Bruno-like proteins modulate flowering time via 3′ UTR-dependent decay of SOC1 mRNA


  • Hyung-Sae Kim,

    1. Division of Bioscience and Bioinformatics, Myongji University, Yongin, Kyunggi-do, South Korea
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  • Nazia Abbasi,

    1. School of Biotechnology and Environmental Engineering, Myongji University, Yongin, Kyunggi-do, South Korea
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  • Sang-Bong Choi

    Corresponding author
    1. School of Biotechnology and Environmental Engineering, Myongji University, Yongin, Kyunggi-do, South Korea
    • Division of Bioscience and Bioinformatics, Myongji University, Yongin, Kyunggi-do, South Korea
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Author for correspondence:

Sang-Bong Choi

Tel: +82 31 330 6199



  • The Bruno RNA-binding protein (RBP) has been shown to initially repress the translation of oskar mRNA during Drosophila oogenesis and later to be involved in a broad range of RNA regulation.
  • Here, we show that homologous constitutive overexpression of each of two Arabidopsis thaliana Bruno-like genes, AtBRN1 and AtBRN2, delayed the flowering time, while the atbrn1 atbrn2-3 double mutant flowered early and exhibited increased expression of APETALA1 (AP1) and LEAFY (LFY) transcripts.
  • Crossing of 35S::AtBRNs with SOC1 101-D plants demonstrated that 35S::AtBRNs suppress an early-flowering phenotype of SOC1 101-D in which the coding sequence (CDS) with the 3′ UTR of SUPPRESSOR OF OVEREXPRESSION OF CONSTANS1 (SOC1) gene is overexpressed. However, this early-flowering phenotype by SOC1 overexpression was maintained in the plants coexpressing 35S::AtBRNs and 35S::SOC1 without the 3′ UTR (3′ UTR).
  • Using yeast three-hybrid, electrophoretic mobility shift, RNA immunoprecipitation, and protoplast transient assays, we found that AtBRNs bind to the 3′ UTR of SOC1 RNA and participate in mRNA decay, which was mediated by the distal region of the SOC1 3′ UTR. Overall, AtBRNs repress SOC1 activity in a 3′ UTR-dependent manner, thereby controlling the flowering time in Arabidopsis.


Post-transcriptional control, including mRNA transport, stability, and translation, plays a crucial role in plant growth and development (Choi et al., 2000; Okita & Choi, 2002). Most of these processes are inevitably achieved either directly or indirectly by RNA-binding proteins (RBPs) (Keene, 2001; Lorković & Barta, 2002). All earlier known RNA recognition motif (RRM)- and K homology (KH)-type RBPs involved in flowering, for example, Flowering Time Control protein A (FCA), FPA and FLOWERING LOCUS K (FLK), act as flowering activators (Macknight et al., 1997; Schomburg et al., 2001; Lim et al., 2004; Mockler et al., 2004). Of functionally identified RBPs, FCAs have been extensively characterized with respect to autoregulation by alternative splicing (Macknight et al., 2002; Quesada et al., 2003) and RNA-mediated FLC chromatin silencing (Baurle et al., 2007; Liu et al., 2007a, 2010; Manzano et al., 2009), thereby autonomously promoting flowering by the repression of FLC expression. It has been previously reported that AtBRUL-1 and AtBRUL-2 (hereafter AtBRN1 and AtBRN2, respectively) belong to the Bruno RBP family (Good et al., 2000). Following the first report of Bruno in flies, a number of Bruno homologs (also known as CELF, CUG-BP, and ETR) have been annotated on the basis of RRM structures in other organisms; for example, six in humans (Barreau et al., 2006), three, including FCA, in Arabidopsis (Good et al., 2000), at least three in Drosophila melanogaster, six in Xenopus laevis (Amato et al., 2005), and one in Caenorhabditis elegans (Good et al., 2000). Drosophila Bruno functions as a translation repressor of Oskar and Nanos mRNA before their localization to the posterior pole of the oocyte (Kim-Ha et al., 1995; Good et al., 2000; Chekulaeva et al., 2006). Bruno binds to cis-acting elements present in the 3′ UTR of Oskar mRNA called Bruno response elements (BREs) with CUP (Kim-Ha et al., 1995; Igreja & Izaurralde, 2011). In more recent studies, the Bruno–CUP–eIF4E complex inhibits the eIF4E–eIF4G interaction, resulting in the translation repression of Oskar mRNA (Wilhelm et al., 2003; Macdonald, 2004; Nakamura et al., 2004). Therefore, translational repression of Oskar mRNA is dependent on the interaction of the 5′ and 3′ UTRs, which is mediated by an eIF4E complex (Wilhelm et al., 2003; Macdonald, 2004). In another model, Bruno participates in forming 50–80S ribonucleoprotein particles (RNPs), which act as silencing particles by assembling on Oskar mRNA and consequently preventing access of the translational machinery (Chekulaeva et al., 2006).

