Site-specific phosphorylation of LRP regulates FLC chromatin looping and flowering

Sheng Fan; Yu Zhang; Shaobo Zhu; Zhenfeng Teng; Qiqi Zhang; Yuanyuan Yan; Lisha Shen

doi:10.1126/sciadv.adz0645

. 2026 Jan 1;12(1):eadz0645. doi: 10.1126/sciadv.adz0645

Site-specific phosphorylation of LRP regulates FLC chromatin looping and flowering

Sheng Fan ¹, Yu Zhang ¹, Shaobo Zhu ¹, Zhenfeng Teng ¹, Qiqi Zhang ¹, Yuanyuan Yan ², Lisha Shen ^1,^3,^*

PMCID: PMC12757049 PMID: 41477849

Abstract

Emerging evidence suggests that multiple RNA recognition motif (RRM)–containing proteins are involved in transcriptional regulation, yet the underlying mechanisms remain largely unclear. Here, we show that the previously uncharacterized LATE-FLOWERING RRM-CONTAINING PROTEIN (LRP) prevents 5′-3′ chromatin looping of the key floral repressor gene FLOWERING LOCUS C (FLC) via direct association with its 5′ and 3′ genomic regions. Loss of LRP function leads to increased 5′-3′ gene looping, reduced RNA polymerase II (Pol II) occupancy at the FLC locus, thereby increasing FLC transcription and delaying flowering in Arabidopsis thaliana. Furthermore, site-specific phosphorylation of LRP at serine 17, mediated by the kinase PRP4KA, is essential for LRP protein stability and its function in preventing FLC chromatin looping and Pol II recruitment. Our findings reveal that site-specific phosphorylation of LRP by PRP4KA precludes FLC gene loop formation, constituting a key mechanism for FLC transcriptional repression to ensure timely flowering.

Phosphorylated LRP by PRP4KA blocks chromatin looping at FLC and represses its transcription to promote flowering in Arabidopsis.

INTRODUCTION

The RNA recognition motif (RRM) is one of the most abundant and evolutionarily conserved RNA binding domains in eukaryotic proteins (1–3). RRMs typically mediate interaction with single-stranded RNA, enabling RRM-containing proteins to participate in diverse aspects of RNA metabolism, including splicing, transport, translation, and decay (2, 4–8). Beyond their canonical roles in posttranscriptional regulation, emerging evidence suggests that several RRM-containing proteins are also associated with chromatins and contribute to transcriptional regulation (9, 10). For instance, some RRM-containing proteins are widespread in active chromatin regions in human cell lines (10), while in mouse embryonic stem cells, they constitute part of the chromatin-associated proteome and may contribute to the stabilization of RNA polymerase II (Pol II) engagement at the transcription sites (9). In Arabidopsis thaliana, 196 RRM-containing proteins have been annotated (3), but most of them are yet to be functionally characterized.

Several RRM-containing proteins have recently been shown to regulate the floral transition, a critical developmental transition from vegetative growth to reproductive growth, through modulating the RNA metabolism processes of FLOWERING LOCUS C (FLC) and its homologs through diverse mechanisms in Arabidopsis (1). FLC encodes a potent floral repressor and acts together with SHORT VEGETATIVE PHASE (SVP) to directly inhibit the expression of two floral pathway integrators, FLOWERING LOCUS T (FT) and SUPPRESSOR OF OVEREXPRESSION OF CONSTANS 1 (SOC1) (11–15). One RRM-containing protein, SISTER OF FCA, is proposed to function as a scaffold that anchors on the FLC locus, facilitating interactions with both positive and negative transcriptional regulators to fine-tune Pol II engagement and FLC transcription (16). Another RRM-containing protein, heterogeneous nuclear RNP R-LIKE PROTEIN (HRLP), represses FLC transcription by inhibiting its cotranscriptional splicing that facilitates DNA:RNA hybrid (R-loop) formation near FLC intron I and reduces RNA Pol II recruitment (17). Several other RRM-containing proteins, such as RZ-1B, RZ-1C, SC35, and SC35-LIKE proteins, have also been shown to influence FLC splicing and expression to regulate flowering (18, 19). In addition, the transcription of FLOWERING LOCUS M/MADS AFFECTING FLOWERING 1 (FLM/MAF1), a close homolog of FLC, is activated by the RRM-containing protein UBA2c, which represses trimethylation of histone 3 at lysine 27 (H3K27me3) and promotes Pol II occupancy (20), highlighting the potential roles of RRM-containing proteins in modulating chromatin states. Notably, FLC transcription is subject to extensive chromatin-based regulatory mechanisms (11, 12, 21), including the formation of a chromatin loop that links its 5′- and 3′-ends (22–24). Despite these advances, whether and how RRM-containing proteins influence transcription and chromatin dynamics remains largely unknown.

Given the important roles of RRM-containing proteins, their functions are likely subject to fine-tuned regulation by posttranslational modifications (PTMs), such as phosphorylation, which could affect protein stability, subcellular localization, and protein-protein interactions (1, 25–27). However, the presence and functional relevance of PTMs on RRM-containing proteins remained largely unexplored in plants. In this study, we identify a previously uncharacterized RRM-containing protein, LATE-FLOWERING RRM-CONTAINING PROTEIN (LRP), as a key regulator of chromatin looping at the FLC locus to ensure flowering at the appropriate time. LRP directly binds both the 5′ and 3′ regions of FLC, thereby preventing the formation of 5′-3′ chromatin loop. Loss of LRP function enhances 5′-3′ gene looping and facilitates RNA Pol II occupancy at the FLC locus, thereby increasing FLC transcription and delaying flowering in A. thaliana. Furthermore, phosphorylation of LRP at serine 17 by the kinase PRP4KA is crucial for its protein stability and function in preventing FLC chromatin looping and Pol II recruitment. Our study reveals that inhibition of chromatin looping at FLC by site-specific phosphorylation of LRP is an integral mechanism underlying FLC transcriptional repression to determine the flowering time in Arabidopsis.

RESULTS

LRP promotes flowering in Arabidopsis

LRP (At3g52660) contains one coiled-coil domain and three RRM domains and shares high sequence similarities with its closest homologs across various plant species (Fig. 1A and fig. S1). To investigate the biological function of LRP, we isolated a transferred DNA (T-DNA) insertion mutant, lrp-1 (CS855920), in the Columbia (Col) background from the Arabidopsis Biological Resource Center and generated lrp-2, which contained a 1 bp of thymine insertion via CRISPR-Cas9–mediated gene editing (Fig. 1, A and B). There was no detectable full-length LRP transcript in lrp-1 (Fig. 1C). Both lrp-1 and lrp-2 exhibited significantly delayed flowering compared to wild type under both long days (LDs) and short days (SDs) (Fig. 1, D and E, and fig. S2A), suggesting that LRP promotes flowering in Arabidopsis.

Fig. 1. — (A) Diagram illustrating *LRP* gene structure and protein domains. The top shows the schematic diagram of the T-DNA insertion site in *lrp-1* (*CS855920*), the mutation site in *lrp-2* (a mutant generated via CRISPR-Cas9–mediated genome editing), and the target site of the AmiR in *AmiR-lrp*. Black boxes, exons in coding region; gray boxes, UTRs; black lines, introns. Arrowheads denote the positions of primers used for amplifying *LRP* transcripts shown in (C). The bottom shows the domain organization of LRP protein predicted by the Simple-Modular Architecture Research Tool. CC, coiled-coli domain. (B) Alignment of genomic sequence and sequencing chromatograms of wild-type (WT) and *lrp-2* at the CRISPR-Cas9 target site. The single guide RNA (sgRNA) sequence and protospacers adjacent motif (PAM) are highlighted. A thymine (T) insertion, shown in red, is observed in *lrp-2*. (C) Semiquantitative reverse transcription PCR analysis reveals the absence of *LRP* transcripts in *lrp-1*. *TUB2* was amplified as an internal control. (D) *lrp* mutants exhibit late flowering under LDs. (E) Flowering time analysis of *lrp-1*, *lrp-2*, and *AmiR-lrp* under LDs (top) and SDs (bottom). Error bars, means ± SD; n = 20 [(E) and (F)] and n = 3 (G). Asterisks in (E) to (G) indicate statistically significant differences compared to WT (*P < 0.05 and **P < 0.01, two-tailed paired Student’s t test). (F and G) The degree of late flowering (F) under LDs correlates with the down-regulation of *LRP* expression (G) in independent *AmiR-lrp* transgenic plants. *LRP* expression levels normalized to *TUB2* expression are shown relative to its level in WT plants set as 100%. (H) Flowering time distribution of T1 transgenic plants of *lrp-1 g4HA-LRP* and *lrp-1 gGFP-LRP* under LDs. (I) Selected *lrp-1 g4HA-LRP* or *lrp-1 gGFP-LRP* fully rescues the late flowering time of *lrp-1*.

