Skip to main content
NCBI home page
Journal of Experimental Botany logoLink to Journal of Experimental Botany
. 2013 Jul 30;64(11):3299–3312. doi: 10.1093/jxb/ert183

Sequence variations of the partially dominant DELLA gene Rht-B1c in wheat and their functional impacts

Wen Wen 1, Qingyan Deng 1, Haiyan Jia 1, Lingzhu Wei 1, Jingbo Wei 1, Hongshen Wan 1, Liming Yang 1, Wenjin Cao 1, Zhengqiang Ma 1,*
PMCID: PMC3733159  PMID: 23918966

Abstract

Rht-B1c, allelic to the DELLA protein-encoding gene Rht-B1a, is a natural mutation documented in common wheat (Triticum aestivum). It confers variation to a number of traits related to cell and plant morphology, seed dormancy, and photosynthesis. The present study was conducted to examine the sequence variations of Rht-B1c and their functional impacts. The results showed that Rht-B1c was partially dominant or co-dominant for plant height, and exhibited an increased dwarfing effect. At the sequence level, Rht-B1c differed from Rht-B1a by one 2kb Veju retrotransposon insertion, three coding region single nucleotide polymorphisms (SNPs), one 197bp insertion, and four SNPs in the 1kb upstream sequence. Haplotype investigations, association analyses, transient expression assays, and expression profiling showed that the Veju insertion was primarily responsible for the extreme dwarfing effect. It was found that the Veju insertion changed processing of the Rht-B1c transcripts and resulted in DELLA motif primary structure disruption. Expression assays showed that Rht-B1c caused reduction of total Rht-1 transcript levels, and up-regulation of GATA-like transcription factors and genes positively regulated by these factors, suggesting that one way in which Rht-1 proteins affect plant growth and development is through GATA-like transcription factor regulation.

Key words: DELLA protein, dominance, GATA transcription factor, sequence variation, wheat.

Introduction

Wheat is a food staple for ~35% of the world population, and the continued population growth is presenting us with a formidable challenge to meet even greater demand for wheat in the near future. Nevertheless, recent data indicate that wheat yield potential is approaching a ceiling because of the lack of a complete understanding of yield formation mechanisms, and the paucity of breakthrough genes in modern cultivars. Because appropriate plant height is a key factor contributing to greater yield potential, investigations to elucidate the functional mechanisms involved in dwarfing are of substantial value to the future of agriculture.

Multiple genetic alternations can result in dwarfism. A common cause is the mutation of genes controlling biosynthesis of the plant hormones gibberellins (GAs), and genes involved in the GA signalling pathway (Peng et al., 1999; Hedden, 2003; Sun and Gubler, 2004). GAs are a group of diterpenoid carboxylic acids, and play a crucial role in the regulation of physiological processes related to plant growth and development, for example seed germination, shoot/stem elongation, cell division, and flower development (Davies, 1985; Fleet and Sun, 2005; Swain and Singh, 2005). Ross et al. (1997) classified GA-related dwarf mutants based on exogenous GA responses into GA-deficient and GA-insensitive types. GA-deficient mutants are typically ascribed to being GA biosynthesis deficient, and the GA-insensitive mutants result from mutations of positive and negative regulatory genes in the GA signalling pathway (Gomi and Matsuoka, 2003). Several negative regulators have been isolated by characterization of the recessive (loss-of-function) slender mutants and dominant (gain-of-function) GA-unresponsive dwarf mutants (Peng et al., 1997; Ikeda et al., 2001). DELLA proteins are well-studied regulators of GA signalling, which in planta function as repressors, and are components of the GID1–DELLA–SCFGID2 complex (Sun and Gubler, 2004; Murase et al., 2008). The complex formation stimulated by GA leads to DELLA protein degradation by means of the 26S proteasome and repression release.

Since the isolation of the DELLA protein gene GAI in Arabidopsis thaliana (Peng et al., 1997), its orthologues have been characterized in several other plants, including d8 in maize (Zea mays), VvGAI in grape (Vitis vinifera), MdRGL1 in apple (Malus×domestica Borkh), BrRGA in Brassica rapa, SLN1 in barley (Hordeum vulgare), Rht in wheat (Triticum aestivum), and SLR1 in rice (Oryza sativa) (Peng et al., 1999; Boss and Thomas 2002; Chandler et al., 2002; Itoh et al., 2002; Muangprom et al., 2005; Foster et al., 2007; Lawit et al., 2010). The encoded proteins share an N-terminal DELLA domain, including DELLA and VHYNP motifs, a poly(S/T) region, and a C-terminal GRAS domain. DELLA domain defects render the proteins resistant to GA-induced degradation, and result in dwarf phenotypes.

More than 20 dwarfing or reduced height genes (Rht) and alleles have been reported in wheat, including GA-insensitive genes (Rht-1: Rht-B1b, Rht-B1c, Rht-B1d, Rht-D1b, Rht-D1c, etc.) and GA-sensitive genes (Rht4, 5, 6, 7, etc.) (Konzak, 1988; Ross et al., 1997). The GA-insensitive and some GA-sensitive genes, such as Rht16 and 18, also cause coleoptile length reduction and leaf elongation, while the remaining GA-sensitive genes do not compromise early plant growth (Ellis et al., 2004). Even at the same locus, different alleles of the dwarfing gene exhibit various effects on plant height (Pearce et al., 2011). For example, of the five alleles detected at the Rht-B1 locus, Rht-B1c (Rht3) exhibited a three times stronger effect on plant height compared with Rht-B1b (Rht1) (Flintham and Gale, 1983; Flintham et al., 1997). Furthermore, Rht-B1c resulted in increased tiller and root number, a higher photosynthetic rate, and a higher chlorophyll content (Zhang et al., 1995; Li et al., 2006). Particularly intriguing was the finding that Rht-B1c exclusively inhibited α-amylase production in ripening grains, and enhanced seed dormancy (Flintham and Gale, 1982; Flintham et al., 1997). However, the mechanisms underlying the functional differences of various alleles remain unclear.

The present study served to elucidate why the Rht-B1c allele has multiple effects on plant growth, and the underlying factors contributing to increased phenotypic expression in reducing plant height. The Rht-B1c genomic sequence covering both the promoter and coding regions was isolated, and it was shown that the sequence insertion in the promoter and some single nucleotide polymorphisms (SNPs) represent evolution’s imprint on the B genome, and the retrotransposon insertion in the coding region leads not only to Rht-B1c transcript change and DELLA motif primary structure disruption, but also to modifications of the Rht-1 gene expression level and responses to GA.

Materials and methods

Plant materials

DS3 and DY3 lines were provided courtsey of Professor Zhao Yinhuai of Jiangsu Academy of Agricultural Sciences (JAAS). They were developed through crossing common wheat S3 and Y3 with the Rht-B1c donor Tom Thumb, and backcrossing for 23 generations using S3 and Y3 as recurrent parents. Segregating F2 populations were developed by selfing BC23F1 plants.

One hundred and twenty-six lines from the core common wheat germplasm collection were obtained from the Chinese Academy of Agricultural Sciences (Supplementary Table S1 available at JXB online). Other plant materials are listed in Supplementary Table S2.

Growth conditions and morphological evaluation

Plant height measurement

The BC23F1:2 families and parentals were planted in a field on the JAAS campus in 2003. The 126 common wheat lines were planted in a field at the Jiangpu Experimental Station (JP) in 2008, 2009, and 2011 for plant height evaluation. The trials were performed in a randomized complete block design, each with two replicates. Each plot of the trials had two 1.5 m long rows spaced 0.5 m apart. Thirty seeds were sown in each row. At the two-leaf stage, 10 evenly distributed plants were retained in each row for further growth. Field management followed common wheat production practices. The main culm height of 10 plants per plot was measured at maturity.

Leaf dimension and chlorophyll content measurements

Plants used for leaf dimension measurements were grown in a field at JP. Each line was grown in 15 1.5-m long rows spaced 27cm apart. At flowering stage, the length and width of the flag leaf, and the first, second, and third leaf below the flag leaf on the main culm were measured. Ten plants were sampled from each line. Leaf area was estimated using the following formula: area=leaf length×leaf width×0.835 (Miralles and Slafer, 1991).

The plants for relative chlorophyll content measurement were grown in two locations, one in a field at JP and one in a field on the Nanjing Agricultural University campus. SPAD readings, a measure of relative leaf chlorophyll content (Kariya et al., 1982), were recorded using a portable Minolta SPAD-502 chlorophyll meter (Spectrum Technologies, Inc., IL, USA) from 9 to 10 o’clock on sunny days. Ten plants at flowering stage were sampled from each line. Five readings were obtained from the proximal to distal end of a fully expanded leaf blade adjacent to the main leaf vein. Mean values were calculated and used as the data entry value.

