2023 Volume 3
Article Contents
About this article
ARTICLE   Open Access    

Floral bud differentiation and mechanism underlying androdioecy of Osmanthus fragrans

  • # These authors contributed equally: Yuanji Han, Yanxia He

More Information
  • Sweet osmanthus is an androdioecious plant; however, the mechanism underlying pistil sterility in male plants is still unclear. Scanning electron microscopy showed that the structure of pollen grains in the stamens does not differ between the sterile cultivar 'Chenghong Dangui' and the fertile cultivar 'Huangchuan Jingui'. Triphenyltetrazolium chloride and fluorescein diacetate staining as well as in vitro culture experiments revealed that pollen grains were active in both cultivars, indicating that the stamens in both 'Chenghong Dangui' and 'Huangchuan Jingui' could develop normally. When the pistils of the fertile cultivar 'Huangchuan Jingui' differentiated, two protrusions formed on the inner side of the stamen primordium, and these gradually developed and fused together to form the ovary, style, and stigma. The pistil of the sterile cultivar 'Chenghong Dangui' also formed two protrusions on the inner side of the stamen during differentiation; however, instead of fusing, two fronds were formed. These results suggest that male sweet osmanthus are formed due to the abortion of pistils during the development of floral organs. Transcriptome sequencing revealed that the expression levels of carpel development gene CRC, AG, and AGL11 were significantly lower in 'Chenghong Dangui' compared with 'Huangchuan Jingui' at different flowering stages, which provide new insight in the molecular mechanism of pistil abortion in 'Chenghong Dangui'. CRC and AG may regulate each other to promote carpel development.
  • Aquaporins (AQPs) constitute a large family of transmembrane channel proteins that function as regulators of intracellular and intercellular water flow[1,2]. Since their first discovery in the 1990s, AQPs have been found not only in three domains of life, i.e., bacteria, eukaryotes, and archaea, but also in viruses[3,4]. Each AQP monomer is composed of an internal repeat of three transmembrane helices (i.e., TM1–TM6) as well as two half helixes that are formed by loop B (LB) and LE through dipping into the membrane[5]. The dual Asn-Pro-Ala (NPA) motifs that are located at the N-terminus of two half helixes act as a size barrier of the pore via creating an electrostatic repulsion of protons, whereas the so-called aromatic/arginine (ar/R) selectivity filter (i.e., H2, H5, LE1, and LE2) determines the substrate specificity by rendering the pore constriction site diverse in both size and hydrophobicity[59]. Based on sequence similarity, AQPs in higher plants could be divided into five subfamilies, i.e., plasma membrane intrinsic protein (PIP), tonoplast intrinsic protein (TIP), NOD26-like intrinsic protein (NIP), X intrinsic protein (XIP), and small basic intrinsic protein (SIP)[1017]. Among them, PIPs, which are typically localized in the cell membrane, are most conserved and play a central role in controlling plant water status[12,1822]. Among two phylogenetic groups present in the PIP subfamily, PIP1 possesses a relatively longer N-terminus and PIP2 features an extended C-terminus with one or more conserved S residues for phosphorylation modification[5,15,17].

    Tigernut (Cyperus esculentus L.), which belongs to the Cyperaceae family within Poales, is a novel and promising herbaceous C4 oil crop with wide adaptability, large biomass, and short life period[2327]. Tigernut is a unique species accumulating up to 35% oil in the underground tubers[2830], which are developed from stolons and the process includes three main stages, i.e., initiation, swelling, and maturation[3133]. Water is essential for tuber development and tuber moisture content maintains a relatively high level of approximately 85% until maturation when a significant drop to about 45% is observed[28,32]. Thereby, uncovering the mechanism of tuber water balance is of particular interest. Despite crucial roles of PIPs in the cell water balance, to date, their characterization in tigernut is still in the infancy[21]. The recently available genome and transcriptome datasets[31,33,34] provide an opportunity to address this issue.

    In this study, a global characterization of PIP genes was conducted in tigernut, including gene localizations, gene structures, sequence characteristics, and evolutionary patterns. Moreover, the correlation of CePIP mRNA/protein abundance with water content during tuber development as well as subcellular localizations were also investigated, which facilitated further elucidating the water balance mechanism in this special species.

    PIP genes reported in Arabidopsis (Arabidopsis thaliana)[10] and rice (Oryza sativa)[11] were respectively obtained from TAIR11 (www.arabidopsis.org) and RGAP7 (http://rice.uga.edu), and detailed information is shown in Supplemental Table S1. Their protein sequences were used as queries for tBLASTn[35] (E-value, 1e–10) search of the full-length tigernut transcriptome and genome sequences that were accessed from CNGBdb (https://db.cngb.org/search/assembly/CNA0051961)[31,34]. RNA sequencing (RNA-seq) reads that are available in NCBI (www.ncbi.nlm.nih.gov/sra) were also adopted for gene structure revision as described before[13], and presence of the conserved MIP (major intrinsic protein, Pfam accession number PF00230) domain in candidates was confirmed using MOTIF Search (www.genome.jp/tools/motif). To uncover the origin and evolution of CePIP genes, a similar approach was also employed to identify homologs from representative plant species, i.e., Carex cristatella (v1, Cyperaceae)[36], Rhynchospora breviuscula (v1, Cyperaceae)[37], and Juncus effusus (v1, Juncaceae)[37], whose genome sequences were accessed from NCBI (www.ncbi.nlm.nih.gov). Gene structure of candidates were displayed using GSDS 2.0 (http://gsds.gao-lab.org), whereas physiochemical parameters of deduced proteins were calculated using ProtParam (http://web.expasy.org/protparam). Subcellular localization prediction was conducted using WoLF PSORT (www.genscript.com/wolf-psort.html).

    Nucleotide and protein multiple sequence alignments were respectively conducted using ClustalW and MUSCLE implemented in MEGA6[38] with default parameters, and phylogenetic tree construction was carried out using MEGA6 with the maximum likelihood method and bootstrap of 1,000 replicates. Systematic names of PIP genes were assigned with two italic letters denoting the source organism and a progressive number based on sequence similarity. Conserved motifs were identified using MEME Suite 5.5.3 (https://meme-suite.org/tools/meme) with optimized parameters as follows: Any number of repetitions, maximum number of 15 motifs, and a width of 6 and 250 residues for each motif. TMs and conserved residues were identified using homology modeling and sequence alignment with the structure resolved spinach (Spinacia oleracea) SoPIP2;1[5].

    Synteny analysis was conducted using TBtools-II[39] as described previously[40], where the parameters were set as E-value of 1e-10 and BLAST hits of 5. Duplication modes were identified using the DupGen_finder pipeline[41], and Ks (synonymous substitution rate) and Ka (nonsynonymous substitution rate) of duplicate pairs were calculated using codeml in the PAML package[42]. Orthologs between different species were identified using InParanoid[43] and information from synteny analysis, and orthogroups (OGs) were assigned only when they were present in at least two species examined.

    Plant materials used for gene cloning, qRT-PCR analysis, and 4D-parallel reaction monitoring (4D-PRM)-based protein quantification were derived from a tigernut variety Reyan3[31], and plants were grown in a greenhouse as described previously[25]. For expression profiling during leaf development, three representative stages, i.e., young, mature, and senescing, were selected and the chlorophyll content was checked using SPAD-502Plus (Konica Minolta, Shanghai, China) as previously described[44]. Young and senescing leaves are yellow in appearance, and their chlorophyll contents are just half of that of mature leaves that are dark green. For diurnal fluctuation regulation, mature leaves were sampled every 4 h from the onset of light at 8 a.m. For gene regulation during tuber development, fresh tubers at 1, 5, 10, 15, 20, 25, and 35 d after tuber initiation (DAI) were collected as described previously[32]. All samples with three biological replicates were quickly frozen with liquid nitrogen and stored at −80 °C for further use. For subcellular localization analysis, tobacco (Nicotiana benthamiana) plants were grown as previously described[20].

    Tissue-specific expression profiles of CePIP genes were investigated using Illumina RNA-seq samples (150 bp paired-end reads) with three biological replicates for young leaf, mature leaf, sheath of mature leaf, shoot apex, root, rhizome, and three stages of developmental tuber (40, 85, and 120 d after sowing (DAS)), which are under the NCBI accession number of PRJNA703731. Raw sequence reads in the FASTQ format were obtained using fastq-dump, and quality control was performed using fastQC (www.bioinformatics.babraham.ac.uk/projects/fastqc). Read mapping was performed using HISAT2 (v2.2.1, https://daehwankimlab.github.io/hisat2), and relative gene expression level was presented as FPKM (fragments per kilobase of exon per million fragments mapped)[45].

    For qRT-PCR analysis, total RNA extraction and synthesis of the first-strand cDNA were conducted as previously described[24]. Primers used in this study are shown in Supplemental Table S2, where CeUCE2 and CeTIP41[25,33] were employed as two reference genes. PCR reaction in triplicate for each biological sample was carried out using the SYBR-green Mix (Takara) on a Real-time Thermal Cycler Type 5100 (Thermal Fisher Scientific Oy). Relative gene abundance was estimated with the 2−ΔΔCᴛ method and statistical analysis was performed using SPSS Statistics 20 as described previously[13].

    Raw proteomic data for tigernut roots, leaves, freshly harvested, dried, rehydrated for 48 h, and sprouted tubers were downloaded from ProteomeXchange/PRIDE (www.proteomexchange.org, PXD021894, PXD031123, and PXD035931), which were further analyzed using Maxquant (v1.6.15.0, www.maxquant.org). Three dominant members, i.e., CePIP1;1, -2;1, and -2;8, were selected for 4D-PRM quantification analysis, and related unique peptides are shown in Supplemental Table S3. Protein extraction, trypsin digestion, and LC-MS/MS analysis were conducted as described previously[46].

    For subcellular localization analysis, the coding region (CDS) of CePIP1;1, -2;1, and -2;8 were cloned into pNC-Cam1304-SubN via Nimble Cloning as described before[30]. Then, recombinant plasmids were introduced into Agrobacterium tumefaciens GV3101 with the helper plasmid pSoup-P19 and infiltration of 4-week-old tobacco leaves were performed as previously described[20]. For subcellular localization analysis, the plasma membrane marker HbPIP2;3-RFP[22] was co-transformed as a positive control. Fluorescence observation was conducted using confocal laser scanning microscopy imaging (Zeiss LMS880, Germany): The wavelength of laser-1 was set as 730 nm for RFP observation, where the fluorescence was excited at 561 nm; the wavelength of laser-2 was set as 750 nm for EGFP observation, where the fluorescence was excited at 488 nm; and the wavelength of laser-3 was set as 470 nm for chlorophyll autofluorescence observation, where the fluorescence was excited at 633 nm.