To further examine the role of LRP in promoting flowering, we created LRP knockdown transgenic plants using artificial microRNA (AmiR) interference (28) specifically targeting the last exon of LRP (Fig. 1A). Among 43 AmiR-lrp independent lines obtained, 38 exhibited varying degrees of late flowering under LDs. We then examined LRP expression levels in six selected lines and found that the extent of late flowering correlated with the degree of LRP down-regulation (Fig. 1, F and G). Notably, the strongest knockdown line, AmiR-lrp#39, which had the lowest LRP expression, showed the latest flowering time (Fig. 1, D to G), indicating that down-regulation of LRP has a dosage-dependent effect on flowering.

To confirm that the late flowering phenotype of lrp-1 is attributed to loss of LRP function, we transformed lrp-1 plants with g4HA-LRP or gGFP-LRP, both constructs containing a 4.3-kb genomic region that included a 2.0-kb 5′ upstream sequence, the entire 2.1-kb coding sequence with introns fused in-frame with a 4HA or green fluorescent protein (GFP) tag immediately after the start codon, and 0.17-kb 3′ untranslated region (3′UTR) (fig. S2B). Most of the T1 lrp-1 g4HA-LRP and lrp-1 gGFP-LRP transgenic plants displayed flowering times comparable to wild type (Fig. 1H), demonstrating that LRP is responsible for the late-flowering phenotype observed in lrp-1. Representative lines of lrp-1 g4HA-LRP and lrp-1 gGFP-LRP that fully rescued the flowering phenotype and contained the transgene at a single locus based on a 3:1 Mendelian segregation ratio were selected for further analysis (Fig. 1I).

LRP functions in the nucleus to promote flowering

Quantitative real-time polymerase chain reaction (qPCR) analysis revealed that LRP was broadly expressed across various tissues, with the highest levels detected in rosette leaves (fig. S2C). Further temporal analysis showed that LRP expression gradually increased in the aerial parts of seedlings from 3 to 11 days after germination (fig. S2D), consistent with its role in promoting flowering. To examine the spatial expression pattern of LRP, we generated gGUS-LRP reporter lines (fig. S2B), and most of the independent transgenic lines exhibited similar GUS staining patterns. In a representative line, strong GUS signals were observed in the leaves and shoot apices of both 3- and 9-day-old seedlings, with a noticeably higher intensity at day 9 (Fig. 2A).

Fig. 2. — (A) GUS staining of a representative *gGUS-LRP* transgenic line shows *LRP* expression in 3-day-old and 9-day-old seedlings. (B) Subcellular localization LRP-GFP in *N. benthamiana* leaf epidermal cells. H2B-RFP, RFP fluorescence of the nuclear reporter (core histone 2B fused to red fluorescent protein); merge, merge of GFP, H2B-RFP, and bright-field images. Scale bar, 10 μm. (C) GFP-LRP localization in the root tip of a 5-day-old *lrp-1 gGFP-LRP* seedling. Merge, merge of GFP and bright field images. Scale bar, 20 μm. (D) Detection of 4HA-LRP proteins in the nuclear and cytosol fractions of *lrp-1 g4HA-LRP #2*, *lrp-1 gNLS-4HA-LRP*, and *lrp-1 gNES-4HA-LRP*. Nuclear and cytosol proteins were extracted from 9-day-old *lrp-1 g4HA-LRP #2*, *lrp-1 gNLS-4HA-LRP*, and *lrp-1 gNES-4HA-LRP.* Anti–histone 3 (H3; Abcam, catalog no. ab1791, RRID:AB_302613) antibody was used as a loading control for nuclear fractions, while the Rubisco large subunit (RbcL) stained with Ponceau S served as a loading control for cytosol fractions. T, total protein; N, nuclear protein; C, cytosol protein. (E) Flowering time distribution of T1 transgenic plants of *lrp-1 gNLS-4HA-LRP* and *lrp-1 gNES-4HA-LRP.*

To understand how LRP affects flowering in response to various flowering signals, we analyzed its expression in various flowering mutants and under different environmental conditions. LRP expression was not greatly changed in photoperiod pathway mutants, the gibberellin (GA)–deficient mutant ga1, or ga1 mutants treated with GA (fig. S3, A to C). Vernalization suppressed the late-flowering phenotype of lrp-1 under LDs (fig. S3, D and E). These results indicate that LRP is not involved in the photoperiod, GA, or vernalization pathway. Given that lrp mutants exhibited late flowering under both LDs and SDs (Fig. 1E) and that this delayed flowering was rescued by vernalization (fig. S3, D and E), LRP likely acts within the autonomous pathway, and its expression was slightly affected in several known autonomous pathway mutants (fig. S3F). Consistently, similar to other mutants in the autonomous pathway (29), lrp-1 exhibited a reduced flowering response to low ambient temperature (fig. S3G).

To examine the subcellular localization of LRP, we first transiently expressed 35S:LRP-GFP in Nicotiana benthamiana leaf epidermal cells and observed that LRP-GFP localized to both the cytoplasm and nucleus (Fig. 2B). Consistently, GFP-LRP was detected in both cytoplasm and nucleus in the root tips of lrp-1 gGFP-LRP, which expressed a functional GFP-LRP protein (Figs. 1I and 2C). To confirm this distribution, we examined LRP protein levels in nuclear and cytosolic fractions from 9-day-old lrp-1 g4HA-LRP seedlings and detected 4HA-LRP in both the nucleus and cytoplasm (Fig. 2D). To determine which subcellular fraction of LRP is functionally important for flowering, we generated gNLS-4HA-LRP and gNES-4HA-LRP constructs, in which a nuclear localization signal (NLS) or a nuclear export signal (NES) was fused immediately after the start codon of g4HA-LRP, respectively, and introduced into the lrp-1 mutant background. Most T1 lrp-1 gNLS-4HA-LRP lines, but not lrp-1 gNES-4HA-LRP lines, rescued the late-flowering phenotype of lrp-1 (Fig. 2E). Further, immunoblot analysis confirmed that gNLS-4HA-LRP and gNES-4HA-LRP were exclusively expressed in the nucleus and cytoplasm, respectively (Fig. 2D). These data collectively suggest that LRP acts in the nucleus to promote flowering.

LRP represses nascent FLC transcription

To investigate how LRP influences flowering time, we performed high-throughput RNA sequencing (RNA-seq) on RNA extracted from 9-day-old lrp-1 and wild-type seedlings to identify potential downstream targets of LRP. A total of 535 differentially expressed genes (DEGs; fold change > 1.5 and P < 0.05) were identified in lrp-1 compared to wild-type, with 291 genes up-regulated and 244 genes down-regulated (Fig. 3A, fig. S4A, and tables S1 and S2). Notably, the expression of FLC, a key floral repressor, was significantly up-regulated in lrp-1 (Fig. 3A and table S1). To confirm this, we examined temporal FLC expression in developing seedlings and observed a marked increase in FLC expression levels in lrp-1 compared to wild-type seedlings (Fig. 3B). This up-regulation was fully restored in three independent transgenic lines of lrp-1 g4HA-LRP (Fig. 3C and fig. S2, E and F). Given that FLC directly represses FT and SOC1 expression (14), the expression levels of these two genes were correspondingly reduced in lrp-1 (fig. S4, B and C). In addition, the expression of two FLC homologs, MAF4 and MAF5, was elevated in lrp-1, whereas no consistent changes were observed for MAF1-3 and SVP (fig. S4, D to I).

The flc-3 (30) mutant completely suppressed the late-flowering phenotype of lrp-1 (Fig. 3, D and E), indicating that the delayed flowering of lrp-1 is primarily due to increased FLC levels. Consistently, the reduced expression of FT and SOC1 in lrp-1 was restored in the flc-3 lrp-1 double mutants (fig. S4J). These results suggest that LRP regulates flowering mainly though FLC.

Given that FLC is a major target of LRP in the regulation of flowering, and that its mature RNA is significantly elevated in lrp-1 mutants (Fig. 3, A to E), we further explored whether this increase is due to changes in FLC transcription. To address this, we examined the abundance of nascent FLC transcripts. The levels of nascent FLC RNA were significantly higher in lrp-1 compared to wild-type plants (Fig. 3, F and G), suggesting that LRP negatively regulates FLC transcription.