Epidermal cell observation

Samples were collected from plants at flowering stage. Tissue coating and microscopic observations were conducted following the procedures described in Wu et al. (2011). Photographs were taken using an Eclipse 80i microscope (NIKON Japan) at ×100 magnification. The length and width of 20 cells for each tissue were measured with an eyepiece graticule at ×200 magnification.

Genomic DNA isolation and amplification

Genomic DNA was extracted from young wheat leaves according to the procedures described by Ma and Sorrells (1995). Bacterial artificial chromosome (BAC) clones carrying Rht-1 genes were obtained by screening a T. aestivum cv. Wangshuibai (2n=6x=42) BAC library representing three haploid genome equivalents using forward primer 5ʹ-CGGAACCGAGGCAAGCAAAAGC-3ʹ, and reverse primer 5ʹ-AAGGGGGCTTAGGAGGAGTTTTACG-3, which amplified all three Rht-1 genes. Positive BAC clones were fingerprinted with restriction enzymes BamHI and XhoI that produced fingerprints distinguishing the three Rht-1 genes based on published sequences (Peng et al., 1999; Febrer et al., 2009). Rht-A1 and Rht-D1 promoter regions were obtained by subcloning the BACs and end sequencing. Rht-B1b and Rht-D1b diagnostic assays followed Ellis et al. (2002), and Rht8 detection was conducted by surveying with the linked microsatellite marker Xgwm261 (Schmidt et al., 2004).

KOD-FX DNA polymerase was employed for PCR amplification (Toyobo, Shanghai, China). Target bands were excised from agarose gels and, following purification, were ligated into the pTA-2 vector (Toyobo) for transformation into competent DH5α cells.

RT–PCR and cDNA cloning

RNA was extracted using Trizol reagent (Invitrogen) following the manufacturer’s protocol, and quantified with Ultrospec 2100 pro (GE Healthcare Biosciences). RNA samples were treated with DNase I (Fermentas) for genomic DNA removal. The RNA sample for 3ʹ-rapid amplification of cDNA ends (RACE) was reverse-transcribed with ThermoScript reverse transcriptase (Invitrogen). RACE PCR was conducted using KOD-FX DNA polymerase (Toyobo) following the procedures described in the SMART™ RACE cDNA Amplification manual (Clontech). The gene-specific primer was BF (Ellis et al., 2002), and the nested primer was 5ʹ-ATGAAGCGGGAGTACCAGGACG-3ʹ.

RNA for reverse transcription–PCR (RT–PCR) was reverse-transcribed with M-MLV reverse transcriptase (Invitrogen). Rht-A1, Rht-B1, and Rht-D1 primers are provided in Supplementar Fig. S2 at JXB online, which were designed based on 3ʹ-untranslated region (UTR) sequences (Supplementary Fig. S2). Rht-B1cS primers were 5ʹ-TGTCGGGGTCGGCGGCTGC-3ʹ and 5ʹ-TCTCTT GTTCCACCAAGGAGAAGGGC-3ʹ. RT–PCR primers for other genes amplified in this study are listed in Supplementary Table S3, and were designed according to expressed sequence tags (ESTs; in the NCBI database) from wheat homologues of the corresponding genes. The wheat α-tubulin gene was amplified as a control to quantify expression levels using primers 5ʹ-ATCTCCAACTCCACCAGTGTCG-3ʹ and 5ʹ-TCATCGCCCTC ATCACCGTC-3ʹ.

Semi-quantitative RT–PCR was performed in a 25 μl total reaction volume. The PCR profile was as follows: 94 °C for 3min; 20–30 cycles at 94 °C for 20 s, 60 °C for 20 s, and 72 °C for 20 s; and a final 5min extension at 72 °C. A 6 μl aliquot from the PCR product of each reaction was electrophoresed on a 2% agarose gel, and viewed under UV light after standard staining with ethidium bromide.

Quantitative RT–PCR amplification was performed using the StepOne™ Real-time PCR instrument (Applied Biosystems) in a 20 μl volume containing 10 μl of SYBR-Green PCR Mastermix (Toyoba), and 10 pmol of each primer. Reactions were conducted in triplicate. Cycle threshold values for each target gene were normalized based on values obtained in corresponding reactions for the wheat α-tubulin gene. Relative expression was estimated employing the 2–ΔΔCT method (Livak and Schmittgen, 2001).

Transient assays

Constructs

The 2.1kb and 2.3kb Rht-B1a and Rht-B1c upstream sequences were amplified using KOD-FX DNA polymerase (Toyobo) with the forward primer containing a PstI restriction site, and the reverse primer containing a SmaI restriction site. The restricted amplicons were used to substitute for the Cauliflower mosaic virus (CaMV) 35S promoter in the CaMV35S–GUS–NOS sequence of a pUC19-based expression vector. Following ligation and transformation into DH5α competent cells, the constructs were confirmed by sequencing.

Transient expression and microscopy

The PDS-1000 particle delivery system (Bio-Rad) was used to introduce the fusion constructs through 1.1mm tungsten particles into epidermal cells of onion (Allium cepa) bulb scales or maize (Zea mays) shoot tips placed on half-strength MS medium agar plates following the manufacturer’s recommended procedure. Maize shoot tips were from seedlings 2–3 d following germination at 25 °C. Bombardment parameters were 1100 psi of pressure, 85mm from the macrocarrier to the samples, and 28 in of mercury vacuum. After bombardment, tissues were incubated in the dark for 12h at 25 °C, and subsequently stained in X-gluc solution for 4h following Jefferson (1987). β-Glucuronidase (GUS) expression was detected under light microscopy (Nikon ECLIPSE 80i).

GA treatment

Seeds were germinated in Petri dishes at room temperature. One-week-old seedlings grown under 16h of light per day were sprayed with 100 μM GA3, and were harvested 1h later for RNA extraction. Samples without GA3 spraying were harvested as controls.

Sequence analysis

Positive clones were sequenced at Takara Bio, Inc., Dalian, China. Basic sequence analysis was conducted with Macvector 10.0 (Accelrys, Oxford, USA). The transcription start site (TSS) was predicted with the plant promoter prediction program TSSP (http://www.softberry.com). Putative cis-acting elements were identified using the PLACE database of Higo et al. (1999).

Statistical analysis

Comparison of quantitative data between samples was subjected to two-tailed and unpaired Student’s t-test.

Accession numbers

The sequence data of Rht-A1a, Rht-B1a, Rht-B1c, and Rht-D1a were deposited in NCBI (http://www.ncbi.nlm.nih.gov) under accession nos KC767924–KC767927.

Results

Effects of Rht-B1c on plant growth and development

Rht-1 genes are key regulators of the GA signalling pathway, which has multiple effects on plant growth and development. Therefore, the Rht-B1c near-isogenic lines D-Sumai 3 (DS3) and D-Yangmai 3 (DY3) were compared with their recurrent parents Sumai 3 (S3) and Yangmai 3 (Y3) for plant height, leaf morphology, epidermal cell length and width, and chlorophyll content.

DS3 and DY3 were dwarfed at the early seedling stage (Fig. 1A, B). At maturity, S3 was ~15–20cm taller than Y3, but DS3 and DY3 were similar in plant height and exhibited an ~50% reduction relative to S3 and Y3 (Fig. 1C–E), compared with a 10–20% height reduction by Rht-B1b and Rht-D1b (Flintham et al., 1997). In the BC23F1:2 families, the plants could be distinctly classified into three groups (a, b, and c; Supplementary Table S4 at JXB online) based on plant height. Groups a and c were similar to S3/Y3 and DS3/DY3, respectively, in plant height, and represented the parental genotypes. Group b exhibited plant height values intermediate to the parental values, suggesting they were heterozygous at Rht-B1. Group b with the Y3 background showed a mean plant height close to the mid-parent values, and the dominant effect mean was nearly zero (Supplementary Table S4), suggesting that Rht-B1c in this background was co-dominant in conditioning plant height. However, the Rht-B1c dominant effects in the three BC23F1:2 families with the S3 background were all negative, with a mean of –5.5 (Supplementary Table S4), indicating partial dominance for reduced height. Consistent with these results, Flintham et al. (1997) reported that the Rht-B1c dominant effects in three different genetic backgrounds varied from –4.4 to –8.5, with a mean of –6.1. However, Rht-B1b and Rht-D1b were either co-dominant or partially recessive for reduced height (Gale et al., 1989; Keyes and Sorrells, 1989; Flintham et al., 1997; Flintham and Gale, 1998).

Fig. 1.

Fig. 1.

Open in a new tab

Phenotypes of Rht-B1c near-isogenic lines. (A and C) Phenotypes of S3 (left) and DS3 (right). (B and D) Phenotypes of Y3 (left) and DY3 (right). (E) Plant height (cm) of the three genotypic groups in S3 and Y3 backgrounds, calculated with pooled data from BC23F2 populations. Error bars indicate the standard deviation. The sample sizes of the two populations were 80 (S3) and 163 (Y3) plants, and the three groups were significantly different from each other in plant height, with P < 0.0001, and statistically fit to a 1:2:1 genotypic ratio (B1a/B1a:B1a/B1c:B1c/B1c). Genotypic classification was verified by a F2:3 progeny test. (This figure is available in colour at JXB online.)