Nevertheless, Bruno specifically disrupts the translation of BRE-containing mRNA, although its specificity may also be controlled by the spatiotemporal expression of proteins that interact with Bruno. For instance, the eIF4E2 transcript is predominantly accumulated in floral tissues and is distinct from eIF4E1, which is constitutively expressed in all organs (Good et al., 2000). FCAs may not have the capacity to bind to BRE or to directly regulate translation repression, because it functions as a component of poly(A) factor and/or is involved in RNA-mediated asymmetric methylation of single and low-copy genes at the chromatin level (Baurle et al., 2007). Based on the early- and late-flowering phenotype observed in atbrns double mutants and 35S::AtBRNs transgenic plants, the genes implicated in the control of flowering time were examined in this study. AtBRNs redundantly repress the activity of the SOC1 protein by interacting with the 3′ UTR of SOC1 mRNA. These results suggest that post-transcriptional coordination occurs in the control of flowering time in addition to previously known regulatory systems occurring at the transcriptional level.

Materials and Methods

Plant materials and growth conditions

Arabidopsis thaliana (L.) Heynh Col-0 seeds were grown on Murashige and Skoog (MS) media (1 × MS, 1% sucrose, and 0.8% agar, pH 5.7) or on soil in a conditioned room at 22°C under long-day (LD, 16 h light (120 μmol photons m−2 s−1) : 8 h dark) or short-day (SD, 8 h light : 16 h dark) cycles. T-DNA insertion mutant alleles were atbrn1 (Salk_041205), atbrn2-1 (Salk_135530), atbrn2-2 (Salk_054409), and atbrn2-3 (Sail_549_D06). The transgenic overexpression lines used in this study – 35S::FT (Kardailsky et al., 1999), SOC1 101-D (Lee et al., 2000), 35S::AGL24 (Yu et al., 2002), and 35S::AP1 (Mandel & Yanofsky, 1995) – are in the Col-0 background. For selection, 50 μg ml−1 kanamycin (Duchefa, Haarlem, the Netherlands), 50 μg ml−1 hygromycin (Duchefa), or 50 μg ml−1 basta (Duchefa) were added to the MS media. Seeds were stratified in darkness at 4°C for 3 d before transferring to the growth room.

Flowering time measurements

Flowering time was measured by counting the number of rosette leaves at the time of flowering. For statistical analysis, 30–50 plants were measured, and a mean was taken for each measurement.

Plasmid construction

For 35S::AtBRNs constructs, full-length coding sequences (CDS) of AtBRN1 (At4g03110) and AtBRN2 (At1g03457) were amplified by PCR from cDNA clones, pda 06133 (RIKEN, Japan) and U60225 (ABRC, Ohio State University, OH, USA), using primers AtBRN1F and AtBRN1R for AtBRN1, and AtBRN2F and AtBRN2R for AtBRN2, respectively. The resulting DNA fragments (1326 and 1290 bp) were digested with NcoI and XbaI and cloned downstream of the 35S dual promoter in the pRTL2 vector (Restrepo et al., 1990). Finally, DNA inserts were generated by PstI digestion and cloned into pCAMBIA3301 (CAMBIA, Canberra, Australia), producing 35S::AtBRN1 and 35S::AtBRN2. For the PAtBRNs::GUS construct, promoter regions spanning 3.8 and 2.0 kb upstream of the translation initiation site of the AtBRN1 and AtBRN2 genes were amplified by PCR from genomic DNA using primers ProAtBRN1F and ProAtBRN1R for AtBRN1, and ProAtBRN2F and ProAtBRN2R for AtBRN2. The resulting DNA fragments were digested with EcoRI and NcoI and then cloned into pCAMBIA 3301, producing PAtBRN1::GUS and PAtBRN2::GUS. For the 35S::GFP:AtBRNs construct, the CDS of AtBRN1 and AtBRN2 were amplified by PCR from cDNA clones, with primers 407F and 105R for AtBRN1 and 408F and 106R for AtBRN2, and cloned into pK7WGF2 (Karimi et al., 2002), producing 35S::GFP:AtBRN1 and 35S::GFP:AtBRN2, respectively. For the yeast three-hybrid assay, hybrid RNA (MS2-SOC1) and fusion protein (VP16 AD-AtBRNs) plasmids were generated using pRH5′ and pYESTrp3 vectors (Invitrogen), respectively. Full-length SOC1 cDNA (1293 bp), including the 5′ UTR and 3′ UTR, was first amplified by reverse transcriptase-polymerase chain reaction (RT-PCR) using the primers 126F and 126R and then cloned into the AatII and SmaI sites of pRH5′ vector, resulting in pMS2-SOC1. CDS of AtBRN1 and AtBRN2 were amplified by PCR from cDNA clones with 124F and 124R for AtBRN1 and with 125F and 125R for AtBRN2, respectively. The resulting DNA fragments were digested with EcoRI and NotI for AtBRN1 and with HindIII and NotI for AtBRN2, followed by cloning into the pYESTrp3 vector, resulting in pVP16-AtBRN1 and pVP16-AtBRN2, respectively. For the electrophoretic mobility shift assay (EMSA), plasmids for GST-AtBRN1 and GST-AtBRN2 fusion proteins were constructed. The CDS of AtBRN1 and AtBRN2 were amplified by PCR using cDNA clones as templates with 231F and 231R for AtBRN1 and 232F and 232R primer pairs for AtBRN2, and subsequently cloned using BamHI and NotI site of pGEX-5X-1 (GE Healthcare, Piscataway, NJ, USA). For the transient protoplast assay, four types of SOC1 DNA fragments were generated with or without the BRE sequence: for example, GFP-SOC1 I (-BRE), GFP-SOC1 II (-BRE), GFP-SOC1 III (-BRE), and GFP-SOC1 IV (+BRE). The DNA fragments were amplified from cDNA clones using primer pairs 445F and 242R for GFP-SOC1 I, 242F and 446R for GFP-SOC1 II, 242F and 447R for GFP-SOC1 II, and 242F and 448R for GFP-SOC1 IV. The fragments were subsequently cloned into p2FGW7 (Karimi et al., 2002), producing constructs 35S::GFP-SOC1 I, 35S::GFP-SOC1 II, 35S::GFP-SOC1 III, and 35S::GFP-SOC1 IV.