    As shown in Table 1, a total of 14 PIP genes were identified from eight tigernut scaffolds (Scfs). The CDS length varies from 831 to 882 bp, putatively encoding 276–293 amino acids (AA) with a molecular weight (MW) of 29.16–31.59 kilodalton (kDa). The theoretical isoelectric point (pI) varies from 7.04 to 9.46, implying that they are all alkaline. The grand average of hydropathicity (GRAVY) is between 0.344 and 0.577, and the aliphatic index (II) ranges from 94.57 to 106.90, which are consistent with the hydrophobic characteristic of AQPs[47]. As expected, like SoPIP2;1, all CePIPs include six TMs, two typical NPA motifs, the invariable ar/R filter F-H-T-R, five conserved Froger's positions Q/M-S-A-F-W, and two highly conserved residues corresponding to H193 and L197 in SoPIP2;1 that were proven to be involved in gating[5,48], though the H→F variation was found in CePIP2;9, -2;10, and -2;11 (Supplemental Fig. S1). Moreover, two S residues, corresponding to S115 and S274 in SoPIP2;1[5], respectively, were also found in the majority of CePIPs (Supplemental Fig. S1), implying their posttranslational regulation by phosphorylation.

    Table 1.  Fourteen PIP genes identified in C. esculentus.
    Gene name Locus Position Intron no. AA MW (kDa) pI GRAVY AI TM MIP
    CePIP1;1 CESC_15147 Scf9:2757378..2759502(–) 3 288 30.76 8.82 0.384 95.28 6 47..276
    CePIP1;2 CESC_04128 Scf4:3806361..3807726(–) 3 291 31.11 8.81 0.344 95.95 6 46..274
    CePIP1;3 CESC_15950 Scf54:5022493..5023820(+) 3 289 31.06 8.80 0.363 94.57 6 49..278
    CePIP2;1 CESC_15350 Scf9:879960..884243(+) 3 288 30.34 8.60 0.529 103.02 6 33..269
    CePIP2;2 CESC_00011 Scf30:4234620..4236549(+) 3 293 31.59 9.27 0.394 101.57 6 35..268
    CePIP2;3 CESC_00010 Scf30:4239406..4241658(+) 3 291 30.88 9.44 0.432 98.97 6 31..266
    CePIP2;4 CESC_05080 Scf46:307799..309544(+) 3 285 30.44 7.04 0.453 100.32 6 28..265
    CePIP2;5 CESC_05079 Scf46:312254..314388(+) 3 286 30.49 7.04 0.512 101.68 6 31..268
    CePIP2;6 CESC_05078 Scf46:316024..317780(+) 3 288 30.65 7.68 0.475 103.06 6 31..268
    CePIP2;7 CESC_05077 Scf46:320439..322184(+) 3 284 30.12 8.55 0.500 100.00 6 29..266
    CePIP2;8 CESC_14470 Scf2:4446409..4448999(+) 3 284 30.37 8.30 0.490 106.90 6 33..263
    CePIP2;9 CESC_02223 Scf1:2543928..2545778(–) 3 283 30.09 9.46 0.533 106.47 6 31..262
    CePIP2;10 CESC_10007 Scf27:1686032..1688010(–) 3 276 29.16 9.23 0.560 106.05 6 26..256
    CePIP2;11 CESC_10009 Scf27:1694196..1696175(–) 3 284 29.71 9.10 0.577 105.49 6 33..263
    AA: amino acid; AI: aliphatic index; GRAVY: grand average of hydropathicity; kDa: kilodalton; MIP: major intrinsic protein; MW: molecular weight; pI: isoelectric point; PIP: plasma membrane intrinsic protein; Scf: scaffold; TM: transmembrane helix.
     | Show Table
    DownLoad: CSV

    To uncover the evolutionary relationships, an unrooted phylogenetic tree was constructed using the full-length protein sequences of CePIPs together with 11 OsPIPs and 13 AtPIPs. As shown in Fig. 1a, these proteins were clustered into two main groups, corresponding to PIP1 and PIP2 as previously defined[10,49], and each appears to have evolved into several subgroups. Compared with PIP1s, PIP2s possess a relatively shorter N-terminal but an extended C-terminal with one conserved S residue (Supplemental Fig. S1). Interestingly, a high number of gene repeats were detected, most of which seem to be species-specific, i.e., AtPIP1;1/-1;2/-1;3/-1;4/-1;5, AtPIP2;1/-2;2/-2;3/-2;4/-2;5/-2;6, AtPIP2;7/-2;8, OsPIP1;1/-1;2/-1;3, OsPIP2;1/-2;4/-2;5, OsPIP2;2/-2;3, CePIP1;1/-1;2, CePIP2;2/-2;3, CePIP2;4/-2;5/-2;6/-2;7, and CePIP2;9/-2;10/-2;11, reflecting the occurrence of more than one lineage-specific whole-genome duplications (WGDs) after their divergence[50,51]. In Arabidopsis that experienced three WGDs (i.e. γ, β, and α) after the split with the monocot clade[52], AtPIP1;5 in the PIP1 group first gave rise to AtPIP1;1 via the γ WGD shared by all core eudicots[50], which latter resulted in AtPIP1;3, -1;4, and -1;2 via β and α WGDs; AtPIP2;1 in the PIP2 group first gave rise to AtPIP2;6 via the γ WGD, and they latter generated AtPIP2;2, and -2;5 via the α WGD (Supplemental Table S1). In rice, which also experienced three WGDs (i.e. τ, σ, and ρ) after the split with the eudicot clade[51], OsPIP1;2 and -2;3 generated OsPIP1;1 and -2;2 via the Poaceae-specific ρ WGD, respectively. Additionally, tandem, proximal, transposed and dispersed duplications also played a role on the gene expansion in these two species (Supplemental Table S1).

    Figure 1.  Structural and phylogenetic analysis of PIPs in C. esculentus, O. sativa, and A. thaliana. (a) Shown is an unrooted phylogenetic tree resulting from full-length PIPs with MEGA6 (maximum likelihood method and bootstrap of 1,000 replicates), where the distance scale denotes the number of amino acid substitutions per site. (b) Shown are the exon-intron structures. (c) Shown is the distribution of conserved motifs among PIPs, where different motifs are represented by different color blocks as indicated and the same color block in different proteins indicates a certain motif. (At: A. thaliana; Ce: C. esculentus; PIP: plasma membrane intrinsic protein; Os: O. sativa).

    Analysis of gene structures revealed that all CePIP and AtPIP genes possess three introns and four exons in the CDS, in contrast to the frequent loss of certain introns in rice, including OsPIP1;2, -1;3, -2;1, -2;3, -2;4, -2;5, -2;6, -2;7, and -2;8 (Fig. 1b). The positions of three introns are highly conserved, which are located in sequences encoding LB (three residues before the first NPA), LD (one residue before the conserved L involved in gating), and LE (18 residues after the second NPA), respectively (Supplemental Fig. S1). The intron length of CePIP genes is highly variable, i.e., 109–993 bp, 115–1745 bp, and 95–866 bp for three introns, respectively. By contrast, the exon length is relatively less variable: Exons 2 and 3 are invariable with 296 bp and 141 bp, respectively, whereas Exons 1 and 4 are of 277–343 bp and 93–132 bp, determining the length of N- and C-terminus of PIP1 and PIP2, respectively (Fig. 1b). Correspondingly, their protein structures were shown to be highly conserved, and six (i.e., Motifs 1–6) out of 15 motifs identified are broadly present. Among them, Motif 3, -2, -6, -1, and -4 constitute the conserved MIP domain. In contrast to a single Motif 5 present in most PIP2s, all PIP1s possess two sequential copies of Motif 5, where the first one is located at the extended N-terminal. In CePIP2;3 and OsPIP2;7, Motif 5 is replaced by Motif 13; in CePIP2;2, it is replaced by two copies of Motif 15; and no significant motif was detected in this region of CePIP2;10. PIP1s and PIP2s usually feature Motif 9 and -7 at the C-terminal, respectively, though it is replaced by Motif 12 in CePIP2;6 and OsPIP2;8. PIP2s usually feature Motif 8 at the N-terminal, though it is replaced by Motif 14 in CePIP2;2 and -2;3 or replaced by Motif 11 in CePIP2;10 and -2;11 (Fig. 1c).

    As shown in Fig. 2a, gene localization of CePIPs revealed three gene clusters, i.e., CePIP2;2/-2;3 on Scf30, CePIP2;4/-2;5/-2;6/-2;7 on Scf46, and CePIP2;10/-2;11 on Scf27, which were defined as tandem repeats for their high sequence similarities and neighboring locations. The nucleotide identities of these duplicate pairs vary from 70.5% to 91.2%, and the Ks values range from 0.0971 to 1.2778 (Table 2), implying different time of their birth. According to intra-species synteny analysis, two duplicate pairs, i.e., CePIP1;1/-1;2 and CePIP2;2/-2;4, were shown to be located within syntenic blocks (Fig. 2b) and thus were defined as WGD repeats. Among them, CePIP1;1/-1;2 possess a comparable Ks value to CePIP2;2/-2;3, CePIP1;1/-1;3, and CePIP2;4/-2;8 (1.2522 vs 1.2287–1.2778), whereas CePIP2;2/-2;4 harbor a relatively higher Ks value of 1.5474 (Table 2), implying early origin or fast evolution of the latter. While CePIP1;1/-1;3 and CePIP2;1/-2;8 were characterized as transposed repeats, CePIP2;1/-2;2, CePIP2;9/-2;10, and CePIP2;8/-2;10 were characterized as dispersed repeats (Fig. 2a). The Ks values of three dispersed repeats vary from 0.8591 to 3.0117 (Table 2), implying distinct times of origin.

    Figure 2.  Duplication events of CePIP genes and synteny analysis within and between C. esculentus, O. sativa, and A. thaliana. (a) Duplication events detected in tigernut. Serial numbers are indicated at the top of each scaffold, and the scale is in Mb. Duplicate pairs identified in this study are connected using lines in different colors, i.e., tandem (shown in green), transposed (shown in purple), dispersed (shown in gold), and WGD (shown in red). (b) Synteny analysis within and between C. esculentus, O. sativa, and A. thaliana. (c) Synteny analysis within and between C. esculentus, C. cristatella, R. breviuscula, and J. effusus. Shown are PIP-encoding chromosomes/scaffolds and only syntenic blocks that contain PIP genes are marked, i.e., red and purple for intra- and inter-species, respectively. (At: A. thaliana; Cc: C. cristatella; Ce: C. esculentus; Je: J. effusus; Mb: megabase; PIP: plasma membrane intrinsic protein; Os: O. sativa; Rb: R. breviuscula; Scf: scaffold; WGD: whole-genome duplication).
    Table 2.  Sequence identity and evolutionary rate of homologous PIP gene pairs identified in C. esculentus. Ks and Ka were calculated using PAML.
    Duplicate 1 Duplicate 2 Identity (%) Ka Ks Ka/Ks
    CePIP1;1 CePIP1;3 78.70 0.0750 1.2287 0.0610
    CePIP1;2 CePIP1;1 77.20 0.0894 1.2522 0.0714
    CePIP2;1 CePIP2;4 74.90 0.0965 1.7009 0.0567
    CePIP2;3 CePIP2;2 70.50 0.1819 1.2778 0.1424
    CePIP2;4 CePIP2;2 66.50 0.2094 1.5474 0.1353
    CePIP2;5 CePIP2;4 87.30 0.0225 0.4948 0.0455
    CePIP2;6 CePIP2;5 84.90 0.0545 0.5820 0.0937
    CePIP2;7 CePIP2;6 78.70 0.0894 1.0269 0.0871
    CePIP2;8 CePIP2;4 72.90 0.1401 1.2641 0.1109
    CePIP2;9 CePIP2;10 76.40 0.1290 0.8591 0.1502
    CePIP2;10 CePIP2;8 64.90 0.2432 3.0117 0.0807
    CePIP2;11 CePIP2;10 91.20 0.0562 0.0971 0.5783
    Ce: C. esculentus; Ka: nonsynonymous substitution rate; Ks: synonymous substitution rate; PIP: plasma membrane intrinsic protein.
     | Show Table
    DownLoad: CSV

    According to inter-species syntenic analysis, six out of 14 CePIP genes were shown to have syntelogs in rice, including 1:1, 1:2, and 2:2 (i.e. CePIP1;1 vs OsPIP1;3, CePIP1;3 vs OsPIP1;2/-1;1, CePIP2;1 vs OsPIP2;4, CePIP2;2/-2;4 vs OsPIP2;3/-2;2, and CePIP2;8 vs OsPIP2;6), in striking contrast to a single one found in Arabidopsis (i.e. CePIP1;2 vs AtPIP1;2). Correspondingly, only OsPIP1;2 in rice was shown to have syntelogs in Arabidopsis, i.e., AtPIP1;3 and -1;4 (Fig. 2b). These results are consistent with their taxonomic relationships that tigernut and rice are closely related[50,51], and also imply lineage-specific evolution after their divergence.