Rht-B1c introduction resulted in significantly shorter and wider leaves at all leaf positions (P < 0.0001). The effects increased as the position approached the spike in the S3 background plants (Fig. 2A); however, in Y3 background individuals, effects on the flag leaf morphology were not as prominent as on the second upper leaves (Fig. 2B). The length and width changes consequently resulted in leaf area variation. DS3 and DY3 flag leaf area was reduced by 8.7% and 4.9%, respectively; both were significantly different (P < 0.0001) from the taller counterparts, namely S3 and Y3. Moreover, applying SPAD units as measurements, it was found that the relative chlorophyll contents in the flag leaf and the leaf directly below the flag leaf were significantly higher in DS3 and DY3 than in S3 and Y3, with P < 0.0001 (Fig. 3). This is consistent with darker green leaves observed in plants with Rht-B1c.

Fig. 2.

Fig. 2.

Open in a new tab

Percentage leaf length reduction and leaf width increase in (A) DS3 and (B) DY3 relative to S3 and Y3. Numbers 1–4 refers to the flag leaf, and the first, second, and third leaf below the flag leaf.

Fig. 3.

Fig. 3.

Open in a new tab

SPAD measurements. (1) Flag leaf; (2) first leaf below the flag leaf for S3, DS3 Y3, and DY3. JP and PL are the field locations in which the plants grew. Ten plants were measured for each line at each location. Error bars indicate the standard deviation. The difference in each leaf between S3 and DS3 and between Y3 and DY3 were significant at P < 0.0001.

Rht-B1c effects on cell morphology were examined through microscopic observation and two-dimensional measurement of the epidermal cells in the peduncle, flag leaf sheath, and blade. The results showed that the epidermal cells of all observed tissues had reduced length and increased width (Fig. 4A). Peduncle epidermal cell morphology for the two Rht-B1 genotypes is provided in Fig. 4B.

Fig. 4.

Fig. 4.

Open in a new tab

Percentage cell length reduction and width increase of epidermal cells in different tissues (A) and microscopic observation of peduncle epidermal cells (×100, bar=100 μm) at flowering stage (B). (This figure is available in colour at JXB online.)

Rht-B1c differs from Rht-B1a by fragment insertions and SNPs in promoters and coding regions

The Rht-B1c allele effects on plant growth and development were distinct from those of Rht-B1b; therefore, it was of interest to determine the sequence characteristics responsible for the observed differences. The full coding Rht-1 sequences were obtained by PCR using a forward primer (5ʹ-CGGAAC CGAGGCAAGCAAAAGC-3ʹ) designed based on one EST sequence (gi55686840) that showed the best homology to the Rht-B1a 5ʹ region (Peng et al., 1997), and a reverse primer (5ʹ-AAGG GGGCTTAGGAGGAGTTTTACG-3) designed based on the 3ʹ-UTRs from a group of Rht-1 3ʹ-RACE products (data not shown). S3 genomic DNA amplification generated three products established by fingerprinting with restriction enzymes BamHI and XhoI. The Rht-B1a-like product exhibited a 67bp 5ʹ-UTR and 147bp 3ʹ-UTR (data not shown). This sequence was used to design one Rht-B-specific reverse primer (5ʹ-GCTTCTCCT TCTTCAAGAGCGTGCG-3ʹ) ending 116bp downstream from the stop codon. S3 and DS3 genomic DNA amplification using the Rht-B-specific reverse primer, and the Rht-B1-specific forward primer BF (Ellis et al., 2002), which initiated 24bp upstream from the start codon, produced an expected 2006bp Rht-B1a sequence in S3, and a 4032bp sequence in DS3. The DS3 sequence differed from Rht-B1a by a 2026bp insertion 151bp downstream from the start codon, two SNPs preceding the indel, and one SNP downstream from the indel (Fig. 5A). These sequence variations were verified by repeating the PCRs.

Fig. 5.

Fig. 5.

Open in a new tab

Sequence variations of Rht-B1c relative to Rht-B1a (A), and the relationship of Veju insertion to plant height (B). In (A), letters and dashed lines represent SNPs between Rht-B1a and Bht-B1c; numbers indicate positions of sequence variations. The positions are numbered by defining the first nucleotide of the start codon as +1. Triangles and inverted triangles represent start and stop codon positions. The small black boxes within the coding regions depict the 5bp (AGGTG) target site duplication. PPT and PBS are the polypurine tract and primer-binding site, respectively. In (B), the values at the bottom represent plant height (cm) of individual F2 plants (not all plants in Supplementary Table S4 at JXB online are provided), a, tall plant group, b, heterozygote plant group, c, dwarf plant group.

Association of the 4kb fragment with plant height was determined by surveying the DS3-derived BC23F1:2 families using one primer (5ʹ-GCCCTCTTGGCTCACTTTTCTCATT-3ʹ) located within the insertion, and one primer (5ʹ-GCTCCAG CTTCTGCGCCACGT-3ʹ) located downstream from the indel. PCR revealed three genotypes, exhibiting co-segregation with variation in plant height (Fig. 5B). A similar result was obtained in the DY3-derived BC23F1:2 families. Assuming that the wheat genome size is equivalent to 17 Gb, only ~2kb of donor DNA will be maintained in a plant derived from 23 backcrossed generations. Consequently, the variable sequence region revealed in the near-isogenic lines should correspond to the introduced Rht-B1c.

The Rht-B1c 2kb insertion was characterized by a 5bp (AGGTG) target site duplication, a 503bp long direct terminal repeat (LTR), and a 1015bp internal domain flanked by LTRs (Fig. 5A). The LTR showed TG–CA inverted terminal repeats. This type of sequence organization was similar to the terminal-repeat retrotransposon in miniature (TRIM) family termed Veju, characterized in wheat (SanMiguel et al., 2002).

Differences between Rht-B1a and Rht-B1c in the sequence upstream from the coding region were examined by isolating a 2135bp and a 2332bp upstream sequence, respectively, from S3 and DS3 via two rounds of PCR-based genome walking (O’Malley et al., 2007). Both fragments were confirmed by genomic DNA PCR amplification using gene-specific primers (5ʹ-AATTCGAACTGCACATACGTATGAAGATG-3ʹ and 5ʹ-CGCCGCCGCCGACCCAGA-3ʹ) flanking the fragments. Relative to the S3 fragment, seven SNPs, and one 197bp sequence insertion upstream from –591 in the DS3 fragment were detected (Fig. 5A). This 197bp indel also co-segregated with the height phenotypes in the BC23F2 populations (data not shown). All Rht-B1c sequence variations were confirmed in ‘Tom Thumb’, the donor parent.

Rht-B1 is more distant from Rht-A1 and Rht-D1 than Rht-A1 is from Rht-D1

The Rht-1 gene upstream sequence variations were examined by cloning Rht-A1a and Rht-D1a sequences corresponding to the Rht-B1 upstream sequence via subcloning of BACs carrying the respective genes. The respective Rht-A1a and Rht-D1a sequences, 3298bp and 4331bp, were structurally unique from Rht-B1c by an 868bp insertion and a 1902bp insertion, respectively, 728bp upstream from the 197bp indel, besides a number of small indels (Supplementary Fig. S1 at JXB online). Regardless of the large indels, Rht-B1 exhibited 89.0% and 92.8% similarity to Rht-A1 and Rht-D1, respectively, and Rht-A1 and Rht-D1 were 90.3% similar. In terms of the 197bp indel, Rht-A1 and Rht-D1 remained consistent in showing the highest similarity. A comparable relationship was observed in the coding regions.

All Rht-B1c sequence variations, except for SNP43, SNP75, and the Veju insertion, are present in Rht-1 genes on A, B, S, and G genomes

Unique Rht-B1c sequence variations relative to Rht-B1 were investigated by cloning Rht-1 sequences up to 1625bp upstream from 848 (Fig. 5A) on A, B, S, and G genomes of the Triticeae tribe by pairing the Rht-B1-specific primer 5ʹ-TTACCTCTTGTGATTCCCACCG-3ʹ or the degenerate primer 5ʹ-CAGAACCGTGTGTCGCATTAGCTA-3ʹ with primer 5ʹ-TGGCGGTGAAGTGGGCGAAC-3ʹ. The 22 examined accessions representing different Triticeae species or subspecies and their genome compositions are listed in Supplementary Table S2 at JXB online. Nucleotide C at SNP43 and nucleotide A at SNP75, as well as the Veju insertion, did not occur in any of these accessions. The other SNPs in the corresponding regions were detected in Rht-1 genes in at least one of the examined genomes (Supplementary Table S2). The Rht-B1c 197bp indel was found in Rht-1 genes of all but the B genomes of tetraploid and hexaploid species, thus probably representing an ancient form of the Rht-B1 sequences, even though there are a few SNPs, and small indels among the orthologues (data not shown). It was concluded that SNP43, SNP75, and the Veju insertion were the candidates resulting in the mutation from Rht-B1a to Rht-B1c.