LRP binds to FLC chromatin and prevents chromatin looping

Although LRP encodes an RNA binding protein with three RRM domains, it does not directly associate with FLC or its antisense transcript COOLAIR (fig. S4K) (31). Since increasing evidence suggests that RNA binding proteins also directly regulate transcription (1, 20), we performed chromatin immunoprecipitation (ChIP) followed by qPCR (ChIP-qPCR) using the lrp-1 g4HA-LRP line to examine whether LRP directly affects FLC transcription. We found that LRP-4HA was associated with the FLC genomic locus (Fig. 3, F and H). Notably, LRP-4HA was primarily enriched at the 5′ region near FLC transcription start site, with additional enrichment at the 3′ region near the translation termination site (Fig. 3H), suggesting that LRP associates with FLC chromatin to directly regulate its transcription.

The 5′ and 3′ regions of the FLC locus are known to form a chromatin loop that facilitates FLC activation (21–24). The association of LRP with both 5′ and 3′ regions of FLC to regulate its transcription prompted us to examine whether LRP affects its 5′-3′ gene looping. Chromosome conformation capture (3C) followed by qPCR revealed that the frequency of the 5′-3′ gene looping at the FLC locus was notably increased in lrp-1 mutants compared to wild type (Fig. 3, F and I). To explore the effect of this gene looping on FLC transcription, we replaced the 3′ genomic region downstream of FLC transcription termination site (TTS) with either a ~0.6-kb sequence corresponding to the 3′ genomic sequence downstream of MAF2 (a homolog of FLC) TTS or a ~0.3-kb NOPALINE SYNTHASE (NOS) terminator (fig. S5A) (23). In contrast to FLC^WT-T, gene looping was absent in FLC^MAF2-T and FLC^NOS-T (fig. S5B). Notably, the remarkable up-regulation of FLC expression observed in FLC^WT-T in the flc-3 lrp-1 background, compared to flc-3, was largely abolished in FLC^MAF2-T and FLC^NOS-T ( fig. S5, C and D). These results suggest that this gene looping is at least partially required for increased FLC expression observed in lrp-1 mutants.

Promoter-terminator gene looping has been suggested to facilitate PoI II local recycling and subsequent transcription reinitiation (32–35). To assess how this enhanced gene looping at FLC influences its transcription, we compared Pol II occupancy at the FLC locus in lrp-1 mutants versus wild-type plants through ChIP assay using an antibody recognizing the C-terminal domain (CTD) of Pol II (anti–Pol II CTD). RNA Pol II enrichment was increased across the FLC locus in lrp-1 mutants (Fig. 3J). Thus, our results suggest that LRP binds at the 5′ and 3′ regions of FLC, inhibits its 5′-3′ chromatin loop formation, and restricts Pol II occupancy to repress FLC transcription (Fig. 3K).

LRP interacts with PRP4KA

Since LRP plays a dosage-dependent role in flowering time regulation (Fig. 1, F and G), we envisaged that its expression or protein activity might be precisely regulated to ensure timely flowering. To identify potential regulators of LRP, we conducted a yeast two-hybrid screening using LRP as a bait against several candidate proteins, including PRP4KA, SET DOMAIN GROUP 8, CURLY LEAF, JUMONJI DOMAIN-CONTAINING PROTEIN 18, EMBRYONIC FLOWER 1 (EMF1), EMF2, RING1A, RING1B, BMI1B, BMI1C, FVE, FLOWERING LOCUS D, FRIGIDA (FRI), FLOWERING LOCUS KH DOMAIN, FLOWERING CONTROL LOCUS A, FPA, and LUMINIDEPENDENS (fig. S6), and isolated PRP4KA (At3g25840), a putative spliceosomal protein kinase (36), which showed interaction with LRP in yeast (Fig. 4A). We then proceed to determine whether PRP4KA is a bona fide interactor of LRP. First, a bimolecular fluorescence complementation (BiFC) assay revealed reconstituted enhanced yellow fluorescent protein (EYFP) fluorescence in the nuclei of N. benthamiana epidermal cells coexpressing nEYFP-LRP and cEYFP-PRP4KA (Fig. 4B), confirming the direct interaction of LRP and PRP4KA in the nucleus in planta. Second, an in vitro glutathione S-transferase (GST) pull-down assay revealed that His-PRP4KA binds to GST-LRP, but not GST alone (Fig. 4C), implying a direct physical interaction. Third, we generated the gPRP4KA-3FLAG transgenic line in the prp4ka-11 mutant background (Fig. 4D) to validate the interaction in vivo. Coimmunoprecipitation (CoIP) assay using protein extracts from 9-day-old F1 seedling of the crosses between lrp-1 g4HA-LRP and prp4ka-11 gPRP4-3FLAG confirmed the in vivo interaction of LRP and PRP4KA (Fig. 4E). Together, these results demonstrate that PRP4KA interacts directly with LRP in the nucleus.

Fig. 4. — (A) Yeast two-hybrid assay of the interaction between LRP and PRP4KA. Transformed yeast cells were grown on SD-Trp/-Leu medium (left) and SD-Trp/-Leu/-His medium supplemented with 3 mM 3-amino-1,2,4-triazole (3AT; right). (B) BiFC analysis of the interaction between LRP and PRP4KA. LRP and PRP4KA were fused to nEYFP and cEYFP to generate nEYFP-LRP and cEYFP-PRP4KA, respectively. Vectors containing nEYFP or cEYFP alone served as negative controls. Merge, merged images of EYFP, H2B-RFP, and bright field. Scale bar, 10 μm. (C) In vitro GST pull-down assay showing the interaction between GST-LRP and His-PRP4KA. Purified His-PRP4KA was divided into two portions and incubated with immobilized GST or GST-LRP. Immunoblotting was performed using anti-His (top) and anti-GST (bottom) antibodies. (D) Schematic diagram showing the T-DNA insertion site in *prp4ka-11* and C to T mutation in *prp4ka-4*. Black boxes, exons; black lines, introns; gray boxes, UTRs. (E) CoIP assay confirming the interaction between LRP and PRP4KA. Nuclear extracts from 9-day-old *prp4ka-11 gPRP4KA-3FLAG #5* and F1 crossed seedlings of *prp4ka-11 gPRP4KA-3FLAG #5* and *lrp-1 g4HA-LRP #2* were immunoprecipitated with anti-HA agarose. Input and coimmunoprecipitated proteins were detected using anti-FLAG (top) and anti-HA (bottom) antibodies. (F) Flowering phenotype of *prp4ka* mutants under LDs and SDs. (G) Flowering time of *prp4ka* mutants under LDs and SDs. Error bars, means ± SD; n = 20. Asterisks indicate statistically significant differences between WT and *prp4ka* mutants (**P < 0.01, two-tailed paired Student’s t test).

LRP and PRP4KA modulate FLC transcription and flowering in the same pathway

PRP4KA has been shown to mediate multiple aspects of development, including flowering (36); however, the mechanisms by which it influences flowering remains unclear. To explore its biological function of PRP4KA, we identified two prp4ka mutants, prp4ka-11 and prp4ka-4 (Fig. 4D and fig. S7A). Both mutants exhibited significantly delayed flowering under LDs and SDs (Fig. 4, F and G). gPRP4KA-3FLAG fully rescued the late-flowering phenotype of prp4ka-11 (fig. S7B), indicating that PRP4KA is responsible for the late-flowering phenotype observed in prp4ka-11. Consistent with its role in promoting flowering, qPCR analysis showed a gradual increase in PRP4KA expression during the floral transition and strong PRP4KA expression in rosette leaves (fig. S7, C and D). This expression pattern was similar to that of LRP (fig. S2, C and D). These results suggest that PRP4KA plays an important role in promoting flowering.