    As described above, phylogenetic and syntenic analyses showed that the last common ancestor of tigernut and rice is more likely to possess only two PIP1s and three PIP2s. However, it is not clear whether the gene expansion observed in tigernut is species-specific or Cyperaceae-specific. To address this issue, recently available genomes were used to identify PIP subfamily genes from C. cristatella, R. breviuscula, and J. effuses, resulting in 15, 13, and nine members, respectively. Interestingly, in contrast to a high number of tandem repeats found in Cyperaceae species, only one pair of tandem repeats (i.e., JePIP2;3 and -2;4) were identified in J. effusus, a close outgroup species to Cyperaceae in the Juncaceae family[36,37]. According to homologous analysis, a total of 12 orthogroups were identified, where JePIP genes belong to PIP1A (JePIP1;1), PIP1B (JePIP1;2), PIP1C (JePIP1;3), PIP2A (JePIP2;1), PIP2B (JePIP2;2), PIP2F (JePIP2;3 and -2;4), PIP2G (JePIP2;5), and PIP2H (JePIP2;6) (Table 3). Further intra-species syntenic analysis revealed that JePIP1;1/-1;2 and JePIP2;2/-2;3 are located within syntenic blocks, which is consistent with CePIP1;1/-1;2, CePIP2;2/-2;4, CcPIP1;1/-1;2, CcPIP2;3/-2;4, RbPIP1;1/-1;2, and RbPIP2;2/-2;5 (Fig. 2c), implying that PIP1A/PIP1B and PIP2B/PIP2D were derived from WGDs occurred sometime before Cyperaceae-Juncaceae divergence. After the split with Juncaceae, tandem duplications frequently occurred in Cyperaceae, where PIP2B/PIP2C and PIP2D/PIP2E/PIP2F retain in most Cyperaceae plants examined in this study. By contrast, species-specific expansion was also observed, i.e., CePIP2;4/-2;5, CePIP2;10/-2;11, CcPIP1;2/-1;3, CcPIP2;4/-2;5, CcPIP2;8/-2;9, CcPIP2;10/-2;11, RbPIP2;3/-2;4, and RbPIP2;9/-2;10 (Table 3 & Fig. 2c).

    Table 3.  Twelve proposed orthogroups based on comparison of representative plant species.
    Orthogroup C. esculentus C. cristatella R. breviuscula J. effusus O. sativa A. thaliana
    PIP1A CePIP1;1 CcPIP1;1 RbPIP1;1 JePIP1;1 OsPIP1;3 AtPIP1;1, AtPIP1;2,
    AtPIP1;3, AtPIP1;4,
    AtPIP1;5
    PIP1B CePIP1;2 CcPIP1;2, CcPIP1;3 RbPIP1;2 JePIP1;2
    PIP1C CePIP1;3 CcPIP1;4 RbPIP1;3 JePIP1;3 OsPIP1;1, OsPIP1;2
    PIP2A CePIP2;1 CcPIP2;1 RbPIP2;1 JePIP2;1 OsPIP2;1, OsPIP2;4,
    OsPIP2;5
    AtPIP2;1, AtPIP2;2,
    AtPIP2;3, AtPIP2;4,
    AtPIP2;5, AtPIP2;6
    PIP2B CePIP2;2 CcPIP2;2 RbPIP2;2 JePIP2;2 OsPIP2;2, OsPIP2;3
    PIP2C CePIP2;3 CcPIP2;3 RbPIP2;3, RbPIP2;4
    PIP2D CePIP2;4, CePIP2;5 CcPIP2;4, CcPIP2;5 RbPIP2;5
    PIP2E CePIP2;5 CcPIP2;5 RbPIP2;6
    PIP2F CePIP2;6 CcPIP2;6
    PIP2G CePIP2;7 CcPIP2;7 RbPIP2;7 JePIP2;3, JePIP2;4
    PIP2H CePIP2;8 CcPIP2;8, CcPIP2;9 RbPIP2;8 JePIP2;5 OsPIP2;6 AtPIP2;7, AtPIP2;8
    PIP2I CePIP2;9, CePIP2;10,
    CePIP2;11
    CcPIP2;10, CcPIP2;11 RbPIP2;9, RbPIP2;10 JePIP2;6 OsPIP2;7, OsPIP2;8
    At: A. thaliana; Cc: C. cristatella; Ce: C. esculentus; Je: J. effuses; Os: O. sativa; Rb: R. breviuscula; PIP: plasma membrane intrinsic protein.
     | Show Table
    DownLoad: CSV

    Tissue-specific expression profiles of CePIP genes were investigated using transcriptome data available for young leaf, mature leaf, sheath, root, rhizome, shoot apex, and tuber. As shown in Fig. 3a, CePIP genes were mostly expressed in roots, followed by sheaths, moderately in tubers, young leaves, rhizomes, and mature leaves, and lowly in shoot apexes. In most tissues, CePIP1;1, -2;1, and -2;8 represent three dominant members that contributed more than 90% of total transcripts. By contrast, in rhizome, these three members occupied about 80% of total transcripts, which together with CePIP1;3 and -2;4 contributed up to 96%; in root, CePIP1;1, -1;3, -2;4, and -2;7 occupied about 84% of total transcripts, which together with CePIP2;1 and -2;8 contributed up to 94%. According to their expression patterns, CePIP genes could be divided into five main clusters: Cluster I includes CePIP1;1, -2;1, and -2;8 that were constitutively and highly expressed in all tissues examined; Cluster II includes CePIP2;2, -2;9, and -2;10 that were lowly expressed in all tested tissues; Cluster III includes CePIP1;2 and -2;11 that were preferentially expressed in young leaf and sheath; Cluster IV includes CePIP1;3 and -2;4 that were predominantly expressed in root and rhizome; and Cluster V includes remains that were typically expressed in root (Fig. 3a). Collectively, these results imply expression divergence of most duplicate pairs and three members (i.e. CePIP1;1, -2;1, and -2;8) have evolved to be constitutively co-expressed in most tissues.

    Figure 3.  Expression profiles of CePIP genes in various tissues, different stages of leaf development, and mature leaves of diurnal fluctuation. (a) Tissue-specific expression profiles of 14 CePIP genes. The heatmap was generated using the R package implemented with a row-based standardization. Color scale represents FPKM normalized log2 transformed counts, where blue indicates low expression and red indicates high expression. (b) Expression profiles of CePIP1;1, -2;1, and -2;8 at different stages of leaf development. (c) Expression profiles of CePIP1;1, -2;1, and -2;8 in mature leaves of diurnal fluctuation. Bars indicate SD (N = 3) and uppercase letters indicate difference significance tested following Duncan's one-way multiple-range post hoc ANOVA (p< 0.01). (Ce: C. esculentus; FPKM: Fragments per kilobase of exon per million fragments mapped; PIP: plasma membrane intrinsic protein)

    As shown in Fig. 3a, compared with young leaves, transcriptome profiling showed that CePIP1;2, -2;3, -2;7, -2;8, and -2;11 were significantly down-regulated in mature leaves, whereas CePIP1;3 and -2;1 were up-regulated. To confirm the results, three dominant members, i.e., CePIP1;1, -2;1, and -2;8, were selected for qRT-PCR analysis, which includes three representative stages, i.e., young, mature, and senescing leaves. As shown in Fig. 3b, in contrast to CePIP2;1 that exhibited a bell-like expression pattern peaking in mature leaves, transcripts of both CePIP1;1 and -2;8 gradually decreased during leaf development. These results were largely consistent with transcriptome profiling, and the only difference is that CePIP1;1 was significantly down-regulated in mature leaves relative to young leaves. However, this may be due to different experiment conditions used, i.e., greenhouse vs natural conditions.

    Diurnal fluctuation expression patterns of CePIP1;1, -2;1, and -2;8 were also investigated in mature leaves and results are shown in Fig. 3c. Generally, transcripts of all three genes in the day (8, 12, 16, and 20 h) were higher than that in the night (24 and 4 h). During the day, both CePIP1;1 and -2;8 exhibited an unimodal expression pattern that peaked at 12 h, whereas CePIP2;1 possessed two peaks (8 and 16 h) and their difference was not significant. Nevertheless, transcripts of all three genes at 20 h (onset of night) were significantly lower than those at 8 h (onset of day) as well as 12 h. In the night, except for CePIP2;1, no significant difference was observed between the two stages for both CePIP1;1 and -2;8. Moreover, their transcripts were comparable to those at 20 h (Fig. 3c).

    To reveal the expression patterns of CePIP genes during tuber development, three representative stages, i.e., 40 DAS (early swelling stage), 85 DAS (late swelling stage), and 120 DAS (mature stage), were first profiled using transcriptome data. As shown in Fig. 4a, except for rare expression of CePIP1;2, -2;2, -2;9, and -2;10, most genes exhibited a bell-like expression pattern peaking at 85 DAS, in contrast to a gradual decrease of CePIP2;3 and -2;8. Notably, except for CePIP2;4, other genes were expressed considerably lower at 120 DAS than that at 40 DAS. For qRT-PCR confirmation of CePIP1;1, -2;1, and -2;8, seven stages were examined, i.e., 1, 5, 10, 15, 20, 25, and 35 DAI, which represent initiation, five stages of swelling, and maturation as described before[32]. As shown in Fig. 4b, two peaks were observed for all three genes, though their patterns were different. As for CePIP1;1, compared with the initiation stage (1 DAI), significant up-regulation was observed at the early swelling stage (5 DAI), followed by a gradual decrease except for the appearance of the second peak at 20 DAI, which is something different from transcriptome profiling. As for CePIP2;1, a sudden drop of transcripts first appeared at 5 DAI, then gradually increased until 20 DAI, which was followed by a gradual decrease at two late stages. The pattern of CePIP2;8 is similar to -1;1, two peaks appeared at 5 and 20 DAI and the second peak was significantly lower than the first. The difference is that the second peak of CePIP2;8 was significantly lower than the initiation stage. By contrast, the second peak (20 DAI) of CePIP2;1 was significantly higher than that of the first one (1 DAI). Nevertheless, the expression patterns of both CePIP2;1 and -2;8 are highly consistent with transcriptome profiling.