Rht-B1c promoter activity and Rht-1 gene cis-element variations

A TSS was identified 163bp upstream from the Rht-B1 gene start codon through plant promoter prediction with TSSP (Fig. 5A). This was confirmed by aligning the sequence with the full-length cDNA (AK332917) obtained through the CAP-trapper method of Kawaura et al. (2009). A putative TATA box was present 36bp upstream from the TSS. Similar sequence features were found in all three Rht-1 genes at the two sites, with the exception of a few indels.

The effects of the Rht-B1c upstream sequence variations on promoter activity were investigated through transient expression assays. The transient expression constructs were prepared by fusing 2.1kb Rht-B1a and 2.3kb Rht-B1c upstream sequences to the GUS reporter gene (Fig. 6A). Initially, following bombardment of the constructs into onion epidermal cells, no GUS activity was detected. However, when the onion epidermal cells were replaced with young maize shoot tips for bombardment, comparable GUS activity was detected for all constructs (Fig. 6B), implying that Rht-B1c and Rht-B1a promoters do not differ substantially in promoter activity despite the sequence variations. Quantitative PCR analysis (results shown below) showed that upstream sequence polymorphisms of all Rht-1 genes exhibited no effects on basic promoter activity.

Fig. 6.

Fig. 6.

Open in a new tab

Rht-B1 promoter-driven transient expression in maize shoot apical cells. (A) Depiction of the binary vector constructs. (B) GUS staining of the transformed shoot apical cells. (This figure is available in colour at JXB online.)

PLACE was used to identify cis-elements in the Rht-1 gene regions starting at 728bp upstream from the 197bp indel. Results showed that 84% of putative upstream regulatory elements of Rht-B1a, which has the shortest target sequence, were found in the other three genes, suggesting substantial conservation in expression regulation. However, putative cis-element evolution was also evident based on predictions. Rht-B1c lost the CACGTG motif predicted in Rht-B1a, and gained eight motifs due to the SNPs and 197bp indel. Three of these gained motifs, including CELLCYCLESC, P1BS, and S1FBOXSORPS1L21, were also not detected in Rht-A1a and Rht-D1a (Supplementary Table S5 at JXB online). With a comparable target sequence length (1593bp and 1616bp, respectively), Rht-A1a and Rht-D1a were distinguished by a total of 25 putative cis-elements (Supplementary Table S5). Consequently, the results suggested that although the basic Rht-1 gene functions in wheat are potentially redundant, their roles in growth and development might have differentiated.

The 197bp indel and coding region SNPs did not contribute to the major dwarfing effect

The influences of the 197bp promoter region insertion, the Veju insertion, and the three coding region SNPs on plant height of 126 wheat cultivars in China’s wheat germplasm pool core collection were investigated. The corresponding regions were examined by PCR amplification using Rht-B1-specific primers 5ʹ-TTACCTCTTGTGATTCCCACCG-3ʹ and 5ʹ-TGGCGGTGAAGTGGGCGAAC-3ʹ. One hundred and sixteen lines produced products identical in size to the corresponding S3 Rht-B1a sequence, indicating that the samples did not possess the 197bp and Veju insertions. These results were confirmed by sequencing products of 22 of these cultivars. The remaining 10 lines generated products longer than the S3 Rht-B1a sequence. Sequencing revealed that seven of these lines possessed the 197bp insertion and were identical to Rht-B1c at the three SNP sites, and three had a 160bp insertion upstream from –356 and were identical to Rht-B1a at these SNP sites. Consequently, the 126 cultivars were classified into three haplotypes (Fig. 7; Supplementary Table S1 at JXB online). SNP43 showed a gain of the HaeIII restriction recognition site in Rht-B1c, and SNP75 detected in Rht-B1c exhibited a loss of the BccI restriction recognition site in Rht-B1a. Therefore, it was possible to examine their presence in wheat cultivars by restriction enzyme digestion of the 123bp PCR products beginning with –24 to 99. The 94 cultivars without sequence data all produced PCR products digestible with BccI but not HaeIII, and were therefore assigned to haplotype III.

Fig. 7.

Fig. 7.

Open in a new tab

Rht-B1 haplotypes in common wheat and the corresponding average plant height (PH in cm).

Based on 3 year field trial data, plant height of the seven cultivars representing haplotype I (i.e. Rht-B1c) exhibited a 90.5cm average. However, one cultivar of this type, Caizihuang, was >150cm in height (Supplementary Table S1 at JXB online). It does not possess a known dwarfing gene. The other six cultivars, all with a plant height <100cm, carried Rht8, Rht-D1b, or dwarfing quantitative trait loci (QTLs) based on the literature, sequencing, or diagnostic assays. This suggests that instead of the 197bp insertion and three SNPs, the Veju insertion is most probably responsible for the extreme Rht-B1c dwarfing capacity. Haplotype II plant height ranged from 80.5cm to 126.4cm, with an average of 110.2cm. Although haplotype III and haplotype II differed in the absence of the 160bp insertion, they exhibited a similar average plant height (Fig. 7), suggesting that this insertion had no effect on dwarfing.

The Veju insertion changes processing and translated products of Rht-B1c transcripts

Since Rht-B1c has one 2026bp insertion and three SNPs in the region corresponding to the Rht-B1a open reading frame (ORF), Rht-B1c transcripts were isolated by amplifying the cDNA beginning at the start codon by 3ʹ-RACE. PCR generated two reproducible products. One product was 363bp in length, and was designated as Rht-B1cS; the other, 2370bp in length, was designated Rht-B1cL. Alignment of Rht-B1cS with Rht-B1a cDNA and Rht-B1c genomic DNA sequences showed that Rht-B1cS resulted from premature polyadenylation of the Rht-B1c transcripts, and it included a 150bp 5ʹ region identical to Rht-B1a, and 186bp of the 5ʹ Veju sequence. The sequences flanking the polyadenylation site contained multiple conserved polyadenylation signal elements (Fig. 8A). Rht-B1cS did not possess a stop codon. In eukaryotes, mRNA lacking a stop codon might be degraded via a non-stop decay pathway (Frischmeyer et al., 2002; van Hoof et al., 2002), which, if also applicable in plants, might explain why Rht-B1cS abundance was 40 times less than that of Rht-B1cL at the mRNA level (Fig. 8B).

Fig. 8.

Fig. 8.

Open in a new tab

Putative polyadenylation signals surrounding the Rht-B1cS polyadenylation site (underlined) (A) and relative expression levels of the two Rht-B1c transcripts in seedling stage leaves (B).

Rht-B1cL contains a 1956bp ORF and a 414bp 3ʹ-UTR. Comparison with the Rht-B1c genomic DNA sequence provided evidence that this transcript resulted from removal of the 1936bp from 149 to 2084 (Fig. 5). Relative to Rht-B1a, the Rht-B1cL ORF retained a 90bp 3ʹ sequence of the Veju insertion, which subsequently resulted in a predicted 30 residue addition following K49, residing within the DELLA motif (Fig. 9). The SNP downstream from the Veju insertion was identified as a nonsense mutation, but the two SNPs before the Veju insertion result in G to R and M to I changes, both of which reside in non-conserved regions (Fig. 9).With the exception of these variations, the predicted Rht-B1cL translation product was identical to Rht-B1a.

Fig. 9.

Fig. 9.

Open in a new tab

N-terminal comparisons of the Rht-B1cL product with other plant DELLA proteins. Arrows indicate the mutated amino acids in the Rht-B1cL product. HvSLN1 (Hordeum vulgare, Q8W127), OsSLR1 (Oryza sativa, NP_001051032), ZmD9 (Zea mays, ABI84225), AtGAI (Arabidopsis thaliana, NP_172945), BrRGA2 (Brassica rapa, Q5BN22), CmGAIP (Cucurbita maxima, Q6EI06), MdRGL1a (Malus×domestica, AAY56751), GmGAI1 (Glycine max, NP_001240948), SlGAI (Solanum lycopersicum, NP_001234365), and VvGAI1 (Vitis vinifera, XP_002284648).

Rht-B1c reduced Rht-1 gene expression levels but not the transcriptional response to GA treatment

Since Rht-B1cL was the major Rht-B1c transcript in cells, its abundance was examined to determine Rht-B1c mRNA levels in the following experiments. Semi-quantitative RT–PCR assays indicated that Rht-B1 was primarily expressed in stems and leaves, and lower expression levels were found in roots and milky stage spikes (Fig. 10A). Overall, the Rht-B1 expression level was consistently lower in DS3 than in S3, irrespective of growth stage (Fig. 10A, B). Seedling treatment with 100 μM GA3 resulted in notable Rht-B1a and Rht-B1c up-regulation, but the Rht-B1c transcription levels remained lower (Fig. 10B). However, the transcription level of Rht-B1c could be higher taking into consideration the Rht-B1cS transcript.