Since PRP4KA interacts with LRP, which represses FLC transcription (Fig. 3), we then examined FLC expression in prp4ka-11 mutants. FLC expression was consistently and markedly elevated in developing prp4ka-11 seedlings compared to wild-type plants during the floral transition (Fig. 5A). Correspondingly, FT and SOC1 expression was down-regulated in prp4ka-11 (fig. S7, E and F). Meanwhile, expression levels of MAF1, MAF4, and MAF5 were also up-regulated in prp4ka-11 (fig. S7G). FLC expression was restored in independent transgenic lines of prp4ka-11 gPRP4KA-3FLAG, which exhibited flowering time comparable to wild-type under LDs (Fig. 5, B and C, and fig. S7, H and I). Moreover, the late-flowering phenotype of prp4ka-11 was largely suppressed by flc-3 (Fig. 5, D and E), suggesting that FLC is a major downstream target of PRP4KA in flowering time control.

prp4ka-11 did not further enhance the late-flowering phenotype of lrp-1 (Fig. 5F), and FLC expression in the prp4ka-11 lrp-1 double mutants remained compared to that in each single mutant (Fig. 5G). Notably, flc-3 largely suppressed the late-flowering phenotype of lrp-1 prp4ka-11 (Fig. 5F and fig. S7J). These results suggest that LRP and PRP4KA function in the same pathway to repress FLC expression, prompting us to examine nascent FLC transcription of the prp4ka-11 and prp4ka-11 lrp-1. Nascent FLC transcript levels were significantly elevated to similar extents in lrp-1, prp4ka-11, and their double mutants as compared to wild type (Fig. 5H). Moreover, the frequency of the 5′-3′ gene looping at the FLC locus was similarly increased in lrp-1, prp4ka-11, and their double mutants relative to wild type (Fig. 5I), accompanied by comparable increases in Pol II occupancy across the FLC locus in these mutants (Fig. 5J). Together, these results suggest that PRP4KA and LRP function in the same pathway to prevent chromatin loop formation at FLC, inhibit RNA Pol II occupancy, and repress FLC transcription, thereby regulating flowering time.

PRP4KA phosphorylates LRP at Ser¹⁷ to enhance its stability

The in vivo interaction between LRP and the kinase PRP4KA and their overlapping function in flowering time regulation prompted us to explore whether PRP4KA phosphorylates LRP. To test this, we conducted in vitro kinase assays using purified kinase domain of PRP4KA (PRP4KA^K) fused to maltose-binding protein (MBP) and GST-LRP^N (N-terminal part of LRP fused to GST) or GST-LRP^C (C-terminal part of LRP fused to GST) proteins (fig. S8A). A mobility shift of GST-LRP^N by PRP4KA^K was observed in phosphate affinity SDS–polyacrylamide gel electrophoresis (PAGE), but not in GST-LRP^C (Fig. 6A and fig. S8A), implying that the shift corresponds to phosphorylated GST-LRP^N. These results indicate that PRP4KA phosphorylates LRP in vitro. We then identified specific phosphorylation site(s) on LRP mediated by PRP4KA by analyzing phosphorylated GST-LRP^N products from the in vitro kinase assay via mass spectrometry. Two potential phosphorylated sites, Ser(S)9 and S17, were identified (Fig. 6B and table S3). These two sites were then individually point-mutated to alanine (A) for an in vitro kinase assay. Notably, the S17A mutation greatly abolished in vitro phosphorylation of GST-LRP^N by MBP-PRP4KA^K (Fig. 6C), indicating that PRP4KA-dependent phosphorylation of LRP primarily occurs at S17. To confirm the in vivo phosphorylation of LRP, we immunoprecipitated 4HA-LRP from the lrp-1 g4HA-LRP transgenic plants using an anti-HA, followed by detection using an anti–phospho-(Ser/Thr) antibody which has been widely used for detecting phosphorylated proteins in plants (37–42). A phosphorylated protein band that overlapped with 4HA-LRP was detected by the anti–phospho-(Ser/Thr) antibody (Fig. 6D), confirming that LRP is phosphorylated in vivo.

Fig. 6. — (A) PRP4KA phosphorylates LRP in vitro. Phosphorylation of GST-LRP^N was detected by immunoblotting on Phos-tag–containing gels and regular SDS-PAGE using anti-GST antibody. (B) Mass spectrometry identification of PRP4KA-mediated phosphorylation sites on LRP. Identified phosphorylated LRP peptides are shown above their corresponding spectra with phosphorylated sites highlighted in red. *m/z*, mass/charge ratio. (C) In vitro kinase assay of WT and mutated GST-LRP^N variants by MBP-PRP4KA^K. LRP^N proteins were separated by SDS-PAGE with (top) or without Phos-tag (bottom) and detected using an anti-GST antibody. (D) Detection of in vivo LRP phosphorylation by an anti–Phospho-(Ser/Thr) antibody. Immunoprecipitated proteins from WT and *lrp-1 g4HA-LRP* were analyzed by immunoblotting using anti–Phospho-(Ser/Thr) and anti-HA antibody antibodies. (E) Immunoblot analysis of 4HA-LRP protein levels in *prp4ka-11*. Total protein extracts from 9-day-old seedlings were analyzed by immunoblotting using an anti-HA antibody. RbcL stained with Ponceau S served as an internal control. (F) Immunoblot analysis of 4HA-LRP protein levels following MG132 treatment. Total protein was extracted from 9-day-old seedlings that were either mock-treated or pretreated with 50 μM MG132 for indicated durations in the presence of cycloheximide (CHX). RbcL stained with Ponceaus S served as the loading control. Protein levels were normalized to RbcL and shown relative to the highest level in each blot set as 1.00. (G) Degradation of LRP and its variants in a cell-free degradation system. Equal amount of total protein extracts from WT and *prp4ka-11* was incubated with GST-LRP^N variants with or without MG132 for the indicated time periods. Remaining LRP^N proteins were detected by an anti-GST antibody. Tubulin served as a loading control. Protein levels were normalized to tubulin and shown relative to the highest level in each blot set as 1.00. Line graphs showing the degradation dynamics are shown in fig. S8D.

To assess the effect of PRP4KA-mediated LRP phosphorylation on LRP function, we generated lrp-1 g4HA-LRP in the prp4ka mutant background and observed a marked reduction in LRP-4HA protein abundance in prp4ka-11 in several independent lines, while LRP-4HA mRNA levels remained unchanged (Fig. 6E and fig. S8, B and C), suggesting that PRP4KA-depdenent phosphorylation of LRP may regulate LRP stability. To test this, we monitored the amount of 4HA-LRP in the prp4ka mutant background in the presence of MG132 and a protein synthesis inhibitor cycloheximide. MG132 treatment greatly increased the stability of 4HA-LRP (Fig. 6F). We next monitored the abundance of GST-LRP^N, a dephosphorylation-mimicking mutant GST-LRP^N(S17A), and a phosphorylation-mimicking mutant GST-LRP^N(S17D) in a cell-free protein degradation system, with or without the proteasome inhibitor N-carbobenzyloxy-l-leucyl-l-leucyl-l-leucinal (MG132). In the absence of MG132, GST-LRP^N degraded more rapidly in prp4ka-11 extracts compared to wild type, while MG132 treatment restored GST-LRP^N levels in both backgrounds (Fig. 6G and fig. S8D). These results indicate that PRP4KA protects LRP from proteasome-mediated degradation. Moreover, GST-LRP^N(S17A) degraded faster than GST-LRP^N or GST-LRP^N(S17D) in both wild-type and prp4ka-11 extracts in the absence of MG132 (Fig. 6G), suggesting that PRP4KA-mediated phosphorylation of LRP on S17 plays an important role on LRP stabilization. Together, these pieces of evidence substantiate that phosphorylation of LRP at S17 by PRP4KA enhances LRP protein stability.

PRP4KA-mediated LRP phosphorylation is critical for FLC repression and flowering time

To investigate the effects of PRP4KA-mediated phosphorylation of LRP on flowering, we transformed lrp-1 with either a dephosphorylation-mimicking construct (g4HA-LRP^S17A) or a phosphorylation-mimicking construct (g4HA-LRP^S17D). Independent transgenic lines of g4HA-LRP^S17D, but not those of g4HA-LRP^S17A, fully recovered the late-flowering phenotype and elevated FLC expression in 1rp-1 (Fig. 7, A to C, and fig. S8E). Notably, the protein level of 4HA-LRP^S17D was slightly higher than 4HA-LRP, whereas 4HA-LRP^S17A was barely detectable (Fig. 7D and fig. S8F), despite comparable mRNA levels among 4HA-LPR, 4HA-LRP^S17D, and 4HA-LRP^S17A (fig. S8E). MG132 treatment in prpk4ka-11 lrp-1 g4HA-LRP^S17A and prp4ka-11 lrp-1 g4HA-LRP^S17D further demonstrate that phosphorylation at S17 enhances LRP protein stability (fig. S8G). Further analyses revealed that the increased 5′-3′ looping and Pol II occupancy at FLC observed in lrp-1 were completely restored in lrp-1 g4HA-LRP and lrp-1 g4HA-LRP^S17D lines, but not in lrp-1 g4HA-LRP^S17A (Fig. 7, E and F). Consequently, nascent FLC transcript levels were rescued in lrp-1 g4HA-LRP and lrp-1 g4HA-LRP^S17D, but not in lrp-1 g4HA-LRP^S17A (Fig. 7G). Together, these results indicate that PRP4KA-mediated LRP phosphorylation at S17 is essential for FLC repression and proper flowering time regulation.