Analyses of RNA expression by β-glucuronidase (GUS) staining and RT-PCR

Histochemical GUS staining and RT-PCR were performed as described previously (Kim et al., 2010). Real-time quantitative RT-PCR (qRT-PCR) was performed as described previously (Abbasi et al., 2010). For qRT-PCR, first-strand cDNA synthesis was performed using 5 μg of total RNA from seedling. Quantification was done using the Bio-Rad iQ5 Real-time PCR detection system with 2× Real-time PCR Pre-mix with Evagreen (Solgent, Korea), according to the manufacturer's instructions. The tubulin expression was used to normalize the expression ratio for each gene. Quantitative RT-PCR experiments were performed using two independent biological samples and with three technical replicates. Fold changes in gene expression were calculated via the inline image method (Livak & Schmittgen, 2001). The primers used for real-time PCR are listed in Supporting Information, Table S1.

Detection of RNA–protein interaction using yeast three-hybrid, electrophoretic mobility shift, and RNA immunoprecipitation assays

A yeast three-hybrid assay was performed using an RNA-Protein Hybrid Hunter™ Kit (Invitrogen). Briefly, yeast L40-ura3 transformants harboring both the hybrid RNA (pMS2:SOC1-R) and hybrid protein (pVP16:AtBRNs) vectors were selected on synthetic media plates lacking uracil and tryptophan and further assayed for β-galactosidase expression on the plates containing X-gal and for growth on selective media lacking histidine. Cotransformants containing both pVP16:AtBRNs and pRH5′ (MS2 only) were employed as a negative control. EMSA was carried out using synthetic biotin-labeled 2′-O-Methyl RNA (49 nt) containing the putative BRE sequence of SOC1 mRNA, the GST-AtBRNs fusion proteins, and the LightShift Chemiluminescent EMSA Kit (Thermo Scientific, Rockford, IL, USA). GST-AtBRNs fusion proteins were induced and purified using a glutathione sepharose CL-4B column (GE Healthcare) according to the manufacturer's instructions. RNA–protein binding reactions were carried out in 5× binding buffer (10 mg BSA, 250 mM KCl, 10 mM MgCl2, 5 mM EDTA, 5 mM DTT, 1% Nonidet P-40, 50% glycerol, and 100 mM Tris-HCl, pH 7.6) by incubating at room temperature for 20 min. To minimize the nonspecific RNA–protein interactions, a poly(A) (300 ng μl−1) competitor was used. The complex was resolved on 8% polyacrylamide gel in TBE buffer (45 mM Tris-borate, pH 8.0, 1.0 mM EDTA) and transferred onto Amersham Hybond™–N+ (GE Healthcare) using an electroblotter. The membrane was blocked, incubated with streptavidin-horseradish peroxidase antibody diluted to 1 : 300, washed and developed according to the manufacturer's protocol.

RNA immunoprecipitation (RIP) was performed as previously described (Peritz et al., 2006). Briefly, Arabidopsis seedling extract was prepared using an immunoprecipitation buffer (100 mM KCl, 5 mM MgCl2, 10 mM Hepes, 1 mM DTT, and 0.5% Nonidet P-40) and then incubated with 3 μg ml−1 polyclonal anti-AtBRN2 or nonimmune rabbit-IgG at 4°C overnight. The RNA–protein immunocomplex was precipitated by protein A-agarose beads (Sigma) and subjected to RT-PCR. The AtBRN2 antibody was raised in rabbits using the specific synthetic peptide (CQQSKNPLFNGLLNS) and purified using an AtBRN2 peptide affinity resin (Peptron Inc., Daejeon, South Korea).