    Figure 4.  Transcript and protein abundances of CePIP genes during tuber development. (a) Transcriptome-based expression profiling of 14 CePIP genes during tuber development. The heatmap was generated using the R package implemented with a row-based standardization. Color scale represents FPKM normalized log2 transformed counts, where blue indicates low expression and red indicates high expression. (b) qRT-PCR-based expression profiling of CePIP1;1, -2;1, and -2;8 in seven representative stages of tuber development. (c) Relative protein abundance of CePIP1;1, -2;1, and -2;8 in three representative stages of tuber development. Bars indicate SD (N = 3) and uppercase letters indicate difference significance tested following Duncan's one-way multiple-range post hoc ANOVA (p < 0.01). (Ce: C. esculentus; DAI: days after tuber initiation; DAS: days after sowing; FPKM: Fragments per kilobase of exon per million fragments mapped; PIP: plasma membrane intrinsic protein).

    Since protein abundance is not always in agreement with the transcript level, protein profiles of three dominant members (i.e. CePIP1;1, -2;1, and -2;8) during tuber development were further investigated. For this purpose, we first took advantage of available proteomic data to identify CePIP proteins, i.e., leaves, roots, and four stages of tubers (freshly harvested, dried, rehydrated for 48 h, and sprouted). As shown in Supplemental Fig. S2, all three proteins were identified in both leaves and roots, whereas CePIP1;1 and -2;8 were also identified in at least one of four tested stages of tubers. Notably, all three proteins were considerably more abundant in roots, implying their key roles in root water balance.

    To further uncover their profiles during tuber development, 4D-PRM-based protein quantification was conducted in three representative stages of tuber development, i.e., 1, 25, and 35 DAI. As expected, all three proteins were identified and quantified. In contrast to gradual decrease of CePIP2;8, both CePIP1;1 and -2;1 exhibited a bell-like pattern that peaked at 25 DAI, though no significant difference was observed between 1 and 25 DAI (Fig. 4c). The trends are largely in accordance with their transcription patterns, though the reverse trend was observed for CePIP2;1 at two early stages (Fig. 4b & Fig. 4c).

    As predicted by WoLF PSORT, CePIP1;1, -2;1, and -2;8 may function in the cell membrane. To confirm the result, subcellular localization vectors named pNC-Cam1304-CePIP1;1, pNC-Cam1304-CePIP2;1, and pNC-Cam1304-CePIP2;8 were further constructed. When transiently overexpressed in tobacco leaves, green fluorescence signals of all three constructs were confined to cell membranes, highly coinciding with red fluorescence signals of the plasma membrane marker HbPIP2;3-RFP (Fig. 5).

    Figure 5.  (a) Schematic diagram of overexpressing constructs, (b) subcellular localization analysis of CePIP1;1, -2;1, and -2;8 in N. benthamiana leaves. (35S: cauliflower mosaic virus 35S RNA promoter; Ce: C. esculentus; EGFP: enhanced green fluorescent protein; kb: kilobase; NOS: terminator of the nopaline synthase gene; RFP: red fluorescent protein; PIP: plasma membrane intrinsic protein).

    Water balance is particularly important for cell metabolism and enlargement, plant growth and development, and stress responses[2,19]. As the name suggests, AQPs raised considerable interest for their high permeability to water, and plasma membrane-localized PIPs were proven to play key roles in transmembrane water transport between cells[1,18]. The first PIP was discovered in human erythrocytes, which was named CHIP28 or AQP1, and its homolog in plants was first characterized in Arabidopsis, which is known as RD28, PIP2c, or AtPIP2;3[3,7,53]. Thus far, genome-wide identification of PIP genes have been reported in a high number of plant species, including two model plants Arabidopsis and rice[10,11,1317,5456]. By contrast, little information is available on Cyperaceae, the third largest family within the monocot clade that possesses more than 5,600 species[57].

    Given the crucial roles of water balance for tuber development and crop production, in this study, tigernut, a representative Cyperaceae plant producing high amounts of oil in underground tubers[28,30,32], was employed to study PIP genes. A number of 14 PIP genes representing two phylogenetic groups (i.e., PIP1 and PIP2) or 12 orthogroups (i.e., PIP1A, PIP1B, PIP1C, PIP2A, PIP2B, PIP2C, PIP2D, PIP2E, PIP2F, PIP2G, PIP2H, and PIP2I) were identified from the tigernut genome. Though the family amounts are comparative or less than 13–21 present in Arabidopsis, cassava (Manihot esculenta), rubber tree (Hevea brasiliensis), poplar (Populus trichocarpa), C. cristatella, R. breviuscula, banana (Musa acuminata), maize (Zea mays), sorghum (Sorghum bicolor), barley (Hordeum vulgare), and switchgrass (Panicum virgatum), they are relatively more than four to 12 found in eelgrass (Zostera marina), Brachypodium distachyon, foxtail millet (Setaria italic), J. effuses, Aquilegia coerulea, papaya (Carica papaya), castor been (Ricinus communis), and physic nut (Jatropha curcas) (Supplemental Table S4). Among them, A. coerulea represents a basal eudicot that didn't experience the γ WGD shared by all core eudicots[50], whereas eelgrass is an early diverged aquatic monocot that didn't experience the τ WGD shared by all core monocots[56]. Interestingly, though both species possess two PIP1s and two PIP2s, they were shown to exhibit complex orthologous relationships of 1:1, 2:2, 1:0, and 0:1 (Supplemental Table S5). Whereas AcPIP1;1/AcPIP1;2/ZmPIP1;1/ZmPIP1;2 and ZmPIP2;1/AcPIP2;1 belong to PIP1A and PIP2A identified in this study, AcPIP2;2 and ZmPIP2;2 belong to PIP2H and PIP2I, respectively (Supplemental Table S5), implying that the last common ancestor of monocots and eudicots possesses only one PIP1 and two PIP2s followed by clade-specific expansion. A good example is the generation of AtPIP1;1 and -2;6 from AtPIP1;5 and -2;1 via the γ WGD, respectively[17].

    In tigernut, extensive expansion of the PIP subfamily was contributed by WGD (2), transposed (2), tandem (5), and dispersed duplications (3). It's worth noting that, two transposed repeats (i.e., CePIP1;1/-1;3 and CePIP2;1/-2;8) are shared by rice, implying their early origin that may be generated sometime after the split with the eudicot clade but before Cyperaceae-Poaceae divergence. By contrast, two WGD repeats (i.e., CePIP1;1/-1;2 and CePIP2;2/-2;4) are shared by C. cristatella, R. breviuscula, and J. effusus but not rice and Arabidopsis, implying that they may be derived from WGDs that occurred sometime after Cyperaceae-Poaceae split but before Cyperaceae-Juncaceae divergence. The possible WGD is the one that was described in C. littledalei[58], though the exact time still needs to be studied. Interestingly, compared with Arabidopsis (1) and rice (2), tandem/proximal duplications played a more important role in the expansion of PIP genes in tigernut (5) as well as other Cyperaceae species tested (5–6), which were shown to be Cyperaceae-specific or even species-specific. These tandem repeats may play a role in the adaptive evolution of Cyperaceae species as described in a high number of plant species[14,41]. According to comparative genomics analyses, tandem duplicates experienced stronger selective pressure than genes formed by other modes (WGD, transposed duplication, and dispersed duplication) and evolved toward biased functional roles involved in plant self-defense[41].

    As observed in most species such as Arabidopsis[10,1417], PIP genes in all Cyperaceae and Juncaceae species examined in this study, i.e., tigernut, C. cristatella, R. breviuscula, and J. effuses, feature three introns with conserved positions. By contrast, zero to three introns was not only found in rice but also in other Poaceae species such as maize, sorghum, foxtail millet, switchgrass, B. distachyon, and barley[54,55], implying lineage/species-specific evolution.

    Despite the extensive expansion of PIP genes (PIP2) in tigernut even after the split with R. breviuscula, CePIP1;1, -2;1, and -2;8 were shown to represent three dominant members in most tissues examined in this study, i.e., young leaf, mature leaf, sheath, rhizome, shoot apex, and tuber, though the situation in root is more complex. CePIP1;1 was characterized as a transposed repeat of CePIP1;3, which represents the most expressed member in root. Moreover, its recent WGD repeat CePIP1;2 was shown to be lowly expressed in most tested tissues, implying their divergence. The ortholog of CePIP1;1 in rice is OsPIP1;3 (RWC-3), which was shown to be preferentially expressed in roots, stems, and leaves, in contrast to constitutive expression of OsPIP1;1 (OsPIP1a) and -1;2[5961], two recent WGD repeats. Injecting the cRNA of OsPIP1;3 into Xenopus oocytes could increase the osmotic water permeability by 2–3 times[60], though the activity is considerably lower than PIP2s such as OsPIP2;2 and -2;2[6163]. Moreover, OsPIP1;3 was shown to play a role in drought avoidance in upland rice and its overexpression in lowland rice could increase root osmotic hydraulic conductivity, leaf water potential, and relative cumulative transpiration at the end of 10 h PEG treatment[64]. CePIP2;8 was characterized as a transposed repeat of CePIP2;1. Since their orthologs are present in both rice and Arabidopsis (Supplemental Table S3), the duplication event is more likely to occur sometime before monocot-eudicot split. Interestingly, their orthologs in rice, i.e., OsPIP2;1 (OsPIP2a) and -2;6, respectively, are also constitutively expressed[61], implying a conserved evolution with similar functions. When heterologously expressed in yeast, OsPIP2;1 was shown to exhibit high water transport activity[62,6466]. Moreover, root hydraulic conductivity was decreased by approximately four folds in OsPIP2;1 RNAi knock-down rice plants[64]. The water transport activity of OsPIP2;6 has not been tested, however, it was proven to be an H2O2 transporter that is involved in resistance to rice blast[61]. More work especially transgenic tests may improve our knowledge of the function of these key CePIP genes.

    Leaf is a photosynthetic organ that regulates water loss through transpiration. In tigernut, PIP transcripts in leaves were mainly contributed by CePIP1;1, -2;1, and -2;8, implying their key roles. During leaf development, in contrast to gradual decrease of CePIP1;1 and -2;8 transcripts in three stages (i.e. young, mature, and senescing) examined in this study, CePIP2;1 peaked in mature leaves. Their high abundance in young leaves is by cell elongation and enlargement at this stage, whereas upregulation of CePIP2;1 in mature leaves may inform its possible role in photosynthesis[67]. Thus far, a high number of CO2 permeable PIPs have been identified, e.g., AtPIP2;1, HvPIP2;1, HvPIP2;2, HvPIP2;3, HvPIP2;5, and SiPIP2;7[6870]. Moreover, in mature leaves, CePIP1;1, -2;1, and -2;8 were shown to exhibit an apparent diurnal fluctuation expression pattern that was expressed more in the day and usually peaked at noon, which reflects transpiration and the fact that PIP genes are usually induced by light[11,7173]. In rice, OsPIP2;4 and -2;5 also showed a clear diurnal fluctuation in roots that peaked at 3 h after the onset of light and dropped to a minimum 3 h after the onset of darkness[11]. Notably, further studies showed that temporal and dramatic induction of OsPIP2;5 around 2 h after light initiation was triggered by transpirational demand but not circadian rhythm[74].