Fig. 7. — (A) Effect of mutated LRP proteins on flowering time under LDs. Representative lines of *lrp-1 g4HA-LRP #2*, *lrp-1 g4HA-LRP^S17A #3*, and *lrp-1 g4HA-LRP^S17D #4* were shown. (B) Flowering time analysis of *lrp-1 g4HA-LRP #2*, *lrp-1 g4HA-LRP^S17A #3*, and *lrp-1 g4HA-LRP^S17D #4* under LDs. Error bars, means ± SD; n = 20. Different letters indicate statistically significant differences (P < 0.05, one-way ANOVA with Tukey’s post hoc test). (C) *FLC* expression levels in 9-day-old seedlings of *lrp-1 g4HA-LRP #2*, *lrp-1 g4HA-LRP^S17A #3*, and *lrp-1 g4HA-LRP^S17D #4*. Error bars [(C), (E), (F), and (G)], means ± SD; n = 3. Different letters in (C), (E), (F), and (G) indicate statistically significant differences (P < 0.05, one-way ANOVA with Tukey’s post hoc test). (D) Immunoblotting analysis of LRP protein levels in *lrp-1 g4HA-LRP #2*, *lrp-1 g4HA-LRP^S17A #3*, and *lrp-1 g4HA-LRP^S17D #4*. Tubulin detected with an anti-tubulin antibody was used as a loading control. (E) Quantitative 3C-qPCR showing relative interaction frequency between the 5′ and 3′ regions of *FLC* in 9-day-old seedlings of *lrp-1 g4HA-LRP #2*, *lrp-1 g4HA-LRP^S17A #3*, and *lrp-1 g4HA-LRP^S17D #4.* Positions of the anchor and primers used for 3C-qPCR are shown in Fig. 3F. (F) ChIP analysis of RNA Pol II enrichment at the *FLC* locus in 9-day-old seedlings of *lrp-1 g4HA-LRP #2*, *lrp-1 g4HA-LRP^S17A #3*, and *lrp-1 g4HA-LRP^S17D #4*. (G) Nascent *FLC* levels in 9-day-old seedlings of *lrp-1 g4HA-LRP #2*, *lrp-1 g4HA-LRP^S17A #3*, and *lrp-1 g4HA-LRP^S17D #4*.

DISCUSSION

As one of the most abundant and evolutionarily conserved RNA binding domains in eukaryotes, RRM typically mediates interactions with RNA to regulate splicing, transport, translation, and decay (1, 2, 8). Beyond these canonical roles in posttranscriptional control, emerging evidence points to RRM-containing proteins also functioning in transcriptional regulation through chromatin association (9, 10), although the underlying mechanism remains largely unknown. The Arabidopsis genome encodes 196 annotated RRM-containing proteins (3), yet most remain functionally uncharacterized. In this study, we identify LRP as a chromatin-associated RRM-containing protein that plays a key role in repressing FLC expression and regulating flowering time (Fig. 8). LRP binds to both the 5′ and 3′ regions of FLC, preventing the 5′-3′ chromatin looping and limiting Pol II engagement. Loss of LRP leads to increased chromatin looping and Pol II recycling at FLC, leading to elevated FLC transcription and delayed flowering. Moreover, phosphorylation of LRP at Ser¹⁷ by PRP4KA is essential for its protein stability and function in chromatin looping and transcriptional repression. In the absence of PRP4KA, LRP is degraded by the proteasome, facilitating chromatin loop and Pol II recycling, thereby activating FLC transcription and delaying flowering.

Fig. 8. — PRP4KA-mediated phosphorylation of LRP at Ser¹⁷ enhances its protein stability, facilitating its binding to both the 5′ and 3′ regions of *FLC*. This binding inhibits the formation of 5′ to 3′ chromatin looping, accompanied by reducing RNA Pol II occupancy and *FLC* transcription, thereby promoting flowering. Loss of PRP4KA leads to LRP degradation, resulting in reduced binding of LRP to the *FLC* chromatin regions. This enhances 5′ to 3′ chromatin looping, which brings the promoter and terminator regions into close physical proximity, likely facilitating the efficient recycling of Pol II for subsequent rounds of transcription, thereby increasing *FLC* transcription and delaying flowering. Created in BioRender. Dbs, Y. (2025) https://biorender.com/0ggad0a.

Our findings establish LRP and PRP4KA as previously uncharacterized flowering promoters in repressing FLC transcription. First, lrp mutants show a daylength-insensitive late-flowering phenotypes under both LDs and SDs, and their flowering is accelerated by vernalization—characteristics typical of autonomous pathway mutants (43, 44). Second, FLC expression is greatly elevated in lrp mutants, and flc-3 completely suppressed its late-flowering phenotype, suggesting that FLC is the primary target of LRP in regulating the floral transition. Third, nascent FLC transcript levels are markedly increased in lrp mutants, indicating that LRP modulates FLC transcription. Moreover, LRP interacts with PRP4KA, whose mutants also exhibit daylength-insensitive late-flowering phenotypes with significantly elevated FLC nascent transcript levels. Notably, flc-3 nearly fully recovered the late-flowering phenotype of prp4ka lrp double mutants, underscoring the key roles of both PRP4KA and LRP in FLC transcriptional repression and flowering time control.

Chromatin architecture is crucial for gene regulation, as it controls spatial genome organization and facilitates interactions between regulatory elements and target genes (45, 46). The floral repressor FLC forms a chromatin loop by connecting its 5′ and 3′ flanking regions, a structure required for active transcription (21–24). Notably, our result suggests that LRP prevents this 5′-3′ FLC loop formation via direct binding to both its 5′ and 3′ regions. No loop is detected at the actively transcribed FLC transgene (23), and several mutants with high FLC expression show no obvious increase in gene looping (24), indicating that active transcription per se does not drive gene loop formation and that gene loop formation is a regulated process. LRP does not physically interact with HON4/5 or BAF60, chromatin regulators that prevents 5′-3′ FLC gene looping (22, 47), nor with components of PRC1 or PRC2 complexes, which have been implicated in chromatin loop formation (fig. S6) (48, 49). How LRP prevents FLC gene looping remains an intriguing question for future investigation. It has been shown that the FRI supercomplex, including FRI, histone methyltransferase, and other chromatin modifiers, are required for FLC 5′-3′ chromatin loop formation (23). It is therefore possible that LRP binding hinders access of the components of FRI supercomplex to the 5′ or 3′ regions of FLC, thereby inhibiting loop formation. This gene looping brings the promoter and terminator regions into close physical proximity, effectively shortening the distance between them and thereby likely facilitating the efficient recycling of Pol II for subsequent rounds of transcription (21, 22, 24, 50, 51). Consistent with this notion, our results show that in the lrp mutant background, where FLC gene looping is enhanced, Pol II enrichment is elevated across the FLC locus. In addition to the 5′-3′ loop, there is a shorter gene loop at FLC formed through interactions between its 5′ region and the first intron, and this loop is associated with FLC repression during vernalization (52). Thus, it will be intriguing to further investigate the potential role of LRP in modulating higher-order chromatin organization in future studies.

Our data show that LRP is evenly distributed throughout the nucleus and directly associates with FLC chromatin, suggesting an unconventional role for an RRM-containing protein in DNA binding. In contrast, its close homolog HRLP has been shown to form phase-separated nuclear bodies near intron I of the nascent FLC RNA to inhibit splicing and enhance R-loop formation (17). Supporting their distinct roles in modulating FLC expression, LRP and HRLP have additive effects on flowering time (fig. S9). Nevertheless, given the sequence similarity between LRP and HRLP, they may also share some RNA targets in other cellular processes. It has been reported that some RNA binding proteins exhibit higher affinity for single-strand DNA than for RNA (20), indicating that RNA binding proteins can directly binds to DNA during transcription, while others are recruited to chromatin through interactions with transcription factors (10). Further characterization of the LRP interactome will be valuable for elucidating the mechanism by which LRP associates with FLC chromatin. Moreover, future structural analysis of LRP, particularly the configuration of its RRM domains and intrinsically disordered sequences, will be crucial for understanding how it recognizes specific genomic loci such as FLC. Such insights could reveal previously unknown modes of nucleic acid recognition by RRM proteins and broaden our understanding of how RNA binding proteins can acquire DNA binding capabilities to influence chromatin looping and transcriptional regulation.