Protein expression analysis

Total soluble proteins were extracted from Arabidopsis using an extraction buffer (125 mM Tris-HCl, pH 8.8/1% SDS/10% glycerol/50 μM Protease Inhibitor Cocktail (Sigma). Protein was separated using 12% sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) and transferred onto a PVDF membrane (Qbiogene Inc., Carlsbad, CA, USA) in solution using an electroblotter. Filters were blocked for 1 h with Tris-buffered saline (TBS) supplemented with 6% nonfat dry milk (Bio-Rad) containing 0.1% Tween 20 and incubated for 2 hs in blocking buffer with anti-SOC1 (1 : 200, Peptron Inc.), anti-GUS, anti-green fluorescent protein (anti-GFP), anti-Lhca2 (1 : 5000, Abcam), and anti-actin (1 : 5000, Sigma) antibodies. The blot was washed three times with TBS plus 0.1% Tween 20 for 15 min each. Anti-rabbit and anti-mouse antibodies (Sigma), conjugated to horseradish peroxidase, were diluted 1 : 10 000 in blocking buffer and then added to the blot for 1 h. The blot was washed and developed using Luminol/Enhancer solution (Thermo Scientific). The SOC1 antibody was raised in rabbit using a specific synthetic peptide (RHTKDRVSTKPVSEEN) and purified using a SOC1 peptide affinity resin (Peptron, Inc.).

Transient assay

Arabidopsis mesophyll protoplasts were isolated from 4-wk-old 35S::AtBRNs plants (Yoo et al., 2007). Protoplasts were transfected with constructs expressing GFP-SOC1 I, GFP-SOC1II, GFP-SOC1III, and GFP-SOC1IV. Protoplasts were incubated for 12 h after transfection, and GFP intensity was measured following the above-mentioned protein expression analysis.

Confocal laser scanning microscopy

Arabidopsis root cells expressing GFP fusion constructs were examined under a laser-scanning confocal microscope (FV1000SPD, Olympus, Tokyo, Japan). GFP signals in the plant tissues were detected by excitation with the 488 nm line of the argon laser and a capturing emission at 522 nm.


Expression of AtBRN1 and AtBRN2

A phylogeny constructed of Bruno-like proteins revealed that AtBRN1 and AtBRN2 are more closely related to rice OsBRNs than to other Bruno-like proteins derived from nonplant organisms, such as the fruit fly, frog, and human (Fig. S1a, Table S2). All three RRM domains, of which two are in the N-terminal half and the third one is on the C-terminal end, are well conserved in AtBRN1 and AtBRN2 (Figs 1a, S1b). However, only two domains are found in FCA, an ELAV-type ribonucleoprotein-1 (Macknight et al., 1997; Good et al., 2000).

Figure 1.

Expression analysis of Bruno. (a) Protein structure of AtBRN1 and AtBRN2. Each protein contains three RNA recognition motifs (RRMs). (b) AtBRN1 (black bars) and AtBRN2 (gray bars) expression in wildtype Arabidopsis thaliana plants. The AtBRN1 transcript from a 10-d-old Col-0 seedling was used as a reference. Bars indicate + SD of the mean of three replicates. (c) The temporal expression patterns of AtBRN1 and AtBRN2 at different growth stages. Protein gel blot analysis of total protein extract from AtBRN1::GUS and AtBRN2::GUS plants, with anti-beta-glucuronidase (anti-GUS) and anti-actin as loading controls. (d) Genomic DNA structure of genes encoding AtBRN1 and AtBRN2, showing the T-DNA insertion sites (left panel). Genotyping and reverse transcriptase-polymerase chain reaction (RT-PCR, right panel) of the atbrn1/atbrn2-3 double mutant. Positions of primers used for genotyping and RT-PCR are indicated below the diagram.

To gain insight into the function of AtBRNs, we first examined the expression levels of AtBRNs by qRT-PCR (Fig. 1b) and GUS histochemical staining (Fig. S2a-w). AtBRN1 showed a higher expression level in the silique, stem, and cauline leaf, while AtBRN2 was highly expressed in the silique, stem, and flower (Fig. 1b). Overall, AtBRN2 was expressed at lower levels than AtBRN1. Consistently, a GUS staining was observed in all of the tissues of PAtBRNs::GUS transgenic plants (Fig. S2a-w). In an immunoblot analysis using a GUS antibody against PAtBRNs::GUS transgenic plants, GUS expression was highest at 7 d after germination (DAG) and gradually decreased as plants reached the flowering stage (Fig. 1c), implying that AtBRN genes are negatively related to flowering.

Flowering time regulation of AtBRNs

We next identified mutant lines for each AtBRN gene: atbrn1 for AtBRN1 and atbrn2-1, atbrn2-2, and arbrn2-3 for AtBRN2. Each single mutant not only showed an unnoticeable phenotype, but atbrn2-1 and atbrn2-2 were also leaky alleles as homozygous for T-DNA yet having knockdown expression activity relative to the wildtype, and an atbrn1 atbrn2-3 double mutant was generated (Fig. 1d). For comparison, 35S::AtBRN1 and 35S::AtBRN2 plants were produced (Fig. S3). Except for flowering time, the overexpression and double mutant plants were indistinguishable from wildtype plants. Overexpression of AtBRNs and atbrn1/atbrn2-3 double mutant plants showed late- and early-flowering phenotypes under LD and SD conditions, respectively (Fig. 2a). In addition, 35S::AtBRNs plants still exhibited the late-flowering phenotypes under SD conditions after vernalization. However, the effect of the atbrn1/atbrn2-3 double mutant was decreased after vernalization (Fig. 2a), indicating that the role of the AtBRNs in the repression of flowering may be related to the promotion of flowering by vernalization.