    As an oil-bearing tuber crop, the main economic goal of tigernut cultivation is to harvest underground tubers, whose development is highly dependent on water available[32,75]. According to previous studies, the moisture content of immature tigernut tubers maintains more than 80.0%, followed by a seed-like dehydration process with a drop of water content to less than 50% during maturation[28,32]. Thereby, the water balance in developmental tubers must be tightly regulated. Like leaves, the majority of PIP transcripts in tubers were shown to be contributed by CePIP1;1, -2;1, and -2;8, which was further confirmed at the protein level. In accordance with the trend of water content during tuber development, mRNA, and protein abundances of CePIP1;1, -2;1, and -2;8 in initiation and swelling tubers were considerably higher than that at the mature stage. High abundances of CePIP1;1, -2;1, and -2;8 at the initiation stage reflects rapid cell division and elongation, whereas upregulation of CePIP1;1 and -2;1 at the swelling stage is in accordance with cell enlargement and active physiological metabolism such as rapid oil accumulation[28,30]. At the mature stage, downregulation of PIP transcripts and protein abundances resulted in a significant drop in the moisture content, which is accompanied by the significant accumulation of late embryogenesis-abundant proteins[23,32]. The situation is highly distinct from other tuber plants such as potato (Solanum tuberosum), which may contribute to the difference in desiccation resistance between two species[32,76]. It's worth noting that, in one study, CePIP2;1 was not detected in any of the four tested stages, i.e., freshly harvested, dried, rehydrated for 48 h, and sprouted tubers[23]. By contrast, it was quantified in all three stages of tuber development examined in this study, i.e., 1, 25, and 35 DAI (corresponding to freshly harvested tubers), which represent initiation, swelling, and maturation. One possible reason is that the protein abundance of CePIP2;1 in mature tubers is not high enough to be quantified by nanoLC-MS/MS, which is relatively less sensitive than 4D-PRM used in this study[30,46]. In fact, nanoLC-MS/MS-based proteomic analysis of 30 samples representing six tissues/stages only resulted in 2,257 distinct protein groups[23].

    Taken together, our results imply a key role of CePIP1;1, -2;1, and -2;8 in tuber water balance, however, the mechanism underlying needs to be further studied, e.g., posttranslational modifications, protein interaction patterns, and transcriptional regulators.

    To our knowledge, this is the first genome-wide characterization of PIP genes in tigernut, a representative Cyperaceae plant with oil-bearing tubers. Fourteen CePIP genes representing two phylogenetic groups or 12 orthogroups are relatively more than that present in two model plants rice and Arabidopsis, and gene expansion was mainly contributed by WGD and transposed/tandem duplications, some of which are lineage or even species-specific. Among these genes, CePIP1;1, -2;1, and -2;8 have evolved to be three dominant members that are constitutively expressed in most tissues, including leaf and tuber. Transcription of these three dominant members in leaves are subjected to development and diurnal regulation, whereas in tubers, their mRNA and protein abundances are positively correlated with the moisture content during tuber development. Moreover, their plasma membrane-localization was confirmed by subcellular localization analysis, implying that they may function in the cell membrane. These findings shall not only provide valuable information for further uncovering the mechanism of tuber water balance but also lay a solid foundation for genetic improvement by regulating these key PIP members in tigernut.

    The authors confirm contribution to the paper as follows: study conception and design, supervision: Zou Z; analysis and interpretation of results: Zou Z, Zheng Y, Xiao Y, Liu H, Huang J, Zhao Y; draft manuscript preparation: Zou Z, Zhao Y. All authors reviewed the results and approved the final version of the manuscript.

    All the relevant data is available within the published article.

    This work was supported by the Hainan Province Science and Technology Special Fund (ZDYF2024XDNY171 and ZDYF2024XDNY156), China; the National Natural Science Foundation of China (32460342, 31971688 and 31700580), China; the Project of Sanya Yazhou Bay Science and Technology City (SCKJ-JYRC-2022-66), China. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

  • The authors declare that they have no conflict of interest.

  • Supplementary Table S1 Primers used in this study.
    Supplemental File 1 Differential expression of genes in the linggeng stage of the 2 cultivars.
    Supplemental File 2 Differential expression of genes in the xiangyan stage of the 2 cultivars.
    Supplemental File 3 Differential expression of genes in the initial flowering stage of the 2 cultivars.
    Supplemental File 4 Differential expression of genes in the full flowering stage of the 2 cultivars.
    Supplemental File 5 Differential expression of genes in the late full flowering stage of the 2 cultivars.
    Supplemental File 6 Enrichment of KEGG pathway in linggeng flowering stage.
    Supplemental File 7 Enrichment of KEGG pathway in xingyan flowering stage.
    Supplemental File 8 Enrichment of KEGG pathway in initial flowering stage.
    Supplemental File 9 Enrichment of KEGG pathway in full flowering stage.
    Supplemental File 10 Enrichment of KEGG pathway in late full flowering stage.
  • [1]

    Dellaporta SL, Calderon-Urrea A. 1993. Sex determination in flowering plants. The Plant Cell 5:1241−51

    doi: 10.1105/tpc.5.10.1241

    CrossRef   Google Scholar

    [2]

    Coen ES, Meyerowitz EM. 1991. The war of the whorls: genetic interactions controlling flower development. Nature 353:31−37

    doi: 10.1038/353031a0

    CrossRef   Google Scholar

    [3]

    Weigel D, Meyerowitz EM. 1994. The ABCs of floral homeotic genes. Cell 78:203−09

    doi: 10.1016/0092-8674(94)90291-7

    CrossRef   Google Scholar

    [4]

    Causier B, Schwarz-Sommer Z, Davies B. 2010. Floral organ identity: 20 years of ABCs. Seminars in Cell & Developmental Biology 21:73−79

    doi: 10.1016/j.semcdb.2009.10.005

    CrossRef   Google Scholar

    [5]

    Cheng Z, Ge W, Li L, Hou D, Ma Y, et al. 2017. Analysis of MADS-box gene family reveals conservation in floral organ ABCDE model of moso bamboo (Phyllostachys edulis). Frontiers in Plant Science 8:656

    doi: 10.3389/fpls.2017.00656

    CrossRef   Google Scholar

    [6]

    Liu J, Fu X, Dong Y, Lu J, Ren M, et al. 2018. MIKCC-type MADS-box genes in Rosa chinensis: the remarkable expansion of ABCDE model genes and their roles in floral organogenesis. Horticuture Research 5:25

    doi: 10.1038/s41438-018-0031-4

    CrossRef   Google Scholar

    [7]

    Shore P, Sharrocks AD. 1995. The MADS-box family of transcription factors. European Journal of Biochemistry 229:1−13

    Google Scholar

    [8]

    Mena M, Ambrose BA, Meeley RB, Briggs SP, Yanofsky MF, et al. 1996. Diversification of C-function activity in maize flower development. Science 274:1537−40

    doi: 10.1126/science.274.5292.1537

    CrossRef   Google Scholar

    [9]

    Yellina AL, Orashakova S, Lange S, Erdmann R, Leebens-Mack J, et al. 2010. Floral homeotic C function genes repress specific B function genes in the carpel whorl of the basal eudicot California poppy (Eschscholzia californica). EvoDevo 1:13

    doi: 10.1186/2041-9139-1-13

    CrossRef   Google Scholar

    [10]

    Lu HW, Klocko AL, Brunner AM, Ma C, Magnuson AC, et al. 2019. RNA interference suppression of AGAMOUS and SEEDSTICK alters floral organ identity and impairs floral organ determinacy, ovule differentiation, and seed-hair development in Populus. New Phytologist 222:923−37

    doi: 10.1111/nph.15648

    CrossRef   Google Scholar

    [11]

    Liu H, Li J, Gong P, He C. 2023. The origin and evolution of carpels and fruits from an evo-devo perspective. Journal of Integrative Plant Biology 65:283−98

    doi: 10.1111/jipb.13351

    CrossRef   Google Scholar

    [12]

    Fourquin C, Vinauger-Douard M, Fogliani B, Dumas C, Scutt CP. 2005. Evidence that CRABS CLAW and TOUSLED have conserved their roles in carpel development since the ancestor of the extant angiosperms. Proceedings of the National Academy of Sciences of the United States of America 102:4649−54

    doi: 10.1073/pnas.0409577102

    CrossRef   Google Scholar

    [13]

    Morel P, Heijmans K, Ament K, Chopy M, Trehin C, et al. 2018. The floral C-lineage genes trigger nectary development in Petunia and Arabidopsis. The Plant Cell 30:2020−37

    doi: 10.1105/tpc.18.00425

    CrossRef   Google Scholar

    [14]

    Gong P, Song C, Liu H, Li P, Zhang M, et al. 2021. Physalis floridana CRABS CLAW mediates neofunctionalization of GLOBOSA genes in carpel development. Journal of Experimental Botany 72:6882−903

    doi: 10.1093/jxb/erab309

    CrossRef   Google Scholar

    [15]

    Bowman JL, Smyth DR, Meyerowitz EM. 1989. Genes directing flower development in Arabidopsis. The Plant Cell 1:37−52

    doi: 10.1105/tpc.1.1.37

    CrossRef   Google Scholar

    [16]

    Yanofsky MF, Ma H, Bowman JL, Drews GN, Feldmann KA, et al. 1990. The protein encoded by the Arabidopsis homeotic gene agamous resembles transcription factors. Nature 346:35−39

    doi: 10.1038/346035a0

    CrossRef   Google Scholar

    [17]

    Davies B, Motte P, Keck E, Saedler H, Sommer H, et al. 1999. PLENA and FARINELLI: redundancy and regulatory interactions between two Antirrhinum MADS-box factors controlling flower development. The EMBO Journal 18:4023−34

    doi: 10.1093/emboj/18.14.4023

    CrossRef   Google Scholar

    [18]

    Kapoor M, Tsuda S, Tanaka Y, Mayama T, Okuyama Y, et al. 2002. Role of petunia pMADS3 in determination of floral organ and meristem identity, as revealed by its loss of function. The Plant Journal 32:115−27

    doi: 10.1046/j.1365-313X.2002.01402.x

    CrossRef   Google Scholar

    [19]

    Hands P, Vosnakis N, Betts D, Irish VF, Drea S. 2011. Alternate transcripts of a floral developmental regulator have both distinct and redundant functions in opium poppy. Annals of Botany 107:1557−66

    doi: 10.1093/aob/mcr045

    CrossRef   Google Scholar

    [20]

    Nakatsuka T, Saito M, Yamada E, Fujita K, Yamagishi N, et al. 2015. Isolation and characterization of the C-class MADS-box gene involved in the formation of double flowers in Japanese gentian. BMC Plant Biology 15:e182

    doi: 10.1186/s12870-015-0569-3

    CrossRef   Google Scholar

    [21]