Since the presence and functional relevance of PTMs on RRM-containing proteins remained largely unexplored in plants, our findings provide insights into the regulatory mechanisms governing these proteins. We have demonstrated that phosphorylation of LRP by PRP4KA at Ser¹⁷ is essential for its protein stability, FLC repression, and flowering time regulation. The kinase PRP4KA interacts with LRP and specifically phosphorylates it at Ser¹⁷ to stabilize it, whereas the unphosphorylated form of LRP is preferentially degraded by the proteasome (Fig. 8). This subsequently affects FLC transcription and contributes to the timely floral transition. Supporting this, the phosphorylation-mimicking mutant 4HA-gLRP^S17D and the dephosphorylation-mimicking mutant 4HA-gLRP^S17A exhibit opposite effects on LRP protein stability, FLC gene looping, Pol II engagement, FLC transcription, and flowering time. Since the expression levels of LRP and PRP4KA increase during seedling development before the floral transition (figs. S2D and S7D), fine-tuning their relative abundance to modulate phosphorylated LRP protein levels is likely essential for the precise regulation of flowering. Nevertheless, how phosphorylation of LRP affects its protein stability remains an open question for future investigation. Such modifications may affect its interactions with E3 ubiquitin ligases, its association with stabilizing binding partners, or induce local conformational changes in its protein structures (53). It is also noteworthy that in addition to LRP, PRP4KA has also been shown to phosphorylate SERRATE (SE) for degradation, and two paralogs of PRP4KA, PRP4KB and PPR4KC, also phosphorylate SE (54, 55). It would be interesting to explore in future studies whether PRP4KB and PRP4KC also phosphorylate LRP or other RRM-containing proteins and assess their broader contributions to posttranscriptional and transcriptional regulation in plants. Furthermore, given the conserved nature of PRP4KA and LRP across various plant species, it would be intriguing to explore whether our identified PRP4KA-LRP-FLC module exists and could be modulated in crop species to ensure timely flowering for subsequent reproductive success.

MATERIALS AND METHODS

Plant materials and growth conditions

A. thaliana plants were grown on soil or Murashige and Skoog plates under LDs (16-hour light/8-hour dark) or SDs (8-hour light/16-hour dark) at 16° ± 1°C, 23° ± 2°C, or 28° ± 2°C. Mutants of lrp-1 (CS855920), prp4ka-11 (CS808147), and prp4ka-4 (CS71818) were obtained from the Arabidopsis Biological Resource Center, and lrp-2 was generated by CRISPR-Cas9–mediated gene editing. Mutants of flc-3 (30), ft-10 (56), co-9 (57), gi-1 (58), fca-2 (59), fpa-7 (31), ld-1 (60), fve-4 (61), flk-2 (62), and fld-3 (63) are in the Columbia-0 background as previously reported.

Plasmid construction and plant transformation

To construct g4HA-LRP, a 4.3-kb genomic fragment of LRP, including an ~2.0-kb promoter, the coding region with introns, and a 0.17-kb 3′UTR, was cloned into pENTR/D-TOPO (Invitrogen) to generate gLRP. A XmaI restriction site was introduced to gLRP immediately after the start codon using the QuikChange Site-Directed Mutagenesis Kit (Stratagene) to generate g(XmaI)LRP. g4HA-LRP, gNLS-4HA-LRP, gNES-4HA-LRP, gGFP-LRP, and gGUS-LRP were generated via cloning the corresponding sequences into the XmaI site based on g(XmaI)LRP. On the basis of the g4HA-gLRP vector, g4HA-LRP^S17A and g4HA-LRP^S17D were generated by overlapping PCR. To construct 35S:GFP-LRP, the full-length coding sequence of LRP was amplified and cloned into pGreen 0229 35S:GFP (64). To construct GST-LRP^N, the N-terminal fragment of LRP was amplified and cloned into pGEX-6p-2. To construct AmiR-LRP, a set of four primers was designed (http://wmd3.weigelworld.org) and used for PCR amplification according to the published protocol (28). The resulting PCR fragment was digested with EcoRI and BamHI and cloned into a modified pENTR vector (41). To construct gPRP4KA-3FLAG, a 5.6-kb genomic fragment of PRP4KA including a 1.2-kb promoter region, the coding regions, and introns was cloned into pENTR/D-TOPO (Invitrogen) to generate gPRP4KA. The sequence of 3FLAG was cloned into gPRP4KA to generate gPRP4KA-3FLAG. The primers used for vector construction are listed in table S4.

Transgenic plants were generated through the floral dipping method (65) using Agrobacterium tumefaciens (GV3101) harboring the desired vectors. AmiR-lrp and gGUS-LRP were transformed into wild-type plants. g4HA-LRP, gNLS-4HA-LRP, gNES-4HA-LRP, gGFP-LRP, g4HA-LRP^S17A, and g4HA-LRP^S17D vectors were transformed into lrp-1 mutants, while gPRP4KA-3FLAG was transformed into prp4ka-11 mutants. Transgenic plants were selected by hygromycin on Murashige and Skoog plates or Basta on soil.

Expression analysis

Total RNA was extracted with the FavorPrep Plant Total RNA Mini kit (Favorgen) and reverse-transcribed using the iScript Reverse Transcription Supermix (Bio-Rad) following the manufacturers’ instructions. qPCR was performed on three biological replicates using the SYBR Green PCR Master Mix (Applied Biosystems) on the CFX384 Real-Time PCR Detection System (Bio-Rad). The expression of TUB2 was included as an internal control. The difference between the cycle threshold (Ct) of target genes and the Ct of TUB2 (∆Ct = Ct_{target gene} − Ct_TUB2) was used to calculate the normalized expression of target genes as previously described (66). The primers used for gene expression analysis are listed in table S4.

GUS staining of the gGUS-LRP reporter line was performed as previously described (67). Seedlings at different developmental stages were fixed in ice-cold 90% acetone for 20 min, followed by three washes with rinse solution [50 mM Na₂HPO₄, 50 mM NaH₂PO4, 0.5 mM K₃Fe(CN)₆, and 0.5 mM K₄Fe(CN)₆]. The tissues were subsequently infiltrated with staining solution (rinse solution supplemented with 2 mM X-Gluc) under vacuum for 10 min and incubated at 37°C. After staining, the tissues were cleared of chlorophyll with an ethanol series and observed under a light microscope in clearing solution (7.5 g of gum arabic, 100 g of chloral hydrate, 5 ml of glycerin, and 30 ml of distilled water).

RNA-seq and analysis

Nine-day-old wild-type and lrp-1 seedlings grown under LDs were collected for RNA extraction with the RNeasy Plus Mini kit (QIAGEN) according to the manufacturer’s instructions. RNA quality and quantity were analyzed by gel electrophoresis and an Agilent Bioanalyzer 2100 system. Sequencing libraries were generated using the NEBNext Ultra II Directional RNA Library Prep Kit, and sequencing was performed on the Illumina Novaseq 6000 platform. Sequencing reads were aligned to the Arabidopsis TAIR10 genomes using STAR 2.7.3a with default parameters. Differential gene expression analysis was performed using DESeq2 v1.18.1 (68).

Chromatin-bound RNA extraction

Chromatin-bound RNA was extracted as previously described (17, 69). Briefly, ~5 g of seedlings was ground into fine powder and then homogenized in Honda buffer [0.44 M sucrose, 10 mM MgCl₂, 20 mM Hepes, 1.25% (w/v) Ficoll, 2.5% (w/v) dextran, 0.5% Triton X-100, 5 mM dithiothreitol (DTT), 1× protease inhibitor, ribonuclease (RNase) inhibitor (20 U/ml), and tRNA (50 ng/μl)]. The mixture was filtered through two layers of Miracloth and centrifuged. The resulting pellet was then resuspended in the resuspension buffer [25 mM tris-HCl (pH 7.5), 50% glycerol, 100 mM NaCl, 1 mM DTT, 0.5 mM EDTA, 1× protease inhibitor, and RNase inhibitor (20 U/ml)] and washed twice with UREA wash buffer [0.5 mM EDTA, 25 mM tris-HCl (pH 7.5), 1% Tween 20, 1 mM DTT, 300 mM NaCl, and 1 M urea]. TRIzol reagent (Thermo Fisher Scientific) was used for RNA extraction. After deoxyribonuclease I (DNase I; Roche) treatment, RNA was further purified with TRIzol reagent. Purified chromatin-bound RNA was used for reverse transcription with FLC gene–specific primers, followed by qPCR analysis to detect nascent FLC RNA levels. Primers used for detecting nascent FLC expression were listed in table S4.