Figure 2.

AtBRNs are involved in the regulation of flowering time. (a) Flowering phenotype of AtBRNs overexpression and mutant Arabidopsis. Arabidopsis was grown under long-day and short-day conditions, as well as short-day conditions after 8 wk of vernalization treatment. Flowering time was measured by counting the number of rosette leaves. *, P < 0.05; Student's t-test. (b) Expression pattern of flowering time genes in Arabidopsis (Col-0, black bars; atbm1/atbm2-3, gray bars). Each gene from 10-d-old Col-0 seedlings was used as a reference. Bars indicate + SD of the mean of three replications.

As indirect evidence of this possibility, the expression of AtBRNs was decreased gradually from the vegetative to the floral stage (Fig. 1c). To further address if AtBRNs are affected by vernalization, we analyzed the expression of AtBRNs after vernalization in wildtype plants and a vernalization-insensitive mutant (vin3) (Sung & Amasino, 2004). Vernalization decreased the expression of AtBRNs in wildtype plants, but the expression was not altered in vin3 plants (Fig. S4). Because vin3 blocked the vernalization response, the effect of vernalization on the expression of AtBRNs was weak.

As AtBRNs control flowering, we next examined the expression levels of regulators of flowering time in the atbrn1 atbrn2-3 double mutant to identify putative target gene(s), for example, FLOWERING LOCUS T (FT), FLOWERING LOCUS C (FLC), FLK, CONSTANS (CO), SOC1, AGAMOUS- LIKE24 (AGL24), AP1, and LFY. The expression levels of most of the flowering pathway genes were not altered compared with those in wildtype plants, except for LFY, AP1, and FT, which were up-regulated 3.3-, 4.2-, and 2.5-fold, respectively (Fig. 2b).

Suppression of the SOC1 101-D phenotype by AtBRN overexpression

To understand the epistatic relationship between AtBRNs and the flowering genes that are potential targets of AtBRNs, we crossed 35S::AtBRNs plants with SOC1 101-D, 35S::AGL24, 35S::LFY, 35S::AP1, and 35S::FT plants. In contrast to the activation tagging mutant SOC1 101-D, which flowers very early (Lee et al., 2000), the 35S::AtBRN1/SOC1 101-D and 35S::AtBRN2/SOC1 101-D crosses flowered identically to wildtype plants (Fig. 3a,b), indicating that AtBRNs repress the SOC1 101-D early-flowering phenotype. In addition to 35S::AtBRNs/SOC1 101-D, the other double-overexpression plants that overexpressed genes downstream of the SOC1 and AtBRN genes flowered early, a phenotype identical to that of the plants that overexpressed a single gene (Fig. 3a,b). This result was verified in the independent SOC1 101-D plants that were transformed with 35S::AtBRNs constructs. All of the 35S::AtBRN1/SOC1 101-D (= 5) and 35S::AtBRN2/SOC1 101-D (= 4) homozygous plants flowered later than the SOC1 101-D plants; however, the lateness of their flowering was dependent on the expression level of AtBRNs (Fig. 4a,b). Taken together, 35S::AtBRNs completely suppressed the SOC1 101-D early-flowering phenotype but did not suppress 35S::AGL24, 35S::AP1, 35S::FT, and 35S::LFY, which suggests that the AtBRNs act upstream of AGL24, AP1, FT, and LFY and downstream of SOC1. Based on the result that 35S::AtBRNs suppress the early-flowering phenotype of SOC1 101-D plants (Fig. 3a,b), although the level of SOC1 mRNA is only slightly affected in atbrn1/atbrn2-3 mutant plants when compared with wildtype plants (Fig. 2b), it is likely that SOC1 mRNA is post-transcriptionally regulated by AtBRNs.

Figure 3.

Overexpression of AtBRNs suppresses the early flowering of SOC1 101-D plants. (a) Rosette leaf numbers and comparison picture of 35S::AtBRNs crossed plants. *, P < 0.05; Student's t-test. Bars indicate + SD of the mean of three replications. (b) Comparison picture of 35S::AtBRNs crossed plants. Bar, 1 cm.

Figure 4.

Effect of overexpression of AtBRNs on flowering time in a SOC1 101-D background. (a) Phenotype comparison between 35S::AtBRNs/SOC1 101-D plants and SOC1 101-D early-flowering plants. *, P < 0.05; Student's t-test. Bars indicate + SD of the mean. (b) AtBRN1 and AtBRN2 expression in 35S::AtBRNs/SOC 101-D plants.