    Klocko AL, Borejsza-Wysocka E, Brunner AM, Shevchenko O, Aldwinckle H, et al. 2016. Transgenic suppression of AGAMOUS genes in apple reduces fertility and increases floral attractiveness. PLoS ONE 11:e0159421

    doi: 10.1371/journal.pone.0159421

    CrossRef   Google Scholar

    [22]

    Eshed Y, Baum SF, Bowman JL. 1999. Distinct mechanisms promote polarity establishment in carpels of Arabidopsis. Cell 99:199−209

    doi: 10.1016/S0092-8674(00)81651-7

    CrossRef   Google Scholar

    [23]

    Yamaguchi T, Nagasawa N, Kawasaki S, Matsuoka M, Nagato Y, et al. 2004. The YABBY gene DROOPING LEAF regulates carpel specification and midrib development in Oryza sativa. The Plant Cell 16:500−09

    doi: 10.1105/tpc.018044

    CrossRef   Google Scholar

    [24]

    Sugiyama SH, Yasui Y, Ohmori S, Tanaka W, Hirano HY. 2019. Rice flower development revisited: regulation of carpel specification and flower meristem determinacy. Plant and Cell Physiology 60:1284−95

    doi: 10.1093/pcp/pcz020

    CrossRef   Google Scholar

    [25]

    Bowman JL, Smyth DR. 1999. CRABS CLAW, a gene that regulates carpel and nectary development in Arabidopsis, encodes a novel protein with zinc finger and helix-loop-helix domains. Development 126:2387−96

    doi: 10.1242/dev.126.11.2387

    CrossRef   Google Scholar

    [26]

    Lee JY, Baum SF, Alvarez J, Patel A, Chitwood DH, et al. 2005. Activation of CRABS CLAW in the nectaries and carpels of Arabidopsis. The Plant Cell 17:25−36

    doi: 10.1105/tpc.104.026666

    CrossRef   Google Scholar

    [27]

    Orashakova S, Lange M, Lange S, Wege S, Becker A. 2009. The CRABS CLAW ortholog from California poppy (Eschscholzia californica, Papaveraceae), EcCRC, is involved in floral meristem termination, gynoecium differentiation and ovule initiation. The Plant Journal 58:682−93

    doi: 10.1111/j.1365-313X.2009.03807.x

    CrossRef   Google Scholar

    [28]

    Alvarez J, Smyth DR. 1999. CRABS CLAW and SPATULA, two Arabidopsis genes that control carpel development in parallel with AGAMOUS. Development 126:2377−86

    doi: 10.1242/dev.126.11.2377

    CrossRef   Google Scholar

    [29]

    Gómez-Mena C, de Folter S, Costa MMR, Angenent GC, Sablowski R. 2005. Transcriptional program controlled by the floral homeotic gene AGAMOUS during early organogenesis. Development 132:429−38

    doi: 10.1242/dev.01600

    CrossRef   Google Scholar

    [30]

    Li X, Yang Y, Zheng W, Hou J. 2002. On Flower-bud Induction in Fruit Trees. Chinese Bulletin of Botany 19:385−95

    doi: 10.3969/j.issn.1674-3466.2002.04.001

    CrossRef   Google Scholar

    [31]

    Ma Y, Dai S. 2004. Flower bud differentiation mechanism of anthophyta. Molecular Plant Breeding 1:539−45

    doi: 10.3969/j.issn.1672-416X.2003.04.014

    CrossRef   Google Scholar

    [32]

    Li J, Dong M, Shang F. 2007. Study on the Flower Bud Differentiation of Osmanthus fragrans 'Dangui' and O. fragrans 'Ziyingui'. Chinese Bulletin of Botany 24:620−23

    doi: 10.3969/j.issn.1674-3466.2007.05.009

    CrossRef   Google Scholar

    [33]

    Wodehouse RP. 1935. Pollen grains. pp. xv + 574. New York: McGraw-Hill Book Company. 323−40 pp.

    [34]

    Walker JW. 1976. Evolutionary significance of the exine in the pollen of primitive angiosperms. In The Evolutionary Significance of Exine, eds. Ferguson IK, Muller J. London: Academic Press. 251−308 pp.

    [35]

    Vernet P, Lepercq P, Billiard S, Bourceaux A, Lepart J, et al. 2016. Evidence for the long-term maintenance of a rare self-incompatibility system in Oleaceae. New Phytologist 210:1408−17

    doi: 10.1111/nph.13872

    CrossRef   Google Scholar

    [36]

    Charlesworth D. 1984. Androdioecy and the evolution of dioecy. Biological Journal of the Linnean Society 22:333−48

    doi: 10.1111/j.1095-8312.1984.tb01683.x

    CrossRef   Google Scholar

    [37]

    Wallander E. 2008. Systematics of Fraxinus (Oleaceae) and evolution of dioecy. Plant Systematics and Evolution 273:25−49

    doi: 10.1007/s00606-008-0005-3

    CrossRef   Google Scholar

    [38]

    Ross MD, Weir BS. 1976. Maintenences of males and females in hermaphrodite populations and the evolution of dioecy. Evolution 30:425−41

    doi: 10.2307/2407568

    CrossRef   Google Scholar

    [39]

    Lloyd DG. 1975. The maintenance of gynodioecy and androdioecy in angiosperms. Genetica 45:325−39

    doi: 10.1007/BF01508307

    CrossRef   Google Scholar

    [40]

    Ross MD. 1982. Five evolutionary pathways to subdioecy. The American Naturalist 119:297−318

    doi: 10.1086/283911

    CrossRef   Google Scholar

    [41]

    Xu L, Wang J, Song L, Wang L. 2009. Preliminary study on the functions of AGAMOUS homologous genes in Pisum sativum (in Chinese). Chinese Science Bulletin 54:3207−12

    doi: 10.1360/972009-724

    CrossRef   Google Scholar

    [42]

    Liljegren SJ, Ditta GS, Eshed Y, Savidge B, Bowman JL, et al. 2000. SHATTERPROOF MADS-box genes control seed dispersal in Arabidopsis. Nature 404:766−70

    doi: 10.1038/35008089

    CrossRef   Google Scholar

    [43]

    Pinyopich A, Ditta GS, Savidge B, Liljegren SJ, Baumann E, et al. 2003. Assessing the redundancy of MADS-box genes during carpel and ovule development. Nature 424:85−88

    doi: 10.1038/nature01741

    CrossRef   Google Scholar

    [44]

    Favaro R, Pinyopich A, Battaglia R, Kooiker M, Borghi L, et al. 2003. MADS-box protein complexes control carpel and ovule development in Arabidopsis. The Plant Cell 15:2603−11

    doi: 10.1105/tpc.015123

    CrossRef   Google Scholar

    [45]

    Mejía N, Soto B, Guerrero M, Casanueva X, Houel C, et al. 2011. Molecular, genetic and transcriptional evidence for a role of VvAGL11 in stenospermocarpic seedlessness in grapevine. BMC Plant Biology 11:57

    doi: 10.1186/1471-2229-11-57

    CrossRef   Google Scholar

    [46]

    Ocarez N, Mejía N. 2016. Suppression of the D-class MADS-box AGL 11 gene triggers seedlessness in fleshy fruits. Plant Cell Reports 35:239−54

    doi: 10.1007/s00299-015-1882-x

    CrossRef   Google Scholar

    [47]

    Zhang S, Tan FQ, Chung CH, Slavkovic F, Devani RS, et al. 2022. The control of carpel determinacy pathway leads to sex determinationin cucurbits. Science 378:543−49

    doi: 10.1126/science.add4250

    CrossRef   Google Scholar

    [48]

    Lenhard M, Bohnert A, Jürgens G, Laux T. 2001. Termination of stem cell maintenance in Arabidopsis floral meristems by interactions between WUSCHEL and AGAMOUS. Cell 105:805−14

    doi: 10.1016/S0092-8674(01)00390-7

    CrossRef   Google Scholar

    [49]

    Lohmann JU, Hong RL, Hobe M, Busch MA, Parcy F, et al. 2001. A molecular link between stem cell regulation and floral patterning in Arabidopsis. Cell 105:793−803

    doi: 10.1016/S0092-8674(01)00384-1

    CrossRef   Google Scholar

    [50]

    Prunet N, Yang W, Das P, Meyerowitz EM, Jack TP. 2017. SUPERMAN prevents class B gene expression and promotes stem cell termination in the fourth whorl of Arabidopsis thaliana flowers. Proceedings of the National Academy of Sciences of the United States of America 114:7166−71

    doi: 10.1073/pnas.1705977114

    CrossRef   Google Scholar

    [51]

    Xu Y, Prunet N, Gan ES, Wang Y, Stewart D, et al. 2018. SUPERMAN regulates floral whorl boundaries through control of auxin biosynthesis. The EMBO Journal 37:e97499

    doi: 10.15252/embj.201797499

    CrossRef   Google Scholar

    [52]

    Yamaguchi N, Huang JB, Xu YF, Tanoi K, Ito T. 2017. Fine-tuning of auxin homeostasis governs the transition from floral stem cell maintenance to gynoecium formation. Nature Communications 8:1125

    doi: 10.1038/s41467-017-01252-6

    CrossRef   Google Scholar

    [53]

    Castañeda L, Giménez E, Pineda B, García-Sogo B, Ortiz-Atienza A, et al. 2022. Tomato CRABS CLAW paralogues interact with chromatin remodeling factors to mediate carpel development and floral determinacy. New Phytologist 234:1059−74

    doi: 10.1111/nph.18034

    CrossRef   Google Scholar

    [54]

    Li Z. 1996. Sectioning of plant tissue. pp. 183. Beijng: Peking University Press. 130−45 pp.

    [55]

    Love MI, Huber W, Anders S. 2014. Moderated estimation of fold change and dispersion for RNA-seq data with DESeq2. Genome Biology 15:550

    doi: 10.1186/s13059-014-0550-8

    CrossRef   Google Scholar

    [56]

    Han Y, Lu M, Yue S, Li K, Dong M, et al. 2022. Comparative methylomics and chromatin accessibility analysis in Osmanthus fragrans uncovers regulation of genic transcription and mechanisms of key floral scent production. Horticulture Research 9:uhac096

    doi: 10.1093/hr/uhac096

    CrossRef   Google Scholar

  • Cite this article

    Han Y, He Y, Yue S, Guo B, Zhu Q, et al. 2023. Floral bud differentiation and mechanism underlying androdioecy of Osmanthus fragrans. Ornamental Plant Research 3:11 doi: 10.48130/OPR-2023-0011
    Han Y, He Y, Yue S, Guo B, Zhu Q, et al. 2023. Floral bud differentiation and mechanism underlying androdioecy of Osmanthus fragrans. Ornamental Plant Research 3:11 doi: 10.48130/OPR-2023-0011

Figures(5)  /  Tables(1)

Article Metrics

Article views(4879) PDF downloads(806)

ARTICLE   Open Access    

Floral bud differentiation and mechanism underlying androdioecy of Osmanthus fragrans

Ornamental Plant Research  3 Article number: 11  (2023)  |  Cite this article

Abstract: Sweet osmanthus is an androdioecious plant; however, the mechanism underlying pistil sterility in male plants is still unclear. Scanning electron microscopy showed that the structure of pollen grains in the stamens does not differ between the sterile cultivar 'Chenghong Dangui' and the fertile cultivar 'Huangchuan Jingui'. Triphenyltetrazolium chloride and fluorescein diacetate staining as well as in vitro culture experiments revealed that pollen grains were active in both cultivars, indicating that the stamens in both 'Chenghong Dangui' and 'Huangchuan Jingui' could develop normally. When the pistils of the fertile cultivar 'Huangchuan Jingui' differentiated, two protrusions formed on the inner side of the stamen primordium, and these gradually developed and fused together to form the ovary, style, and stigma. The pistil of the sterile cultivar 'Chenghong Dangui' also formed two protrusions on the inner side of the stamen during differentiation; however, instead of fusing, two fronds were formed. These results suggest that male sweet osmanthus are formed due to the abortion of pistils during the development of floral organs. Transcriptome sequencing revealed that the expression levels of carpel development gene CRC, AG, and AGL11 were significantly lower in 'Chenghong Dangui' compared with 'Huangchuan Jingui' at different flowering stages, which provide new insight in the molecular mechanism of pistil abortion in 'Chenghong Dangui'. CRC and AG may regulate each other to promote carpel development.