ChIP assay

ChIP assays were performed as previously described (67). Seedlings were collected 9 days after germination and fixed in MC buffer [10 mM KH₂PO₄ (pH 7.0), 50 mM NaCl, and 100 mM sucrose] with 1% formaldehyde for 40 min on ice under vacuum. Fixed tissues were then homogenized in liquid nitrogen, and chromatin was extracted and sonicated to generate DNA fragments ranging from 200 to 500 bp. 4HA-LRP were immunoprecipitated by anti-HA agarose conjugate (Sigma-Aldrich), while Pol II was immunoprecipitated using anti–RNA Pol II CTD antibody (Abcam, catalog no. ab26721, RRID:AB_777726) bound to Protein G Plus agarose beads (Santa Cruz Biotechnology). Fold enrichment of each fragment was determined by qPCR, with a genomic fragment of TUB2 serving as an internal control. All ChIP experiments were performed three times, yielding similar results. Primers used for ChIP assays were listed in table S4.

3C assay

The 3C assay was performed as previously described (70, 71). Briefly, 9-day-old seedlings were fixed with 1% (v/v) formaldehyde, and nuclei were isolated with nuclear isolation buffer [15 mM Pipes (pH 6.8), 150 mM sucrose, 5 mM MgCl₂, 60 mM KCl, 15 mM NaCl, 1 mM CaCl₂, 1 mM phenylmethylsulfonyl fluoride, 0.9% Triton X-100, and protease inhibitor cocktail]. The nuclei were treated with 0.3% (w/v) SDS at 65°C for 40 min and 37°C for 20 min with gentle shaking, followed by sequestration with 2% Triton X-100 at 37°C for 60 min. Chromatin was then digested with 400 U of Dpn II (New England Biolabs) overnight at 37°C, and the enzyme was inactivated by adding 1.6% (w/v) SDS and incubating at 65°C for 20 min. Ligation was performed using 200 U of DNA ligase at room temperature for 5 hours. Reverse cross-linking and protein digestion were carried out by adding proteinase K and incubating overnight at 65°C. DNA was then purified using phenol-chloroform-isoamyl alcohol extraction and ethanol precipitation. A 10-kb genomic FLC plasmid subjected to the same procedure was used as a control template to normalize the relative interaction frequency. An FLC genomic region without Dpn II cutting site was amplified as an internal control for qPCR. Primers used for 3C-qPCR were listed in table S4.

Yeast two-hybrid assay

The coding sequences of various genes were amplified and cloned into pGBKT7 (BD) or pGADT7 (AD) vectors (Clontech) using the ClonExpress II One Step Cloning Kit (Vazyme). Various combinations of AD and BD vectors were cotransformed into AH109 cells using the Yeastmaker Yeast Transformation System 2 (Clontech). The transformed yeast cells were cultured on selection media. Primers used for vector construction were listed in table S4.

BiFC analysis

The full-length coding sequences of LRP and PRP4KA were cloned into the primary pSAT1 vectors (72). The resulting cassette, including the constitutive promoters and fusion proteins, were cloned into the pGreen binary vector pHY105 (73). Agrobacterium suspensions harboring the desired vector combinations were coinfiltrated into the abaxial surfaces of N. benthamiana leaves, and leaves were examined under a confocal microscope 2 days after infiltration. Primers used for vector construction were listed in table S4.

In vitro GST pull-down assay

The coding region of LRP was cloned into the pGEX-6p-2 vector (Pharmacia). GST and GST-LRP proteins were produced by inducing Escherichia coli Rosetta (DE3) harboring the corresponding vectors using 0.4 mM isopropyl-β-d-thiogalactopyranoside (IPTG) overnight at 16°C. The soluble GST and GST-LRP proteins were isolated, immobilized on glutathione sepharose beads (Amersham Biosciences), and incubated with protein purified from His-PRP4KA. Proteins retained on the beads were resolved by SDS-PAGE and detected using the anti-HIS antibody (Santa Cruz Biotechnology, catalog no. sc-8036, RRID:AB_627727).

Coimmunoprecipitation

Seedlings of prp4ka-11 gPRP4KA-3FLAG #5 and F1 crosses of prp4ka-11 gPRP4KA-3FLAG #5 and lrp-1 g4HA-LRP #2 were harvested at 9 days after germination. Nuclei were isolated using the nuclear isolation buffer [20 mM KCl, 25% glycerol, 20 mM tris (pH 7.0), 30 mM β-mercaptoethanol, 2.5 mM MgCl₂, 0.7% Triton X-100, 2 mM EDTA (pH 8.0), 250 mM sucrose, and 1× protease inhibitor cocktail (Roche)]. The isolated nuclei were resuspended in IP buffer [50 mM Hepes (pH 7.5), 150 mM KCl, 10 μM ZnSO₄, 0.05% SDS, 5 mM MgCl₂, 1% Triton X-100, and 1× protease inhibitor cocktail] and incubated with anti-HA agarose conjugate (Sigma-Aldrich) for 4 hours at 4°C. Nuclear protein extracts (input) and the immunoprecipitated proteins were resolved by SDS-PAGE and detected using anti-HA (Santa Cruz Biotechnology, catalog no. sc-7392 HRP, RRID:AB_2894930) and anti-FLAG (Thermo Fisher, catalog no. MA1-91878-HRP, RRID:AB_2537626) antibodies.

Phos-tag SDS-PAGE

Phos-tag SDS-PAGE was performed as previously described (74). Phos-tag (50 μM; Wako Pure Chemical Industries) and MnCl₂ (100 μM) were added to the SDS-PAGE separation gels. After electrophoresis, the Phos-tag gel was washed three times with wash buffer (25 mM tris-HCl, 192 mM glycine, 20% methanol, and 10 mM EDTA) and then three times with transfer buffer (25 mM tris-HCl, 192 mM glycine, and 20% methanol). After washing, the protein gel was blotted, blocked, and subsequently probed with anti-GST (Santa Cruz Biotechnology) or anti-HA (Santa Cruz Biotechnology) antibodies.

In vitro kinase assay

The sequences of LPR^N and LRP^C were cloned into the pGEX-6p-2 vector (Pharmacia), while the sequence of PRP4KA^K was cloned into pMAL-C2X (New England Biolabs). The recombinant proteins were produced by E. coil rosette (DE3) cells with the induction by 0.4 mM IPTG overnight at 16°C. Soluble GST, GST-LRP^N, and GST-LRP^C were purified with glutathione sepharose beads (Amersham Biosciences), while MBP and MBP-PRP4KA^K proteins were purified using amylose resin (New England Biolabs). The concentrations of purified proteins were determined by Bradford assay (Bio-Rad).

The in vitro kinase assay was performed as described previously (74, 75). Briefly, 1 μg of purified MBP-PRP4KA^K protein was incubated with 1 μg of GST-LRP^N in 12 μl of kinase buffer [25 mM tris (pH 7.5), 12 mM MgCl₂, 1 mM DTT, and 1 mM adenosine triphosphate (ATP)] at 37°C for 30 min. The reaction products were then analyzed by 8% SDS-PAGE with or without Phos-tag and detected by anti-GST (Santa Cruz Biotechnology). In addition, the reaction products were also analyzed by a mass spectrum on a TripleTOF 5600 system (AB Sciex) to identify PRP4KA-mediated phosphorylation sites on LRP. Candidate target sites were then mutated into Ala and subjected to in vitro kinase assay to confirm PRP4KA target sites on LRP.

Protoplast isolation and transformation

Arabidopsis protoplasts were isolated from 2-week-old flc-3 and flc-3 lrp-1 plants grown under LDs and transfected as described previously (76). The vectors containing genomic sequences of FLC^WT-T, FLC^MAF-T, or FLC^NOS-T were transfected into the isolated Arabidopsis protoplast. FLC gene looping and expression were determined as above described.

Cell-free degradation assay

Cell-free degradation assay was performed as previously described with minor modifications (74). Nine-day old wild-type and prp4ka-11 plants grown under LDs were collected for total protein extraction using the extraction buffer [25 mM tris-HCl (pH 7.4), 150 mM NaCl, 1% NP-40, 1 mM EDTA, and 5% glycerol] supplemented with freshly added 1× protease inhibitor cocktail (Roche). Protein concentrations were measured using the Bradford assay (Bio-Rad). GST-LRP^N (200 ng) was incubated with 500 μg of total protein extract in the reaction buffer [20 mM tris-HCl (pH 7.5), 10 mM MgCl₂, 1 mM DTT, and 1 mM ATP] with or without 50 μM MG132 at 37°C for 60, 120, and 180 min. GST-LRP^N protein levels in different reactions were analyzed by SDS-PAGE followed by detection using anti-GST antibody (Santa Cruz Biotechnology). Band intensities were quantified using ImageJ software (US National Institutes of Health).

