for OsPHR2 assay in this study, the appropriate concentration of Triton X‐100 is 0.5%. One gram cross‐linked tissue was ground into fine power in liquid nitrogen, then transferred to 50 mL falcon tubes. Twenty milliliter of ice‐cold nuclei lysis buffer A was added and the material resuspended evenly, followed by centrifugation at 400 g at 4 °C for 5 min. The supernatant was then decanted and the pellet was resuspended by adding 10 mL nuclei lysis buffer B containing 0.5% Triton X‐100, followed by incubation on ice for 5–10 min. The resulting lysate (~10 mL) was transferred to five 2‐mL tubes and the pellet was collected by centrifugation for 4 min at 400 g , 4 °C. The supernatant was then decanted and the pellet was resuspended by adding 10 mL nuclei wash buffer (10 mm Tris pH 8.0, 150 mm NaCl, 0.5 mm spermidine, 0.1% protease inhibitor cocktail). The nuclei suspension was filtered through an autoclaved 500‐mesh sieve to remove larger tissue debris. The nuclei were collected in 2‐mL tubes by centrifugation for 4 min at 400 g , 4°C and all the nuclei was combined into a fresh 1.5 mL tube and washed three times using 1 mL nuclei wash buffer. We estimated nuclei yield based on the pellet volume produced by low‐speed centrifugation (400 g ). For one gram cross‐linked tissue, ~100 μL nuclei in volume was finally obtained. In setting up for the antibody reaction, 100 μL nuclei in volume (equals ~20 μg chromatin) was resuspended in 1 mL antibody buffer (50 mm Tris pH 8.0, 1 mm EDTA, 150 mm NaCl, 0.5 mm spermidine, 1 mg/mL BSA, 0.1% protease inhibitor cocktail, 0.05% w/v digitonin). Each reaction consisted of 250 μL above nuclei suspension. We set up two replicates each for normal mouse IgG (12–371; Millpore) and anti‐FLAG antibody (ab125243; Abcam). Two microlitre (2 μg) antibody was added to 250 μL nuclei suspension and incubated with gentle rotation (12 rpm) at 4°C overnight. After the antibody reaction, the nuclei were collected by centrifugation for 4 min at 400 g , 4°C and washed gently for 5 min using 800 μL IP wash buffer (10 mm Tris pH 8.0, 150 mm NaCl, 0.5 mm spermidine, 0.1% protease inhibitor cocktail, 0.05% v/v digitonin). One μg of rabbit anti‐mouse IgG H&L (HRP) (ab6728; Abcam) in 250 μL IP wash buffer was added to each reaction and incubated with gentle rotation (12 rpm) at room temperature for one hour. Afterwards, the nuclei were collected by centrifugation for 4 min at 400 g , 4°C and washed three times using 800 μL IP wash buffer to remove the unbound secondary antibody. For transposase incubation, 2 μL of biotinylated pA‐Tn5 (8 pmol, commercial source, e.g Vazyme or Abclonal) in 250 μL transposase incubation buffer (10 mm Tris pH 8.0, 300 mm NaCl, 0.5 mm spermidine, 0.1% protease inhibitor cocktail, 0.05% w/v digitonin) was added to each reaction and incubated with gentle rotation (12 rpm) at room temperature for 2 h. Subsequently, the products were the nuclei were collected by centrifugation for 4 min at 400 g , 4 °C and washed three times using 800 μL IP wash buffer to remove unbound pA‐Tn5. For tagmentation, 300 μL tagmentation buffer was added (10 mm Tris pH 8.0, 300 mm NaCl, 0.5 mm spermidine, 0.1% protease inhibitor cocktail, 10 mm MgCl2, 0.05% w/v digitonin) and incubated at 37 °C for one hour. For DNA extraction, 30 μL 10% SDS and 10 μL 0.5 m EDTA were added to stop the tagmentation reaction, and the tubes were incubated at 65 °C overnight for reverse cross‐linking. The next day, 300 μL DNA extraction buffer was added to each tube and DNA was extracted as described (Paterson et al., 1993). The resulting DNA was dissolved in 26 μL ddH2O and take 1 μL for measuring concentration by NanoDrop. For biotin‐SA purification of biotinylated fragments, a total of 10–20 μL SA magnetic beads (10 mg/mL, e.g. Sera‐Mag Streptavidin‐Blocked Magnetic SpeedBeads, Cytiva) were washed twice with 500 μL 1× Western blot (WB) buffer (5 mm Tris–HCl, pH7.5, 0.5 mm EDTA, 1 m NaCl, 0.02% Tween‐20), once with 500 μL 2× WB buffer and finally resuspended in 50 μL of 2× WB buffer (25 μL each for a two‐step purification). We performed a two‐step purification for each sample: The DNA sample (25 μL in ddH2O) was first mixed with 25 μL SA magnetic beads in 2× WB buffer in a PCR tube, and incubated at 37 °C for 20 min in the PCR machine without heat lid. After incubation, place the tubes in a magnetic stand for 5 min to collect the beads. Take a second PCR tube containing the supernatant from first PCR tube (50 μL) and the remaining beads (collected on a magnetic stand before the supernatant from the first PCR tube was added) and incubate once more at 37 °C for 20 min. The resulting beads from two tubes were combined together and washed twice with 500 μL 1× WB buffer, once with 500 μL 10 mm Tris buffer (pH 8.0) and finally resuspended in 50 μL ddH2O. Libraries for deep sequencing were constructed by on‐beads PCR, with the PCR reaction (100 μL) set up as follows: 50 μL beads with affinity purified DNA, 20 μL ddH2O, 20 μL of 5× TruePrep Amplify Enzyme Buffer (Vazyme, Nanjing, Jiangsu, China), 4 μL of 10 uM N50X Primer, 4 μL of 10 μm N70X Primer and 2 μL of TruePrep Amplify Enzyme (Vazyme). The adaptor pooling guide strategy from Illumina was followed. Each PCR solution was mixed gently and 100 μL mineral oil added on top to prevent evaporation. PCR was performed as described (Tao et al., 2020), starting with a low cycle number and adding 2 cycles as needed until visible PCR bands were obtained on agarose gel analysis. After PCR, the SA beads were removed and PCR products purified using commercial DNA purification beads (e.g. AMPure XP beads). Finally, the library was redissolved in 25 μL ddH2O for quality control (QC) and deep sequencing. For B‐CUT&Tag‐qPCR, we performed the B‐CUT&Tag procedure as described above, except that after DNA extraction, the DNA was redissolved in 30 μL ddH2O, from which 5 μL was taken out for use as ‘input’ in a subsequent qPCR assay. The remaining 25 μL was subjected to biotin‐SA purification of biotinylated fragments as described above. The DNA‐bound SA beads were finally resuspended in 50 μL 30 mm NaOAc pH 9.0 and 95% formamide, then incubated at 90°C for 10 min to release the DNA. A magnetic stand was again used to collect the supernatant (~50 μL). qPCR reactions were performed using 1 μL purified DNA and 1 μL input DNA as templates and performed in triplicate. The gene‐specific qPCR primers were designed according to the same criteria as used with ChIP‐qPCR, which was designed to cover ~250 bp sequences flanking the putative TF‐binding sites. The relative proportion of target content was calculated using the ∆Ct method, taking the Ct values of the IgG sample as control. That is, the Ct values of the B‐CUT&Tag antibody sample were normalized to the Ct values of the IgG control, ∆Ct = CtAb – CtIgG. Fold enrichment of the target region in each antibody sample was then determined relative to the IgG control: Fold enrichment = 2−∆Ct. Alternatively, the Ct values of each CUT&Tag sample, whether antibody or IgG, can be normalized using its corresponding input as control: ∆Ct = Ctsample – Ctinput. The percentage of input for each B‐CUT&Tag sample is then calculated as: Percent input = 2−∆Ct × 100%.