    • In flowering plants, cross-pollination is a common mode of reproduction and promotes genetic variation, gene exchange, and species adaptation. Plants have evolved a variety of reproductive systems involving cross-pollination at the individual and population levels, such as monoecy, dioecy, andromonoecy, gynomonoecy, and androdioecy[1].

      Based on studies of Arabidopsis thaliana and snapdragon mutants, Coen & Meyerowitz and Causiera & Meyerowitz proposed the well-known ABC model, which suggests that the formation of the four whorls of floral organs is a result of the combined action of class A, B, and C genes[2,3]. Subsequent studies identified class D and E genes, and the flower organ development model gradually developed into an ABCDE model[46], with MADS-box genes accounting for the majority of genes in the model[7]. In core dicotyledons, A and E functional genes determine the development of the calyx; A, B, and E functional genes determine the development of petals; B, C, and E functional genes determine the development of stamens; and C and E functional genes determine the development of carpels[7].

      Studies have shown that the class C MADS-box gene AGAMOUS (AG)[811] and the non-MADS-box gene CRABS CLAW (CRC)[1214] play important roles in carpel development. AG regulates the differentiation of carpels and interacts with other genes to control floral determinacy. In Arabidopsis, the loss of AG function usually results in the absence of carpels, and the carpels are replaced by indeterminate perianth whorls[15,16]. The loss-of-function of AG and class C homeotic genes shows similar phenotypes in other plants, such as Antirrhinum[17], petunia[18], opium poppy (Papaver somniferum)[19], Nicotiana benthamiana[20], and apple (Malus domestica)[21].

      CRC regulates the development of tissues derived from the abaxial side of the carpel primordium[22]. In plants with mutations in CRC and CRC orthologs, the carpel is replaced with petals, stamens, or a flower[23,24]. CRC genes influence carpel development in various plants, such as A. thaliana[25,26], Eschscholzia californica[27], Pisum sativum[12], Petunia hybrid[13], and Physalis floridan[14]. CRC is a master regulator in the gene regulatory network determining floral morphology, particularly carpel formation[11]. Both CRC and AG are core transcription factors that can regulate each other to promote carpel development[28,29].

      Sweet osmanthus (Osmanthus fragrans Lour.) is a well-known ornamental and aromatic plant in China. Preliminary studies have found that sweet osmanthus is an androdioecious breeding system; however, the molecular mechanism underlying the formation of the androdiecious reproductive system in the species has not been determined. In this study, morphology and molecular analyses were combined to investigate the mechanism underlying pistil sterility in androdiecious sweet osmanthus plants. The results of this study will provide a theoretical basis for the conservation of germplasm resources and the cultivation of new cultivars of sweet osmanthus.

    • According to the observations of paraffin sections, flower bud differentiation in sweet osmanthus can be divided into five stages: involucre differentiation, primordium differentiation, terminal flower perianth differentiation, stamen differentiation, and pistil differentiation. Involucre differentiation was completed at the end of June, when the growth cone gradually flattened and widened, producing two protrusions on its surface, the involucre primordium (Fig. 1af). As the cells grew, the involucre and primordia became larger and bent upward until the two primordia were close together (Fig. 1g). At the beginning of July, the growth points inside the involucre gradually expanded into a hemispherical shape, and the central tip and both sides of the lower part showed darker staining. The growth points gradually expanded and separated to form multiple raised semicircles. At this time, the rudimentary structure of cymose inflorescence (Fig. 1h) was already visible. The protrusions on both sides of the base of the involucre formed the lateral flower primordium (Fig. 1i), while the central protrusion formed the terminal flower primordium. During the perianth differentiation stage, the terminal flower primordium gradually flattened, and two small protrusions were formed on both sides, corresponding to the calyx primordia (Fig. 1jl). Since the calyx of sweet osmanthus is very short, the calyx primordium stopped growing after a short period, and two protrusions formed at the center. These protrusions were the petal primordia and continued to grow and bend into a curved petal shape (Fig. 1m, n). This period proceeded rapidly, and the differentiation of the perianth primordia was completed in less than a month. At the time of the formation of petal primordia, the stamen primordia began to appear, and two small protrusions (Fig. 1o) appeared on the inner side of petals at the end of July; these developed gradually, enlarged, and finally formed the stamen primordia (Fig. 1p). At this time, the filaments and anthers also began to differentiate. At the beginning of August, after the stamen primordia of the fertile 'Huangchuan Jingui' matured, two small protrusions were formed on its inner side. With continuous growth, the protrusions moved towards each other and gradually fused to form a carpel, forming a slit in the center (Fig. 1q). The carpel primordia continued to develop, expanded into an ovary, and formed a style and a stigma (Fig. 1r & s). Two small protrusions (Fig. 1t) were also formed on the inner side of the stamens of sterile 'Chenghong Dangui'; however, the protrusions gradually elongated and did not fuse (Fig. 1u). Instead, two fronds were formed, and ovary development was not seen (Fig. 1v). The pistil of 'Huangchuan Jingui' was able to develop normally to form fruits after fertilization (Fig. 2a & b), while pistil abortion was observed in 'Chenghong Dangui', with no fruit formation (Fig. 2c & d).

      Figure 1. 

      Floral bud differentiation of sweet osmanthus. (a) − (g) Involucre differentiation. (h), (i) flower primordium differentiation. (j) − (n) Terminal flower perianth differentiation. (o), (p) Stamen differentiation. (q) − (s) Pistil differentiation. (t) − (v) Pistil differentiation (sterile). in: Involucre primordium, sf: Side flower primordium, se: Sepal primordium, pe: Petal primordium, an: Stamen primordium, pi: Pistil primordium.

      Figure 2. 

      Development of the pistil of the two cultivars. (a) The full flowering stage of 'Huangchun Jingui' cultivar. (b) Fruit formation of 'Huangchun Jingui' cultivar. (c) The full flowering stage of 'Chenghong Dangui' cultivar. (d) Pistil abortion and no fruit formation in 'Chenghong Dangui' cultivar.

    • Scanning electron microscopy results showed that the pollen grains of both cultivars ('Huangchuan Jingui' and 'Chenghong Dangui') were oblong and round with three lobes in polar view (Fig. 3a & b); they were classified as N3P4C5 according to the NPC classification system. The outer wall of the pollen had a reticulate pattern, with fine and small reticulation. The germination pore was a 3-pore groove, and the poles of the pore groove were narrow and became wide in the middle (Fig. 3ch). The mean polar axis length of 'Huangchuan Jingui' was 18.0 μm, while that of 'Chenghong Dangui' was 18.6 μm. Both cultivars had small pollen with significant differences in polar axis length (Table 1). In addition, the P/E, pore groove length (L), and L/P values differed significantly between the two cultivars (Table 1). After triphenyltetrazolium chloride (TTC) staining and observing under a microscope, the pollen grains of both 'Huangchuan Jinghui' and 'Chenghong Dangui' were stained red or light red (Fig. 3i,j). After fluorescein diacetate (FDA) staining and observing under fluorescence microscope, the pollen grains of both 'Huangchuan Jingui' and 'Chenghong Dangui' exhibited green fluorescence (Fig. 3k,l), indicating that the pollen grains of both cultivars were active. The results of aniline blue staining showed that regardless of whether the pollen of 'Huangchuan Jingui' or 'Chenghong Dangui' was used to pollinate the stigma of 'Huangchuan Jingui', the germinated pollen tubes could be seen on the stigma of the pistil style of the 'Huangchuan Jingui' (Fig. 3m,n). These results indicated that pollen grains of both male and bisexual flowers were active on the stigma of the pistil.

      Figure 3. 

      Morphological characteristics and activity of pollen grains of the two cultivars. (a) − (h) Morphological characteristics of pollen grains. (i) − (n) Activity of pollen grains of the two cultivars.

      Table 1.  Measurement results of pollen.

      CultivarsPolar axis (P) (μm)Equator axis (E) (μm)P/EPore groove length (L) (μm)L/P
      'Huangchuan Jingui'18.0 ± 1.0110.8 ± 1.051.63 ± 0.1112.4 ± 1.520.70 ± 0.08
      'Chenghong Dangui'18.6 ± 0.66*10.9 ± 0.541.70 ± 0.12*13.9 ± 1.21*0.77 ± 0.05*
      * P < 0.05
    • To investigate the differential expression of genes related to floral organ development in 'Huangchuan Jingui' and 'Chenghong Dangui' and their effects on the development of carpels and pistils, a transcriptome sequencing analysis of the floral organs of 'Huangchuan Jingui' and 'Chenghong Dangui' at the linggeng stage, xingyan stage, initial flowering stage, full flowering stage, and late full flowering stage was performed. Compared with 'Chenghong Dangui', the numbers of upregulated differentially expressed genes (DEGs) were 4,937, 4,644, 3,981, 4,165, 4,910 during the linggeng stage, xingyan stage, initial flowering stage, full flowering stage and late full flowering stage, while the downregulated DGEs were 3,842, 3,341, 3,003, 3,462, 4,090 respectively in 'Huangchuan Jingui' (Supplemental Files 15). From the KEGG enrichment analysis results, we found that the most enriched DEGs exists in the plant-pathogen interaction pathway of the five flowering stages, with 209, 204, 178, 222, and 252 DEGs, respectively. The secondly DEGs enrichment pathway is plant hormone signal transduction, which contains 162, 163, 140, 190, and 226 DEGs in the five flowering stages (Supplemental Files 610). Thus, we presumed that the development of sweet osmanthus flower organs may be greatly influenced by hormones.