Fig. 10.

Fig. 10.

Open in a new tab

Rht-1 expression profiling. (A) Rht-B1 expression in different tissues revealed through semi-quantitative RT–PCR. (B) Rht-1 gene expression following GA treatment revealed via real-time PCR. In A, RNA was isolated from roots, stems, leaves, and spikes of wheat seedlings at the milk stage. In B, total RNA was isolated from 10-day-old seedlings before and 1h after spraying with 100 μM GA3. The experiment was repeated three times. The wheat α-tubulin gene was amplified as a control to quantify expression levels.

Rht-A1a and Rht-D1a transcript levels and their responses to GA3 treatment in DS3 and S3 plants were also investigated (Fig. 10B). Both genes were expressed at a lower level in DS3 than in S3. Similar to Rht-B1 genes, Rht-A1a and Rht-D1a expression was up-regulated in DS3 and S3 following the GA3 application; however, the expression levels still remained lower in DS3. These results provided evidence that Rht-B1c decreased the transcript abundance of Rht-1 genes, but did not affect induction of expression by GA.

Rht-B1c up regulated genes involved in cell elongation, chlorophyll biosynthesis, and chloroplast development

As Rht-B1c plant lines exhibited shorter cells and higher chlorophyll content (Fig. 3), S3 and DS3 seedling leaves were evaluated to determine if Rht-B1c regulated cell elongation and chlorophyll biosynthesis through GATA-like transcription factors, similar to what was reported in A. thaliana (Richter et al., 2010; Hudson et al., 2011). Quantitative RT–PCR for GATA22, the wheat GATA-like transcription factor gene, showed that DS3 expressed almost three times more of this gene relative to S3 (Fig. 11A). GA3 application decreased its expression in both lines; nevertheless, the expression level remained notably higher in DS3 than in S3. At the tillering stage, DS3 still maintained a higher GATA22 expression level (Fig. 11B). The expression of four wheat genes homologous to HEMA1, GUN4, CAO1, and PorB was also examined, and they all had higher expression levels in DS3 (Fig. 11B). These four genes were positively regulated for chlorophyll biosynthesis by GATA transcription factors in A. thaliana (Hudson et al. 2011).

Fig. 11.

Fig. 11.

Open in a new tab

Expression profiling of genes putatively regulated by DELLA proteins. (A) GATA22 in 1-week-old S3 and DS3 seedlings following GA3 treatment. (B) HEMA1, GUN4, CAO1, PorB, CAB, and rbcS in tillering stage leaves of S3 and DS3.

Chiang et al. (2012) demonstrated that GATA transcription factors modulated chloroplast biogenesis. Therefore, to determine if Rht-B1c affected chloroplast development and photosynthesis, the expression of rbcS and CAB genes was measured for chlorophyll a/b binding. rbcS showed an almost 6-fold increase in the expression level, and the CAB genes also displayed a significantly higher level in DS3 (Fig. 11B).

Discussion

Sequence characteristics of Rht-B1c and its homologues

Pearce et al. (2011) and Wu et al. (2011) reported Rht-B1c cDNA cloning. Wu et al. (2011) also isolated this gene’s coding region genomic DNA. In the present study, using homology-based cloning and genome walking strategies, the full genomic DNA sequence of Rht-B1c and Rht-B1a, including both the coding region and the upstream 2kb sequence, was cloned. In agreement with Wu et al. (2011), the present results revealed that in the coding region, Rht-B1c differed from Rht-B1a by one 2026bp insertion 151bp downstream from the start codon. The retrotransposon was determined to be different from Veju_S and Veju_L elements (SanMiguel et al., 2002; Sabot et al., 2005). Furthermore, it was found that the Rht-B1c coding region carried three SNPs, with SNP43 and SNP75 preceding the insertion, and SNP723 following the insertion. This was consistent with the findings of Pearce et al. (2011). However, the SNP A617 reported by Pearce et al. (2011) was not confirmed. SNP723 does not cause a codon change. In the 2kb upstream region, Rht-B1c was characterized by one 197bp insertion preceding –591, and seven SNPs (Fig. 5A). Genetic analyses showed that the insertions co-segregated with plant height in segregating populations.

Among the insertions and SNPs, the Veju insertion was the only one not detected in the Rht-B1 genes of all 126 surveyed common wheat cultivars, and in the Rht-1 orthologues present in the A, B, S, and G genomes of the Triticeae species, the close relatives of wheat. Therefore, unlike most identified DELLA mutants resulting from deletion (gai), premature termination (rga-1, Rht-B1b, Rht-B1d, Rht-B1e, Rht-D1b, and slr1), or amino acid substitutions (rga-2, sln1d, and Brrga1-d) (Peng et al., 1997, 1999; Silverstone et al., 1998; Pearce et al., 2011; Ikeda et al., 2001; Chandler et al., 2002; Muangprom et al., 2005), the Veju insertion clearly emerged as the mutation responsible for the origin of the Rht-B1c allele from Rht-B1.

SNPs C43 and A75 in Rht-B1c caused a codon change, but an association with dwarfing was not observed. Furthermore, the substituted amino acids reside in a non-conserved region, which is not implicated in the DELLA–GID1 interaction (Murase et al., 2008), suggesting no crucial functional relationship. SNPs C43 and A75 were identified in seven of the 126 cultivars with SNP G723 and the 197bp upstream insertion (haplotype I in Fig. 7). Sequences homologous to the 197bp insertion and SNPs T–131 and G723 were resolved in Rht-B1 orthologues in S genome carriers Aegilops speltoides and A. longissima, and G genome carriers T. timopheevii and T. araraticum. Because the S genome was viewed as the ancestor of B and G genomes (Wang et al., 1997), these results suggested that Rht-B1c originated from the Veju insertion into an ancient Rht-B1 haplotype that experienced point mutations from G43 to C43 and G75 to A75. All Rht orthologues detected in A and D genomes possessed the 197bp insertion, providing additional support for its ancestral origin. The 197bp insertion, together with the seven SNPs detected in the Rht-B1c upstream region, appeared to exhibit negligible effects on basic promoter activity. In common wheat, haplotype III (Figure 7) was the most common haplotype, differing from the S genome orthologs by the G723 to A723 mutation in the coding region and the loss of 197-bp fragment. All detected tetraploid species and hexaploid sub-species of wheat examined in our study belonged to this haplotype. Haplotype II, the most rare haplotype, might have originated from haplotype III by insertion of the 160-bp fragment.

Compared with the DNA coding region, the upstream regions of Rht-1 genes in various genomes were much more diverse, characterized by a number of indels and SNPs. Generally speaking, Rht-A1 and Rht-D1 were more similar relative to Rht-B1. Sequence variations in the upstream regions of these genes caused loss or gain of the predicted cis-elements. In A. thaliana, five DELLA protein genes have been reported with overlapping but differentiated functions (Dill and Sun, 2001; Tyler et al., 2004). Gallego-Bartolomé et al. (2010) suggested that this functional diversification largely relied on changes in gene expression patterns rather than on the capacity of the protein to interact with different regulatory partners. It would be interesting to ascertain if upstream sequence variations in Rht-1 genes caused functional diversification. The polyploid nature of wheat could surely allow such modifications.

The influence of Veju insertion on Rht-B1c expression and function

Two Rht-B1c transcripts were identified, termed Rht-B1cS and Rht-B1cL. Rht-B1cL is identical to the Rht-B1c cDNA reported by Pearce et al. (2011) and Wu et al. (2011). The transcript encodes a protein with 30 extra residues within the N-terminal DELLA motif, due to retention of the 90bp 3ʹ Veju sequence in mature mRNA. Disruption of the DELLA motif could cause failure of its interaction with the GA receptor GID1, which is required for GA–GID1–DELLA complex formation underlying GA signal transduction (Murase et al., 2008). Wu et al. (2011) found that rice plants transformed with Rht-B1c cDNA were much shorter than those transformed with Rht-B1b.

Rht-B1cS contained only the 336bp 5ʹ region initiating at the start codon. The multiple conserved polyadenylation signal elements surrounding the putative Rht-B1cS polyadenylation site and the presence of two Rht-B1c transcripts in cells suggested that Rht-B1cS resulted from alternative polyadenylation during RNA processing. Real-time RT–PCR analysis revealed that Rht-B1cL accounted for ~97.5% of the Rht-B1c transcripts in seedling leaves. However, this proportion might actually be much less since Rht-B1cS probably has a short lifetime due to the absence of a stop codon. The mRNA without a stop codon in Saccharomyces cerevisiae would trigger a non-stop decay pathway (van Hoof et al., 2002; Frischmeyer et al., 2002, Vasudevan et al., 2002). Indeed, Rht-B1c transcripts in DS3 were substantially less than Rht-B1a transcripts in S3, regardless of tissue types. If Rht-B1a in S3 and Rht-B1c in DS3 are assumed to have comparable expression levels, generation of short-lived Rht-B1cS would result in less Rht-B1cL. The truncated polypeptide translated from Rht-B1cS possessed a truncated DELLA motif, and therefore probably does not serve a major role in GA signalling.