3′ UTR-mediated SOC1 mRNA destabilization by AtBRNs

As the Drosophila Bruno represses translation of Oskar mRNA during localization to the posterior end of the egg cell by binding to the BREs of the 3′ UTR sequence, [U(G/A)U(A/G)U(G/A)U] (Kim-Ha et al., 1995; Chekulaeva et al., 2006; Snee et al., 2007, 2008), AtBRNs are likely to bind to cis-element(s) on the SOC1 mRNA. Indeed, SOC1 mRNA has a potential BRE (UAUGUAU) cis-element located in the 3′ UTR. To test the possibility that AtBRNs bind to this putative BRE, we first performed a yeast three-hybrid assay (SenGupta et al., 1996) to detect the interaction between AtBRN proteins and the full-length SOC1 transcript (5′ UTR+CDS+3′ UTR). Coexpression of AtBRN proteins and SOC1 mRNA activated HIS3 and lacZ gene expression, while coexpression of AtBRNs and the empty vector did not (Fig. 5a). We then examined the direct interaction of AtBRNs with the SOC1 3′ UTR sequence by a gel retardation assay using purified GST-AtBRN fusion proteins and a biotin-labeled SOC1 3′ UTR (Fig. 5b). GST-AtBRNs bound to the 3′ UTR of SOC1 mRNA (lanes 3 and 8), while control GST did not bind to RNA (lanes 2 and 7). In a competition assay, a 10-fold molar excess of the cold substrate (SOC1 3′ UTR) reduced the formation of the SOC1-AtBRN1 RNP complex (lanes 4 and 9), and a 50-fold molar excess of competitor inhibited the complex formation almost completely (lanes 5 and 10). To further verify the interaction of AtBRNs and SOC1 mRNA, we performed RNA immunoprecipitation (RIP) using an AtBRN2 antibody (an AtBRN1 antibody was not available), which could recognize specific AtBRN protein followed by RT-PCR. To show specificity of anti-AtBRN2 antibody, we performed a western blot analysis. A high degree of specificity is observed for the anti-AtBRN2 antibody detecting a band c. 48 kDa in size in wildtype and 35S::AtBRN2 seedlings and not in atbrn2 knockout plants. As a loading control, we probed the blot with anti-actin antibody (Fig. S5). The SOC1 transcript was not detected in precipitates obtained using preimmune serum, whereas the transcript was amplified in the precipitates obtained using the AtBRN2 antibody (Fig. 5c), demonstrating that AtBRN2 interacts with SOC1 mRNA in planta.

Figure 5.

AtBRNs bind to the 3′ UTR of SOC1 mRNA in vitro and in vivo. (a) Yeast three-hybrid assay using the AtBRNs and SOC1 full-length RNA. (b) Electrophoretic mobility shift assay (EMSA) using GST-AtBRNs protein and biotin-labeled 2′-O-methyl SOC1 3′ UTR RNA. (c) The AtBRN2–RNA complexes were immunoprecipitated with anti-AtBRN2 peptide antibodies from Arabidopsis seedling extract, followed by reverse transcriptase-polymerase chain reaction (RT-PCR). Bars indicate + SD of the mean. (d) Green fluorescent protein (GFP) localization in transgenic Arabidopsis root cells expressing 35S::GFP, 35S::AtBRN1:GFP, and 35S::AtBRN2:GFP. Bar, 5 μm.

In the root cells of 35S::GFP:AtBRN1 and 35S::GFP:AtBRN2 plants, GFP signals were exclusively localized in the cytoplasm. However, GFP-AtBRN2 forms speckles within the cytoplasm (Fig. 5d), suggesting that AtBRNs do not have nuclear functions such as alternative splicing and polyadenylation that are exhibited by the other Bruno-like protein FCA (Quesada et al., 2003). Considering the localization of AtBRNs and the SOC1 mRNA which must be transported into the cytosol for translation, it is most likely that AtBRNs bind SOC1 mRNA in cytosol. To investigate this possibility, we checked the subcellular localization of AtBRNs-GFP in the presence of overexpressed SOC1 3′ UTR. To direct AtBRNs to the nucleus, we have fused the nuclear localization signal (NLS) to the downstream of AtBRNs. The fusion protein AtBRNs-NLS-GFP was confined to the nucleus. We also cotransformed 35S::AtBRNs:NLS:GFP with either 35S::SOC1(3′ UTR) lacking 3′ UTR or 35S::SOC1(+3′ UTR) containing 3′ UTR. When SOC1(3′ UTR) mRNA was coexpressed with AtBRNs-NLS-GFP in protoplasts, there was no significant difference in AtBRNs-NLS-GFP localization compared with the control where AtBRNs-NLS-GFP was solely expressed (Fig. 6). However, when SOC1(+3′ UTR) was coexpressed, the AtBRNs-NLS-GFP localized in cytoplasm, indicating that AtBRNs-NLS-GFP present in the nucleus moved to the cytoplasm for binding to 3′ UTR of cytoplasmic SOC1 mRNA. Further, when AtBRNs-NLS-GFP and SOC1(+3′ UTR) were coexpressed in los4-1 mutant that is defective in mRNA export and thus accumulates mRNA in the nucleoplasm (Gong et al., 2005), the fusion protein AtBRNs-NLS-GFP was mainly localized to the nucleus. Before AtBRNs-NLS-GFP moves to nucleus, some proteins that bind SOC1 (+3′ UTR) mRNA appeared to remain in the cytoplasm. All together, cytoplasmic localization of AtBRNs-NLS-GFP is the result of binding of AtBRNs to SOC1 mRNA.

Figure 6.