ChIP was performed as described (Haring et al., 2007) with modifications. Briefly, cross‐linking and nuclei isolation were performed as described in ‘B‐CUT&Tag‐seq & B‐CUT&Tag‐qPCR’ part above. For AtSPL9 ChIP‐seq assay using the same amount of nuclei as that for AtSPL9 B‐CUT&Tag‐seq, 100 μL nuclei (equals ~20 μg chromatin) isolated from 1 g cross‐linked tissue was resuspended in 320 μL ice‐cold nuclei lysis buffer (50 mm Tris pH 8.0, 10 mm EDTA, 0.4% w/v SDS, 0.1% protease inhibitor cocktail), chromatin sonication was then performed with M220 Focused‐ultrasonicator (Covaris) using the following setups: Peak Power 75.0, Duty Factor 20.0, Cycles/Brust 200, Avg. Power 15.0, for a total of 10 min. After sonication, the clear supernatant, which contains sonicated chromatin was collected after centrifuge at 12 000 g at 4°C for 10 min and was adjusted to 420 μL using ice‐cold nuclei lysis buffer. A 400 μL aliquot of sonicated chromatin was frozen in liquid nitrogen and stored at −80°C for further use after checking the sonication efficiency. To check the efficiency of sonication, the remaining 20 μL aliquot of sonicated chromatin was mixed with 560 μL TE buffer (10 mm Tris pH 8.0, 1 mm EDTA), 20 μL 5 m NaCl and 20 μL 20% SDS, reverse cross‐linking was performed at 65 °C overnight and the DNA was purified using phenol/chloroform method as described (Tao et al., 2020) and was dissolved in 50 μL ddH2O. Five microlitre of the resulting DNA was analysed by 2% agarose gel electrophoresis, the chromatin was sheared to a length between 200 bp and 600 bp. For immunoprecipitation, the reaction was setup by adding 100 μL of sonicated chromatin, 900 μL ChIP Ab incubation buffer (50 mm Tris pH 8.0, 1 mm EDTA, 0.1% v/v Triton X‐100, 150 mm NaCl, 10 μg/mL BSA), 20 μL Protein A + G magnetic beads (CAT#16–663; Millpore) and 2 μL normal mouse IgG or anti‐FLAG antibody. Set two tubes each for IgG control and anti‐FLAG antibody (equals ~5 μg chromatin/reaction) and incubated the tubes overnight at 4 °C with gentle agitation (Rotator at 12 rpm). After immunoprecipitation, the washing, elution, reverse cross‐linking steps were performed as described (Haring et al., 2007). After reverse cross‐linking, the DNA was purified using phenol/chloroform method as described (Tao et al., 2020) and was dissolved in 30 μL ddH2O. The resulting DNA was used for NGS or qPCR.