Since the aberrant Rht-B1c protein cannot interact with GID1 (Pearce et al. 2011; Wu et al. 2011), it would not be degraded through the GA-stimulated GID1–DELLA–SCFGID2 complex, and consequently would accumulate in cells. It is proposed that this could in turn down-regulate Rht-1 gene expression through a feedback mechanism. When external GA3 is applied, DELLA protein degradation encoded by Rht-A1a and Rht-D1a could provide the induction signal for Rht-1 gene expression. However, this induction was limited compared with S3 due to the aberrant Rht-B1c protein.

Plant morphogenesis affected by Rht-B1c

Rht-1 genes were first known for their role in determining plant height. Rht-1 mutants varied in their genetic and physiological effects on plant growth and development. In contrast to the co-dominant or partially recessive Rht-B1b and Rht-D1b, Rht-B1c was co-dominant or partially dominant for reduced height, as shown in this study and others (Uddin and Marshall, 1989; Flintham et al., 1997; Jošt et al., 2011). This was in disagreement with the report of Wu et al. (2011), in which Rht-B1c was claimed to be completely dominant. Different from Rht-B1b and Rht-D1b that reduced cell length in vegetative organs and had no affect on cell width or cell division (Miralles et al., 1998), Rht-B1c notably reduced cell length, and substantially increased cell width.

It has become clear that Rht-1 genes encode GA signalling proteins critical for GA-regulated growth and development. Using GA-deficient and altered GA-response A. thaliana mutants, Cowling and Harberd (1999) demonstrated that GA regulated cell elongation in hypocotyls grown under light and dark conditions. GAI and RGA signalling elements, both of which are DELLA proteins, were integral in these processes (Alabadí et al., 2004). DELLAs block the transcriptional activity of phytochrome-interacting factors (PIF) by binding the DNA recognition domains in PIFs (de Lucas et al., 2008; Feng et al., 2008). PIF factors, including PIF3, PIF4, and PIF5, positively regulate hypocotyl elongation, and are antagonistically regulated by light and GA (Ni et al., 1998; Kim et al., 2003; de Lucas et al., 2008; Feng et al., 2008; Kunihiro et al., 2011). Based on the GA signalling ‘relief of repression’ regulatory model, degradation failure in the aberrant Rht-B1c protein could result in a constitutive block of PIF-like transcription factor activity in wheat, subsequently resulting in shorter cells and dwarfing. Richter et al. (2010) identified two homologous GATA-type transcription factors, GNC and GNL, in A. thaliana that functioned downstream from DELLA and PIF regulators in germination, greening, elongation growth, and flowering time regulation. Their results indicated that DELLA proteins provided positive regulation of GATA-type transcription factors by inhibiting PIFs that are GNC repressors. In agreement with these observations, it was shown here that GATA22 expression in Rht-B1a and Rht-B1c lines was significantly reduced by GA application. Nevertheless, relative to the Rht-B1a lines, the GATA22 expression in the Rht-B1c line consistently maintained a markedly higher expression level, even following GA application, suggesting that PIF-like regulators could remain partially inhibited even after destabilization of the normal Rht-A1a and Rht-D1a proteins by GA treatment. The observations regarding the GATA22 expression levels resulting from GA treatment are analogous to the heterozygous state of Rht-B1c in terms of Rht cellular protein composition, which underlies the partial dominance nature of this gene. This was also consistent with the observation that increased accumulation of the mutated Rht-D1 protein exhibited stronger dwarfing effects (Li et al., 2012).

Rht-1 gene participation in chlorophyll biosynthesis and regulation of chloroplast biogenesis

Leaf is the main photosynthetic organ, and efficient photosynthesis is critical for photosynthate formation and yield. A number of experiments suggested that GA-mediated signalling participates in chlorophyll biosynthesis and chloroplast biogenesis regulatory processes. For example, both DS3 and DY3 wheat had higher chlorophyll content; a semi-dwarf near-isogenic line with Rht-B1b and Rht-D1b had significantly higher leaf photosynthetic rates, and chlorophyll and Rubisco concentrations compared with the tall wheat (Morgan et al. 1990). Feng et al. (2008) and Cheminant et al. (2011) demonstrated that DELLA proteins played a critical role in regulating chlorophyll and carotenoid biosynthetic pathways by PIF interactions. Several authors indicated that PIF factors were inhibitory to chlorophyll biosynthesis (Kim et al., 2003; Feng et al., 2008; Moon et al., 2008; Shin et al., 2009; Stephenson et al., 2009; Cheminant et al., 2011). In this process, GATA-type transcription factors were involved and functioned downstream from PIFs (Richter et al., 2010; Hudson et al., 2011). Thus, it was not surprising to find that Rht-B1c exhibited a positive regulation of GATA-type transcription factors and chlorophyll biosynthesis-related genes, such as HEMA1, GUN4, CAO1, and PorB.

Rht-B1c participated in the regulation of chloroplast biogenesis, in addition to chlorophyll biosynthesis modulation, as shown in the distinct rbcS and CAB expression patterns between Rht-B1a and Rht-B1c lines, and reported by Jiang et al. (2012). In A. thaliana this regulatory process also required GATA-type regulators (Chiang et al., 2012) and PIFs (Shin et al., 2009). In the pifQ A. thaliana mutant grown under dark conditions, both chlorophyll biosynthetic and photosynthetic genes were highly overexpressed (Shin et al., 2009). In agreement with these findings, in GA-deficient/sensitive mutants, such as ga1-3, the expression of both chlorophyll biosynthetic and photosynthetic genes was de-repressed (Alabadí et al., 2004; Cheminant et al., 2011).

Crop dwarfing was a critical factor leading to the first ‘Green revolution’. It is related to a comprehensive suite of traits, including ideal plant type, harvest index, photosynthetic efficiency, and assimilate partition, among other attributes, which are all determining factors in increasing yield potential. Pearce et al. (2011), Wu et al. (2011), and the present study clearly demonstrated that Rht-B1c, as a Rht-B1a mutant unique among other known DELLA protein gene mutants, affects many aspects of growth and development. Characterization of its structure and regulation of growth- and development-related genes will be extremely valuable in designing new crop improvement strategies. However, this is still not sufficient, since in monocots, that include most food staple cereal crops, only one DELLA protein gene locus per genome has been identified, the mechanisms underlying the GA-mediated regulation and cross-talk of GA signalling with other regulatory pathways and environmental cues, particularly the coordinative control of various physiological functions, remains poorly understood. Therefore, isolation and characterization of genes downstream from the DELLA regulator, and elucidation of their interplay with DELLA protein genes, is our next challenge.

Supplementary data

Supplementary data are available at JXB online

Figure S1. Upstream structures of Rht-1 genes.

Figure S2. 3ʹ UTR sequence alignment of Rht-1 genes.

Table S1. Rht-B1 haplotypes and plant height for 126 wheat cultivars.

Table S2. Rht-1 sequence variations in Triticeae tribe species.

Table S3. Primer sequences used in RT–PCR of downstream genes.

Table S4. Plant height of the three Rht-B1 genotypes in the individual BC23F1:2 families and the estimated dominant effect

Table S5. Distinct cis-elements predicted for wheat Rht-1 genes.

Supplementary Data
supp_64_11_3299__index.html (1.4KB, html)

Acknowledgements

This work was partially supported by the National Science Foundation of China (grant no. 30025030), the Ministry of Science and Technology (grant nos 2002CCA01300 and J00-A-006-1), fundamental research funds for the Central Universities (KYZ201202-4), and 111 project of the Ministry of Education (grant no. B08025).