AtBRNs bind to the 3′ UTR of SOC1 mRNA in vivo. AtBRNs bind specifically to SOC1 3′ UTR in vivo. Green fluorescent protein (GFP) localization in protoplasts expressing 35S::AtBRN:GFP, 35S::AtBRNs:NLS:GFP/35S::SOC1 (-UTR), and 35S::AtBRNs:NLS:GFP/35S::SOC1 (+UTR). Bar, 100 μm.

Next, to investigate whether AtBRNs exert repression activity in plants in a SOC1 3′ UTR-dependent manner, we compared the flowering phenotype of the 35S::SOC1(CDS)/35S::AtBRNs plants with the SOC1 101-D/35S::AtBRNs plants in which an entire SOC1 gene, including the CDS and 3′ UTR, was expressed. 35S::SOC1(CDS)/35S::AtBRNs plants exhibited an early-flowering phenotype similar to the 35S::SOC1(CDS) plants, while SOC1 101-D/35S::AtBRNs plants exhibited a normal flowering phenotype similar to wildtype plants (Fig. 7a,b), indicating that AtBRNs suppress the early flowering caused by SOC1 overexpression only in the presence of the 3′ UTR of SOC1 mRNA. We then examined the expression levels of SOC1 mRNA and protein in Col-0, SOC1 101-D, 35S::SOC1(CDS), 35S::AtBRNs/SOC1 101-D, and 35S::AtBRNs/35S::SOC1(CDS) plants. 35S::AtBRNs/SOC1 101-D plants displayed greatly reduced SOC1 transcript and protein abundances, whereas 35S::AtBRNs/35S::SOC1(CDS) plants produced almost the same abundances of SOC1 transcript and protein as did 35S::SOC1(CDS) plants (Fig. 7c). Overexpression of AtBRN1 and AtBRN2 reduced SOC1 transcripts to 8.3 and 16.6%, respectively, in the SOC1 101-D background, whereas these transcripts were reduced to 17.8 and 48.4% in the wildtype background. The SOC1 protein was not detected in wildtype seedlings, most likely because of the low hybridization capacity of the antibody. However, overall expression levels of SOC1 RNA and protein correlate well with the flowering phenotype in the plants examined here (Fig. 7a–c).

Figure 7.

AtBRNs suppress SOC1 activity via binding to the SOC1 mRNA 3′ UTR. (a) Phenotype comparison between 35S::AtBRNs/SOC1 101-D crossed plants and 35S::AtBRNs/35S::SOC1 (-UTR) crossed plants. Bar, 1 cm. (b) Rosette leaf numbers and comparison picture of 35S::AtBRNs/SOC1 101-D crossed plants and 35S::AtBRNs/35S::SOC1 (-UTR) crossed plants. *, P < 0.05; Student's t-test. (c) The expression pattern of SOC1 RNA and protein in wildtype and crossed plants. Anti-actin is shown as the loading control. (d) Graphical representation of band intensities obtained from 35S::AtBRNs leaf protoplast transfected with the GFP-SOC1 I, GFP-SOC1II, GFP-SOC1III, and GFP-SOC1IV constructs. (35S::AtBRN1, black bars; 35S::AtBRN2, gray bars.) Bars indicate + SD of the mean of three replications.

To further examine if the 3′ UTR of SOC1 mRNA is involved in the decay of SOC1 mRNA, we performed a transient assay using protoplasts derived from 35S::AtBRNs transgenic plants. Truncated 35S::GFP:SOC1 3′ UTR constructs, for example, SOC I, SOCII, SOCIII, and SOCIV (Fig. S6), were transformed into protoplasts, and then GFP-SOC1 expression was tested using a GFP antibody. The GFP level was greatly reduced only in the presence of the distal 3′ UTR where the putative BRE (UAUGUAU) is located (Fig. 7d,S7). Consequently, our finding demonstrates that AtBRNs redundantly regulate flowering time by a 3′ UTR-dependent SOC1 mRNA decay.


To ensure reproductive success, flowering time is sophisticatedly regulated by developmental processes that integrate various external and internal factors in plant. Recent studies showed that post-transcriptional control also plays important roles in flowering time regulation (Bezerra et al., 2004; Doyle et al., 2005; Kuhn et al., 2007; Liu et al., 2007b). In our study, AtBRNs regulate the flowering time through the post-transcriptional control of SOC1 activity. The atbrn1/atbrn2-3 double mutant flowered early under LD and SD conditions, and under SD conditions after vernalization treatment (Fig. 2a), and the early-flowering phenotype was associated with increased expression of AP1, LFY, and FT. Although these integrators have overlapping functions, SOC1 definitely regulates LFY expression (Lee & Lee, 2010). Consistently, the LFY transcript showed the highest rate of increase in these integrators. SOC1 RNA, which is not regulated by AtBRNs, could activate LFY expression in atbrn1/atbrn2-3. Considering that up-regulation of AGL24 and SOC1 increases each other's expression (Michaels et al., 2003; Liu et al., 2008), AGL24 expression was slightly increased in proportion to the SOC1 expression in double mutant. Genetic analyses of epistasis indicate that AtBRNs directly regulate SOC1 or act downstream of SOC1 in the signaling pathway of flowering. Consistent with this epistasis result, AtBRNs directly bind 3′ UTR of SOC1 RNA in vitro and in vivo, thereby presumably tuning the SOC1 activity. In summary, the early-flowering phenotype of the double mutant is the result of the increased expression of these flowering integrators, except for AGL24.