Cross-linking immunoprecipitation and qPCR

The cross-linking immunoprecipitation and qPCR (CLIP-qPCR) was performed as previously described with some modifications (77, 78). Briefly, 9-day-old WT and lrp-1 g4HA-LRP seedlings were irradiated on ice with 254-nm ultraviolet (UV) light at a dose of 600 mJ/cm² in a Stratalinker 2400 UV cross-linker. Plants were then frozen and ground into fine powders in liquid nitrogen. Samples were then lysed in lysis buffer [50 mM tris-HCl (pH 7.4), 100 mM NaCl, 1% Igepal CA-630, 0.1% SDS, 0.5% sodium deoxycholate, and 1× protease inhibitor cocktail], partially digested with RNase I, precleared with Protein A/G Plus Agarose (Santa Cruz Biotechnology) for 1 hour at 4°C, and then incubated with anti-HA agarose conjugate (Sigma-Aldrich) for 3 hours at 4°C. The bound beads were washed three times with wash buffer I [50 mM tris-HCl (pH 7.4), 1 M NaCl, 1 mM EDTA, 1% Igepal CA-630, 0.1% SDS, and 0.5% sodium deoxycholate] and once with wash buffer II [20 mM tris-HCl (pH 7.4), 10 mM MgCl₂, 0.1 M KCl, and 0.02% Triton X-100]. Following DNase digestion, the RNA was further purified with TRIzol reagent (Thermo Fisher Scientific) and reversed transcribed with gene-specific primers. Primers used for CLIP-qPCR assay were listed in table S4.

Statistical analysis

IBM SPSS Statistics software (version 19) was used to conduct statistical analyses. A two-tailed paired Student’s t test was used to assess significant differences between two groups, while differences among multiple groups were analyzed using one-way analysis of variance (ANOVA), followed by Tukey’s post hoc tests.

Acknowledgments

We thank the Arabidopsis Biological Resource Centre for providing seeds. We thank the Protein and Proteomics Centre in the Department of Biological Sciences, National University of Singapore for mass spectrometry service. We thank Y. Hao and members of Shen laboratory for discussion and comments on the manuscript.

Funding:

This work was supported by the National Research Foundation Competitive Research Programme (NRF-CRP22–2019-0001), Singapore Food Story R&D Programme (SFS_RND_SUFP_001_04), and the intramural research support from Temasek Life Sciences Laboratory.

Author contributions:

L.S. conceived this study. S.F. and L.S. designed the experiments. S.F., Y.Z., S.Z., Z.T., Q.Z., and L.S. performed the experiments. S.F., Y.Y., and L.S. analyzed data and wrote the paper. All authors read and approved the manuscript.

Competing interests:

The authors declare that they have no competing interests.

Data and materials availability:

All data and code needed to evaluate and reproduce the results in the paper are present in the paper and/or the Supplementary Materials. Sequencing reads are available in the National Center for Biotechnology Information (NCBI) Gene Expression Omnibus (GEO) database under the accession number: PRJNA1256247.

Supplementary Materials

The PDF file includes:

Figs. S1 to S9

Legends for tables S1 to S3

Table S4

sciadv.adz0645_sm.pdf^{(2.1MB, pdf)}

Other Supplementary Material for this manuscript includes the following:

Tables S1 to S3

sciadv.adz0645_tables_s1_to_s3.zip^{(85.6KB, zip)}

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Associated Data

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

Supplementary Materials

Figs. S1 to S9

Legends for tables S1 to S3

Table S4

sciadv.adz0645_sm.pdf^{(2.1MB, pdf)}

Tables S1 to S3

sciadv.adz0645_tables_s1_to_s3.zip^{(85.6KB, zip)}

Data Availability Statement

[R1] 1.Fan S., Zhang Y., Zhu S., Shen L., Plant RNA-binding proteins: Phase separation dynamics and functional mechanisms underlying plant development and stress responses. Mol. Plant 17, 531–551 (2024). [DOI] [PubMed] [Google Scholar]

[R2] 2.Clery A., Blatter M., Allain F. H., RNA recognition motifs: Boring? Not quite. Curr. Opin. Struct. Biol. 18, 290–298 (2008). [DOI] [PubMed] [Google Scholar]

[R3] 3.Lorković Z. J., Barta A., Genome analysis: RNA recognition motif (RRM) and K homology (KH) domain RNA-binding proteins from the flowering plant Arabidopsis thaliana. Nucleic Acids Res. 30, 623–635 (2002). [DOI] [PMC free article] [PubMed] [Google Scholar]

[R4] 4.Corley M., Burns M. C., Yeo G. W., How RNA-binding proteins interact with RNA: Molecules and mechanisms. Mol. Cell 78, 9–29 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]

[R5] 5.Koster T., Marondedze C., Meyer K., Staiger D., RNA-binding proteins revisited - the emerging Arabidopsis mRNA interactome. Trends Plant Sci. 22, 512–526 (2017). [DOI] [PubMed] [Google Scholar]

[R6] 6.Maris C., Dominguez C., Allain F. H., The RNA recognition motif, a plastic RNA-binding platform to regulate post-transcriptional gene expression. FEBS J. 272, 2118–2131 (2005). [DOI] [PubMed] [Google Scholar]

[R7] 7.Lunde B. M., Moore C., Varani G., RNA-binding proteins: Modular design for efficient function. Nat. Rev. Mol. Cell Biol. 8, 479–490 (2007). [DOI] [PMC free article] [PubMed] [Google Scholar]

[R8] 8.Prall W., Sharma B., Gregory B. D., Transcription is just the beginning of gene expression regulation: The functional significance of RNA-binding proteins to post-transcriptional processes in plants. Plant Cell Physiol. 60, 1939–1952 (2019). [DOI] [PubMed] [Google Scholar]

[R9] 9.Shao W., Bi X., Pan Y., Gao B., Wu J., Yin Y., Liu Z., Peng M., Zhang W., Jiang X., Ren W., Xu Y., Wu Z., Wang K., Zhan G., Lu J. Y., Han X., Li T., Wang J., Li G., Deng H., Li B., Shen X., Phase separation of RNA-binding protein promotes polymerase binding and transcription. Nat. Chem. Biol. 18, 70–80 (2022). [DOI] [PubMed] [Google Scholar]

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PERMALINK

Site-specific phosphorylation of LRP regulates FLC chromatin looping and flowering

Sheng Fan

Yu Zhang

Shaobo Zhu

Zhenfeng Teng

Qiqi Zhang

Yuanyuan Yan

Lisha Shen

Roles

Abstract

INTRODUCTION

RESULTS

LRP promotes flowering in Arabidopsis

Fig. 1. LRP promotes flowering in Arabidopsis.

LRP functions in the nucleus to promote flowering

Fig. 2. LRP functions within the nucleus to regulate flowering.

LRP represses nascent FLC transcription

Fig. 3. LRP inhibits FLC gene looping and transcription.

LRP binds to FLC chromatin and prevents chromatin looping

LRP interacts with PRP4KA

Fig. 4. LRP interacts with PRP4KA.

LRP and PRP4KA modulate FLC transcription and flowering in the same pathway

Fig. 5. PRP4KA functions in the same pathway as LRP to repress FLC transcription.

PRP4KA phosphorylates LRP at Ser17 to enhance its stability

Fig. 6. PRP4KA phosphorylates LRP mainly at Ser17 to stabilize it.

PRP4KA-mediated LRP phosphorylation is critical for FLC repression and flowering time

Fig. 7. PRP4KA-mediated phosphorylation of LRP inhibits FLC chromatin looping and transcription.

DISCUSSION

Fig. 8. A model depicting the regulation of flowering time through site-specific phosphorylation of LRP by PRP4KA.

MATERIALS AND METHODS

Plant materials and growth conditions

Plasmid construction and plant transformation

Expression analysis

RNA-seq and analysis

Chromatin-bound RNA extraction

ChIP assay

3C assay

Yeast two-hybrid assay

BiFC analysis

In vitro GST pull-down assay

Coimmunoprecipitation

Phos-tag SDS-PAGE

In vitro kinase assay

Protoplast isolation and transformation

Cell-free degradation assay

Cross-linking immunoprecipitation and qPCR

Statistical analysis

Acknowledgments

Funding:

Author contributions:

Competing interests:

Data and materials availability:

Supplementary Materials

The PDF file includes:

Other Supplementary Material for this manuscript includes the following:

REFERENCES

Associated Data

Supplementary Materials

Data Availability Statement

ACTIONS

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PRP4KA phosphorylates LRP at Ser¹⁷ to enhance its stability

Fig. 6. PRP4KA phosphorylates LRP mainly at Ser¹⁷ to stabilize it.