References

  1. Alabadí D, Gil J, Blázquez MA, García-Martínez JL. 2004. Gibberellins repress photomorphogenesis in darkness. Plant Physiology 134, 1050–1057 [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Boss PK, Thomas MR. 2002. Association of dwarfism and floral induction with a grape ‘green revolution’ mutation. Nature 416, 847–850 [DOI] [PubMed] [Google Scholar]
  3. Chandler P, Marion-Poll A, Ellis M, Gubler F. 2002. Mutants at the slender1 locus of barley cv Himalaya. Molecular and physiological characterization. Plant Physiology 129, 181–190 [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Cheminant S, Wild M, Bouvier F, Pelletier S, Renou JP, Erhardt M, Hayes S, Terry MJ, Genschik P, Achard P. 2011. DELLAs regulate chlorophyll and carotenoid biosynthesis to prevent photooxidative damage during seedling deetiolation in Arabidopsis . The Plant Cell 23, 1849–1860 [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Chiang YF, Zubo YO, Tapken F, Kim FJ, Lavanway AM, Howard L, Pilon M, Kieber JJ, Schaller GE. 2012. Functional characterization of the GATA transcription factors GNC and CGA1 reveals their key role in chloroplast development, growth, and division in Arabidopsis . Plant Physiology 160, 332–348 [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Cowling R, Harberd NP. 1999. Gibberellins control Arabidopsis hypocotyl growth via regulation of cellular elongation. Journal of Experimental Botany 50, 1351–1357 [Google Scholar]
  7. Davies PJ. 1995. The plant hormones: their nature, occurrence, and functions. In: Davies PJ, ed. Plant hormones: physiology, biochemistry and molecular biology. Boston: Kluwer, 13–38 [Google Scholar]
  8. de Lucas M, Davière JM, Rodríguez-Falcón M, Pontin M, Iglesias-Pedraz JM, Lorrain S, Fankhauser C, Blázquez MA, Titarenko E, Prat S. 2008. A molecular framework for light and gibberellin control of cell elongation. Nature 451, 480–484 [DOI] [PubMed] [Google Scholar]
  9. Dill A, Sun TP. 2001. Synergistic derepression of gibberellin signaling by removing RGA and GAI function in Arabidopsis thaliana . Genetics 159, 777–785 [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Ellis MF, Rebetzke GJ, Chandler P, Bonnett D, Spielmeyer F, Richards RA. 2004. The effect of different height reducing genes on the early growth of wheat. Functional Plant Biology 31, 583–589 [DOI] [PubMed] [Google Scholar]
  11. Ellis MF, Spielmeyer F, Gale KR, Rebetzke GJ, Richards RA. 2002. ‘Perfect’ markers for the Rht-B1b and Rht-D1b dwarfing genes in wheat. Theoretical and Applied Genetics 105, 1038–1042 [DOI] [PubMed] [Google Scholar]
  12. Febrer M, Wilhelm E, Al-Kaff N, Wright J, Powell F, Bevan MF, Boulton MI. 2009. Rapid identification of the three homoeologues of the wheat dwarfing gene Rht using a novel PCR-based screen of three-dimensional BAC pools. Genome 52, 993–1000 [DOI] [PubMed] [Google Scholar]
  13. Feng SF, Martinez C, Gusmaroli G, et al. 2008. Coordinated regulation of Arabidopsis thaliana development by light and gibberellins. Nature 451, 475–479 [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Fleet CM, Sun TP. 2005. A DELLAcate balance: the role of gibberellin in plant morphogenesis. Current Opinion in Plant Biology 8, 77–85 [DOI] [PubMed] [Google Scholar]
  15. Flintham JE, Angus FJ, Gale MD. 1997. Heterosis, overdominance for grain yield and alpha-amylase activity in F1 hybrids between near-isogenic Rht dwarf and tall wheats. Journal of Agricultural Sciences 129, 371–378 [Google Scholar]
  16. Flintham JE, Gale MD. 1982. The Tom Thumb dwarfing gene, Rht3 in wheat I. Reduced preharvest sprouting damage to bread making quality. Theoretical and Applied Genetics 62, 121–126 [DOI] [PubMed] [Google Scholar]
  17. Flintham JE, Gale MD. 1983. The Tom Thumb dwarfing gene, Rht3 in wheat. II Effects on height, yield and grain quality. Theoretical andd Applied Genetics 66, 249–256 [DOI] [PubMed] [Google Scholar]
  18. Flintham JE, Gale MD. 1998. Plant height and yield components of inbred isogenic and F1 hybrid dwarf wheat. Journal of Applied Genetics 39, 73–83 [Google Scholar]
  19. Foster T, Kirk C, Jones FT, Allan AC, Espley R, Karunairetnam S, Rakonjac J. 2007. Characterisation of the DELLA subfamily in apple (Malus×domestica Borkh.). Tree Genetics and Genomes 3, 187–197 [Google Scholar]
  20. Frischmeyer PA, van Hoof A, O’Donnel K, Guerrerio AL, Parker R, Dietz FC. 2002. An mRNA surveillance mechanism that eliminates transcripts lacking termination codons. Science 295, 2258–2261 [DOI] [PubMed] [Google Scholar]
  21. Gale MD, Salter AM, Angus FJ. 1989. The effect of dwarfing genes on the expression of heterosis for grain yield in F1 hybrid wheat. In: Maluszynski M, ed. Current options for cereal improvement. Dordrecht: Kluwer Academic, 49–62 [Google Scholar]
  22. Gallego-Bartolomé J, Minguet EG, Marín JA, Prat S, Blázquez MA, Alabadí D. 2010. Transcriptional diversification and functional conservation between DELLA proteins in Arabidopsis . Molecular Biology and Evolution 27, 1247–1256 [DOI] [PubMed] [Google Scholar]
  23. Gomi K, Matsuoka M. 2003. Gibberellin signalling pathway. Current Opinion in Plant Biology 6, 489–493 [DOI] [PubMed] [Google Scholar]
  24. Hedden P. 2003. The genes of the Green Revolution. Trends in Genetics 19, 5–9 [DOI] [PubMed] [Google Scholar]
  25. Higo K, Ugawa Y, Iwamoto M, Korenaga T. 1999. Plant cis-acting regulatory DNA elements (PLACE) database: 1999. Nucleic Acids Research 27, 297–300 [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. Hudson D, Guevara D, Yaish MF, Hannam C, Lon N, Clarke JD, Bi YM, Rothstein SJ. 2011. GNC and CGA1 modulate chlorophyll biosynthesis and glutamate synthase (GLU1/Fd-GOGAT) expression in Arabidopsis . PLoS One 6, e26765. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Ikeda A, Ueguchi-Tanaka M, Sonoda Y, Kitano F, Koshioka M, Futsuhara Y, Matsuoka M, Yamaguchi J. 2001. slender rice, a constitutive gibberellin response mutant is caused by a null mutation of the SLR1 gene, an orthologue of the height-regulating gene GAI/RGA/RHT/D8 . The Plant Cell 13, 999–1010 [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Itoh F, Ueguchi-Tanaka M, Sato YY, Ashikari M, Matsuoka M. 2002. The gibberellin signaling pathway is regulated by the appearance and disappearance of SLENDER RICE1 in nuclei. The Plant Cell 14, 57–70 [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Jefferson RA. 1987. Assaying chimeric genes in plants: the GUS gene fusion system. Plant Molecular Biology Reporter 5, 387–405 [Google Scholar]
  30. Jiang X, Li F, Wang T, Peng C, Wang F, Wu F, Wang X. 2012. Gibberellin indirectly promotes chloroplast biogenesis as a means to maintain the chloroplast population of expanded cells. The Plant Journal 72, 768–780 [DOI] [PubMed] [Google Scholar]
  31. Jošt M, Samobor V, Srečec S. 2001. Pleiotropic effect of Rht3 dwarfing gene on some traits of wheat (Tr. aestivum L. em Thell). Poljoprivreda (Osijek) 7, 10–16 [Google Scholar]
  32. Kariya K, Matsuzaki A, Machida F. 1982. Distribution of chlorophyll content in leaf blade of rice plant. Japanese Journal of Crop Science 51, 134–135 [Google Scholar]
  33. Kawaura K, Mochida K, Enju A, et al. 2009. Assessment of adaptive evolution between wheat and rice as deduced from full-length common wheat cDNA sequence data and expression patterns. BMC Genomics 10, 271–281 [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Keyes G, Sorrells ME. 1989. Rht1 and Rht2 semidwarf genes effect on hybrid vigor and agronomic traits of wheat. Crop Science 29, 1442–1447 [Google Scholar]
  35. Kim JY, Yi FK, Choi G, Shin B, Son PS, Choi GS. 2003. Functional characterization of phytochrome interacting factor 3 in phytochrome-mediated light signal transduction. The Plant Cell 15, 2399–2407 [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Konzak CF. 1988. Genetic analysis, genetic improvement and evaluation of induced semi-dwarf mutants in wheat. In: Semi-dwarf cereal mutants and their use in cross-breeding III research coordination meeting, 16–20 December 1985. International Atomic Energy Agency, Vienna, Austria, 39–50 [Google Scholar]