Unlike SOC1 101-D plants that show early flowering and contain 3′ UTR in their transcripts driven by their native promoter, 35S::AGL24, 35S::AP1, 35S::FT, and 35S::LFY plants overexpress only protein CDS. Hence, it is possible for one to postulate that in the presence of 3′ UTR, the transcripts of flowering genes driven by 35S promoter might be targets of AtBRNs. However, our results and other previous reports showed that the transcripts of these flowering genes are not direct targets of AtBRNs. First, when SOC1 mRNA without 3′ UTR is overexpressed, these flowering gene transcripts containing 3′ UTR were increased in planta (Michaels et al., 2003; Liu et al., 2008; Lee & Lee, 2010). If these increased transcripts are regulated as the increased SOC1 transcript, AtBRN would suppress their transcript abundance. In our study, however, 35S::SOC1(-UTR) showed the early flowering, whereas AtBRNs did not rescue their early-flowering phenotype. We also analyzed SOC1, AP1, LFY, and FT expression in the precipitates obtained using preimmune serum and the AtBRN2 antibody. Besides SOC1 RNA, other tested transcripts were not detected, suggesting that, of the flowering gene transcripts tested in the current study, only the SOC1 transcript is able to interact directly with AtBRN2.

In this study, 35S::AtBRNs completely suppressed the SOC1 101-D early-flowering phenotype; therefore, AtBRNs act as repressors of the SOC1 flowering integrator. Although other mRNAs could be targets of AtBRNs, it is very likely that SOC1 mRNA is a prominent target of AtBRNs. AtBRNs suppressed the SOC1 mRNA activity via binding to SOC1 mRNA 3′ UTR, which was shown by in vitro and protoplast assays. The phenotypic consequence of the repression of SOC1 mRNA translation was seen when SOC1 mRNA was overexpressed with or without the 3′ UTR. In order to elucidate how AtBRNs regulate SOC1 activity, we will consider possible mechanisms for AtBRN-mediated repression.

To maintain the dosage effect of a particular mRNA in the cell, mRNA decay is an effective method of post-transcriptional regulation (Wilusz & Wilusz, 2004). The process of mRNA decay requires specific nucleases, cis-acting sequences, and trans-acting factors (Beelman & Parker, 1995). Cis-elements in mRNAs play important roles in regulating their degradation through their interaction with RBPs. Of these, AU-rich elements (AREs) (Shaw & Kamen, 1986) and GU-rich elements (GREs) (Vlasova et al., 2008) are well-characterized cis-acting elements involved in mRNA degradation. Although AREs and GREs are absent in the SOC1 mRNA, a putative BRE is located near the distal end of the 3′ SOC1 mRNA. Thus, we first considered the possibility that AtBRNs are involved in mRNA decay. In the 35S::AtBRNs/SOC1 101-D plants, where SOC1 mRNA was overaccumulated, SOC1 mRNA was rapidly removed. Therefore, it is very likely that the BRE in the SOC1 mRNA might be a binding sequence that is responsible for regulating its stability through an interaction with AtBRNs. However, other sequences than BREs might be involved in Bruno protein–target mRNA interaction. In the initial stages of our Bruno protein study, we carried out in vitro binding assays to identify mRNA targets for AtBRNs proteins. We found that, although a number of genes such as At2g32460, At5g58140, At5g13180, At5g64270, and At5g53480 contain BREs in the 3′ UTR region of their mRNAs, they fail to form a complex with AtBRN proteins. Although the SOC1 protein was below the level of detection in 35::AtBRNs/SOC1 101-D plants, the transcript and protein abundances of SOC1 were lower in 35::AtBRNs/SOC1 101-D than in SOC1 101-D. Overall, the SOC1 protein was reduced in proportion to its transcript in 35::AtBRNs/SOC1 101-D plants (Fig. 7c). Similar to the translational repression of Oskar mRNA by the Drosophila Bruno, which is involved in female and male gametogenesis and also early embryogenesis (Kim-Ha et al., 1995), AtBRNs repress mRNA translation via mRNA decay and consequently affect flowering, which is distinct from the role of the Drosophila Bruno protein.


We thank Dr Ilha Lee for providing the SOC1 101-D plants and Ji Hoon Ahn for the 35S::FT and 35S::AP1 plants. T-DNA insertion mutant seeds, atbrn1 (Salk_041205), atbrn2-1 (Salk_135530), atbrn2-2 (Salk_054409), and atbrn2-3 (Sail_549_D06), were obtained from Arabidopsis Biological Resource Center (ABRC) (Ohio State University). This work was supported by grants from the Next-Generation BioGreen21 Program, Rural Development Administration (no. PJ008206), and National Research Foundation funded by the Ministry of Education, Science and Technology (no. 2011-0011281), Korea.