A total of 1 μg RNA per sample was used as input material for RNA sample preparations. Sequencing libraries were generated using the NEBNext® Ultra™ II RNA Library Prep Kit for Illumina® (NEB, Ipswich, MA, USA) following the manufacturer's recommendations, and index codes were added to attribute sequences to each sample. Three biological replicates were set up for each inflorescence sample from 28‐day‐old pSPL9::3xflag‐rSPL9 and WT plants.

Bioinformatic analysis of mRNA‐seq data was performed as described (Pertea et al., 2016). Bioinformatic analysis of B‐CUT&Tag‐seq data and ChIP‐seq data were likewise performed as described for CUT&Tag‐seq (Tao et al., 2020). Only uniquely mapped and de‐duplicated reads were used for peak calling by macs2 (q‐value cut‐off 0.05).

All data supporting the findings of this study are available in the article or its Supplementary Information or from the corresponding author upon reasonable request. Source data generated in this study, including the raw data from B‐CUT&Tag, CUT&Tag and mRNA‐seq, have been deposited in the National Center for Biotechnology Information Sequence Read Archive under the accession number PRJNA746662.

The authors have declared that no competing interests exist.

X.T., S.X. and J.W. conceived the project and designed the experiments. X.T. performed the experiments. X.T., S.X., J.W., X.G., G.H., Y.H., J.W., F.X., S.F. and G.C. analysed the data and interpreted the results. X.T., S.X. and J. W. wrote the manuscript.

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