  37. Kunihiro A, Yamashino T, Nakamichi N, Niwa Y, Nakanishi F, Mizuno T. 2011. Phytochrome-interacting factor 4 and 5 (PIF4 and PIF5) activate the homeobox ATHB2 and auxin-inducible IAA29 genes in the coincidence mechanism underlying photoperiodic control of plant growth of Arabidopsis thaliana . Plant and Cell Physiology 52, 1315–1329 [DOI] [PubMed] [Google Scholar]
  38. Lawit SJ, Wych FM, Xu D, Kundu S, Tomes DT. 2010. Maize DELLA proteins dwarf plant8 and dwarf plant9 as modulators of plant development. Plant and Cell Physiology 51, 1854–1868 [DOI] [PubMed] [Google Scholar]
  39. Li XP, Lan SQ, Feng YR. 2006. Effects on agronomic chracteristics of Rht-B1b, Rht-D1b, Rht-B1c wheat dwarfing genes and theirs combinations in artificial climate chamber. Journal of Hebei Agricultural Sciences 10, 14–18 [Google Scholar]
  40. Li Y, Xiao J, Wu J, et al. 2012. A tandem segmental duplication (TSD) in green revolution gene Rht-D1b region underlies plant height variation. New Phytologist 196, 282–291 [DOI] [PubMed] [Google Scholar]
  41. Livak KJ, Schmittgen TD. 2001. Analysis of relative gene expression data using real-time quantitative PCR and the 2–ΔΔCT method. Methods 25, 402–408 [DOI] [PubMed] [Google Scholar]
  42. Ma ZQ, Sorrells ME. 1995. Genetic analysis of fertility restoration in wheat using restriction fragment length polymorphisms. Crop Science 3, 1137–1143 [Google Scholar]
  43. Miralles DJ, Katz SD, Colloca A, Slafer GA. 1998. Floret development in near isogenic wheat lines differing in plant height. Field Crops Research 59, 21–30 [Google Scholar]
  44. Miralles DJ, Slafer GA. 1991. A simple model for non-destructive estimates of leaf area in wheat. Cereal Research Communications 19, 439–444 [Google Scholar]
  45. Moon J, Zhu L, Shen F, Huq E. 2008. PIF1 directly and indirectly regulates chlorophyll biosynthesis to optimize the greening process in Arabidopsis . Proceedings of the National Academy of Sciences, USA 105, 9433–9438 [DOI] [PMC free article] [PubMed] [Google Scholar]
  46. Morgan JA, LeCain DR, Wells R. 1990. Semidwarfing genes concentrate photosynthetic machinery and affect leaf gas exchange of wheat. Crop Science 30, 602–608 [Google Scholar]
  47. Muangprom A, Thomas SG, Sun TP, Osborn TC. 2005. A novel dwarfing mutation in a green revolution gene from Brassica rapa . Plant Physiology 137, 931–938 [DOI] [PMC free article] [PubMed] [Google Scholar]
  48. Murase K, Hirano Y, Sun TP, Hakoshima T. 2008. Gibberellin-induced DELLA recognition by the gibberellin receptor GID1. Nature 456, 459–463 [DOI] [PubMed] [Google Scholar]
  49. Ni M, Tepperman J.M, Quail PH. 1998. PIF3, a phytochrome-interacting factor necessary for normal photoinduced signal transduction, is a novel basic helix–loop–helix protein. Cell 95, 657–667 [DOI] [PubMed] [Google Scholar]
  50. O’Malley RC, Alonso JM, Kim CJ, Leisse TJ, Ecker JR. 2007. An adapter ligation-mediated PCR method for high-throughput mapping of T-DNA inserts in the Arabidopsis genome. Nature Protocols 2, 2910–2917 [DOI] [PubMed] [Google Scholar]
  51. Pearce S, Saville R, Vaughan SP, et al. 2011. Molecular characterisation of Rht1 dwarfing genes in hexaploid wheat. Plant Physiology 157, 1820–1831 [DOI] [PMC free article] [PubMed] [Google Scholar]
  52. Peng JR, Carol P, Richards DE, King KE, Cowling RJ, Murphy GP, Harberd NP. 1997. The Arabidopsis GAI gene defines a signalling pathway that negatively regulates gibberellin responses. Genes and Development 11, 3194–3205 [DOI] [PMC free article] [PubMed] [Google Scholar]
  53. Peng J, Richards DE, Hartley NM, et al. 1999. ‘Green revolution’ genes encode mutant gibberellin response modulators. Nature 400, 256–261 [DOI] [PubMed] [Google Scholar]
  54. Richter R, Behringer C, Muller IK, Schwechheimer C. 2010. The GATA-type transcription factors GNC and GNL/CGA1 repress gibberellin signaling downstream from DELLA proteins and phytochrome-interacting factors. Genes and Development 24, 2093–2104 [DOI] [PMC free article] [PubMed] [Google Scholar]
  55. Ross JJ, Murfet IC, Reid JB. 1997. Gibberellin mutants. Physiologia Plantarum 100, 550–560 [Google Scholar]
  56. Sabot F, Sourdille P, Bernard M. 2005. Advent of a new retrotransposon structure: the long form of the Veju elements. Genetica 125, 325–332 [DOI] [PubMed] [Google Scholar]
  57. SanMiguel PJ, Ramakrishna F, Bennetzen JL, Busso CS, Dubcovsky J. 2002. Transposable elements, genes and recombination in a 215-kb contig from wheat chromosome 5Am . Functional and Integrative Genomics 2, 70–80 [DOI] [PubMed] [Google Scholar]
  58. Schmidt AL, Gale KR, Ellis MF, Giffard PM. 2004. Sequence variation at a microsatellite locus (XGWM261) in hexaploid wheat (Triticum aestivum) varieties. Euphytica 135, 239–246 [Google Scholar]
  59. Shin J, Kim K, Kang F, Zulfugarov IS, Bae G, Lee CF, Lee D, Choi G. 2009. Phytochromes promote seedling light responses by inhibiting four negatively-acting phytochrome-interacting factors. Proceedings of the National Academy of Sciences, USA 106, 7660–7665 [DOI] [PMC free article] [PubMed] [Google Scholar]
  60. Silverstone AL, Ciampaglio CN, Sun TP. 1998. The Arabidopsis RGA gene encodes a transcriptional regulator repressing the gibberellin signal transduction pathway. The Plant Cell 10, 155–169 [DOI] [PMC free article] [PubMed] [Google Scholar]
  61. Sun TP, Gubler F. 2004. Molecular mechanism of gibberellin signaling in plants. Annual Review of Plant Biology 55, 197–223 [DOI] [PubMed] [Google Scholar]
  62. Swain SM, Singh DP. 2005. Tall tales from sly dwarves: novel functions of gibberellins in plant development. Trends in Plant Science 10, 123–129 [DOI] [PubMed] [Google Scholar]
  63. Tyler L, Thomas SG, Hu JF, Dill A, Alonso JM, Ecker JR, Sun TP. 2004. DELLA proteins and gibberellin-regulated seed germination and floral development in Arabidopsis . Plant Physiology 135, 1008–1019 [DOI] [PMC free article] [PubMed] [Google Scholar]
  64. Uddin MN, Marshall DR. 1989. Effects of dwarfing genes on yield and yield components under irrigated and rainfed conditions in wheat (Triticum aestivum L.). Euphytica 42, 127–134 [Google Scholar]
  65. van Hoof A, Frischmeyer PA, Dietz FC, Parker R. 2002. Exosome-mediated recognition and degradation of mRNAs lacking a termination codon. Science 295, 2262–2264 [DOI] [PubMed] [Google Scholar]
  66. Vasudevan S, Peltz SF, Wilusz CJ. 2002. Non-stop decay—a new mRNA surveillance pathway. BioEssays 24, 785–788 [DOI] [PubMed] [Google Scholar]
  67. Wang GZ, Miyashita NT, Tsunewaki K. 1997. Plasmon analyses of Triticum (wheat) and Aegilops: PCR-single-strand conformational polymorphism (PCR-SSCP) analyses of organellar DNAs. Proceedings of the National Academy of Sciences, USA 94, 14570–14577 [DOI] [PMC free article] [PubMed] [Google Scholar]
  68. Wu K, Wang JY, Kong ZX, Ma ZQ. 2011. Characterization of a single recessive yield trait mutant with elevated endogenous ABA concentration and deformed grains, spikelets and leaves. Plant Science 180, 306–312 [DOI] [PubMed] [Google Scholar]
  69. Wu J, Kong X, Wan J, et al. 2011. Dominant and pleiotropic effects of a GAI gene in wheat results from lack of interaction between DELLA and GID1. Plant Physiology 157, 2120–2130 [DOI] [PMC free article] [PubMed] [Google Scholar]
  70. Zhang R, Lu J, Ma G, Wu Y, Fang M, Zhao Y. 1995. Effects of Rht3 gene and 4B-chromosome with Rht3 on photosynthetic carbon-assimilation of wheat. Acta Genetica Sinica 22, 264–271 [Google Scholar]

Associated Data

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

Supplementary Materials

Supplementary Data
supp_64_11_3299__index.html (1.4KB, html)
supp_ert183_jexbot098962_file006.pdf (512.9KB, pdf)
supp_ert183_jexbot098962_file003.xls (26.5KB, xls)
supp_ert183_jexbot098962_file005.xls (59KB, xls)
supp_ert183_jexbot098962_file004.doc (35.5KB, doc)
supp_ert183_jexbot098962_file001.xls (58.5KB, xls)
supp_ert183_jexbot098962_file002.xls (44KB, xls)

Articles from Journal of Experimental Botany are provided here courtesy of Oxford University Press

RESOURCES