      MADS-box genes are key genes for the development of floral organs, and the transcriptome sequencing results showed that, compared with levels in 'Chenghong Dangui', 7, 8, 6, 8, and 10 MADS-box genes were up-regulated in 'Huangchuan Jingui' during the linggeng stage, xingyan stage, initial flowering stage, full flowering stage, and late full flowering stage, respectively (Fig. 4). Among these, class A genes included AP1 and FUL, which were up-regulated in the linggeng stage and the xingyan stages of the flowering organs of 'Huangchuan Jingui'. Class B genes included GLOBOSA and PISTILLATA, which were up-regulated in the floral organs of 'Huangchuan Jingui' at both the full flowering and the late full flowering stages. The class C gene AGAMOUS (AG) was up-regulated in the floral organs of 'Huangchuan Jingui' at both the initial and full flowering stages. In addition, several AGAMOUS-like (AGL) genes, such as AGL11, AGL12, AGL81, AGL17, and AGL81, were up-regulated in the floral organs of 'Huangchuan Jingui' in different flowering stages (Fig. 4). These genes may also be involved in the development of floral organs and carpel formation. In particular, the expression of AGL11 was significantly up-regulated in all five periods of flowering organ development of 'Huangchuan Jingui'. In addition, the CRC gene, a non-MADS-box gene involved in carpel development, was significantly up-regulated in all five periods in the floral organs of 'Huangchuan Jingui'.

      Figure 4. 

      Differential expression of genes related to flower organ development in the two cultivars. (a) Differential expression genes related to flower organ development in the linggeng stage. (b) Differential expression genes related to flower organ development in the xingyan stage. (c) Differential expression genes related to flower organ development in the initial flowering stage. (d) Differential expression genes related to flower organ development in the full flowering stage. (e) Differential expression genes related to flower organ development in the late full flowering stage. Y: 'Huangchuan Jingui', D: 'Chenghong Dangui'. Y1 and D1: Linggeng stage, Y2 and D2: Xingyan stage, Y3 and D3: Initial flowering stage, Y4 and D4: Full flowering stage, Y5 and D5: Late full flowering stage. Three replicates for each analysis.

    • A transcriptome sequencing analysis showed that the CRC gene was significantly up-regulated in the floral organs of 'Huangchuan Jingui' across all five periods, compared with levels in 'Chenghong Dangui', while the AG gene was up-regulated in the floral organs of 'Huangchuan Jingui' at the initial and full flowering stages. Previous studies have shown that CRC and AG are two key genes that determine carpel formation[10,11,13,14]. A dual luciferase assay showed that the activity of the AG gene promoter was 1.43-fold higher in tobacco leaves co-transformed with 35S::CRC and AGpro::LUC plasmids as compared to that in control tobacco leaves transformed with only AGpro::LUC plasmids (Fig. 5a). Compared with activity in control tobacco leaves transformed with only CRCpro::LUC plasmids, the activity of the CRC gene promoter was up-regulated by 1.45-fold in tobacco leaves co-transformed with 35S::AG and CRCpro::LUC plasmids (Fig. 5b).

      Figure 5. 

      Transcriptional regulation of CRC and AG genes. (a) The effector and reporter plasmids used in dual-LUC assays. REN: Renilla luciferase, LUC: Firefly luciferase. (b) The AG promoter activity (LUC/REN ratio) of tobacco leaves coinfiltration with Agrobacteria carrying effector and reporter. (c) The CRC promoter activity (LUC/REN ratio) of tobacco leaves coinfiltration with Agrobacteria carrying effector and reporter. The data represent the means ± SD of three replicates from three independent experiments. * P < 0.05.

    • Flower bud differentiation is an important process in the transition of plants from the vegetative to the reproductive stage[30]. Understanding the mechanism underlying flower bud differentiation in plants is important for formulating reasonable cultivation measures for flowering regulation, implementing them to improve annual production of ornamental plants, and clarifying the genetic regulation of plant traits[31]. In this study, the leaf-like structure surrounded by each inflorescence of sweet osmanthus was considered as the involucre; accordingly, the bract differentiation period could be more appropriately termed as the involucre differentiation period. The period from the emergence of the inflorescence primordium to the differentiation of the calyx primordium involves the emergence of the embryonic cyme of sweet osmanthus, and the terminal flowers and lateral flowers differentiate later. Therefore, the inflorescence primordium and flower primordium emerge simultaneously at this stage, which is more appropriately referred to as the flower primordium differentiation period. The emergence of the flower primordium and the formation of the perianth primordium are short and continuous processes; accordingly, the sepal and petal differentiation stages should be combined into the perianth differentiation stage. Therefore, in this study, flower bud differentiation in sweet osmanthus was divided into five periods: involucre differentiation, floral primordium differentiation, terminal perianth differentiation, stamen differentiation, and pistil differentiation.

      Previous studies have shown that when the pistil of fertile varieties differentiate, the carpel primordium first appears as a protrusion and develops gradually, forming a small hole in the center, which extends and fuses to form the carpel primordium. In sterile varieties, two protrusions form on the inner side of the stamen, and these do not fuse at the end but form two fronds[32]. In this study, regardless of whether the variety was fertile or sterile, two protrusions formed initially on the inner side of the stamen primordia. However, in the later growth period of the fertile 'Huangchuan Jingui', the two protrusions fused together and gradually developed into the ovary, style, and stigma. The 'small hole' would be the slit between the two protrusions when they fuse.

      The shape and size of plant pollen, the type of germination pore, and surface characteristics of plant pollen are important features for evolutionary analyses and plant classification. Previous studies have shown that a longer pollen is related to a smaller surface-to-volume ratio and more derived taxa, while the most derived pollen shows reticulate, striped reticulate, or fine reticulate outer wall characteristics[33,34]. The results of this study revealed that the surface of the pollen grains of the two varieties of sweet osmanthus had reticulate patterns, indicating that their pollen was highly evolved. Among the two varieties, the P/E value of 'Huangchuan Jingui' was smaller than that of 'Chenghong Dangui', indicating that the pollen grains of 'Huangchuan Jingui' were rounder while those of 'Chenghong Dangui' were longer. Hence, 'Chenghong Dangui' may be more evolved than 'Huangchuan Jingui'.

      Androdioecy is an extremely rare plant reproductive system where in both male and hermaphroditic plants are present. It has been reported in only a few plants (<0.005%) and is usually found in Oleaceae[1,35]. Androdioecy is considered an intermediate state between monoecious and dioecious plants[36,37], probably resulting from the sterility of pistils in monoecious plants[36, 3840]. In the ABCDE model of floral organ development, class C functional genes (e.g., AG) determine the development of pistils and carpels. Class C genes have been found in Zeamays[6], E. californica[9], P. sativum[41], Populus alba[10], A. thaliana[16,42,43] and other plants. In this study, the class C functional gene AG was up-regulated in both the initial and full flowering stages of 'Huangchuan Jingui', suggesting that these stages are critical periods for carpel development. The ectopic expression of the STK (AGL11) gene in ag-deficient mutants can promote carpel development[44]. Hence, AGL11 is an important gene for the development of carpels, ovules, and fruits[4346]. In this study, the expression levels of AGL11 in 'Huangchuan Jingui' across five periods, including the ling geng stage, xingyan stage, initial flowering stage, full flowering stage, and late full flowering stage, were significantly higher than those of 'Chenghong Dangui', indicating that this gene is an important determinant of the development of carpels and ovules in sweet osmanthus. Previous studies have shown that the CRC gene contributes to carpel and fruit development. Arabidopsis CRC is primarily required for the elaboration of carpel morphology[25,28], and mutations in the CRC gene contribute to the female-to-male transition in melons[47]. In this study, the expression level of the CRC gene in all five stages was significantly higher in 'Huangchuan Jingui' than that in 'Chenghong Dangui', suggesting that this gene is involved in the development of carpels in sweet osmanthus.

      Previous studies have shown that both CRC and AG are important genes for carpel development, and AG can promote the expression of CRC genes[13,4853]. The results of this study showed that AG in sweet osmanthus can bind to the promoter and thereby regulate the expression of CRC, and CRC can bind to the promoter and regulate the expression of AG. The results of this study suggest that both genes, CRC and AG, may regulate each other and promote the expression of downstream genes, thereby determining carpel development in sweet osmanthus. Genes that function downstream of CRC and AG and their mechanisms of action needs further investigation.

    • The flower buds and flower organs at the linggeng stage, xingyan stage, initial flowering stage, full flowering stage, and late full flowering stage of both the 'Huangchuan Jingui' and the 'Chenghong Dangui' cultivars were harvested from the campus of Henan University (China).

    • The experimental materials were washed with water and fixed with formalin-acetic acid-alcohol (FAA) solution. Flower buds of different growth parts and sizes were collected and cut into slices according to the conventional paraffin sectioning method[54], with a thickness of 8 μm. Sections were stained with alum-hematoxylin, sealed with neutral gum, and observed and photographed under an OLYMPUS BX60 microscope.

    • The anthers were peeled off and placed in a dry place to allow the pollen to disperse naturally. The pollen was spread on double-sided conductive adhesive, vacuum-dried, sprayed with gold coating, and observed and photographed under a JSM5600LV scanning electron microscope. The polar axis length (P), equatorial axis length (E), and germination groove length (L) were measured using a screen measurement tool, and the P/E and L/P values were calculated. In total, 30 pollen grains of each variety were measured and average values were obtained for each parameter. SPSS 22.0 statistical software was used to analyze the measurement data. Difference is considered significant at P < 0.05. There were three independent repeated experiments for the analysis.

    • The pollen grains at the initial flowering stage were taken and stained with 2,3,5-triphenyltetrazolium chloride (TTC) staining solution and fluorescein diacetate (FDA) staining solution, and images were obtained under a light microscope and a fluorescence microscope, respectively. The viability of pollen was examined by in vitro culture, using aniline blue staining and fluorescence microscopy for observation and photography.

    • RNA sequencing (RNA-seq) was performed by Frasergen (Wuhan, China). Transcriptome datasets were generated using the Illumina HiSeq 2,500 sequencing platform (San Diego, CA, USA). Differential expression analysis was performed using the DESeq2R package[55]. Expression levels of differentially expressed genes were computed using the following formula: FPKM = cDNA fragments / [mapped fragments (millions) × transcript length (kb)]. Fold Change ≥ 2 and FDR (False Discovery Rate) < 0.01 were used as differential expression genes screening criteria.

    • The cDNA sequences of AG and CRC were amplified and inserted into the pHBT vector (effectors). The promoter regions of AG (2.07 kb) and CRC (2.00 kb) were amplified and ligated into the pGreenII 0800-Luc vector (reporters). The primers for amplification are listed in Supplemental Table S1. EHA105 strains containing effectors and reporters were cotransformed into 5-week-old N. benthamiana leaves. Agroinfiltration was carried out following the method described by Han et al.[56]. The infiltrated plants were cultured at 23 °C under continuous lighting for 72 h. The LUC and REN activities were detected using a dual-luciferase assay kit (Solarbio, China). There were three independent repeated experiments for the analysis. Difference is considered significant at P < 0.05.

      • This research were supported by the Henan Province Major Research Fund of Public Welfare (No. 201300110900), National Natural Science Fund in China (No. U1604114), Basic Research Project of Key Scientific Research Program of higher education institutions in Henan Province (No. 20zx015).

      • The authors declare that they have no conflict of interest.

      • # These authors contributed equally: Yuanji Han, Yanxia He

      • Copyright: © 2023 by the author(s). Published by Maximum Academic Press, Fayetteville, GA. This article is an open access article distributed under Creative Commons Attribution License (CC BY 4.0), visit https://creativecommons.org/licenses/by/4.0/.
    Figure (5)  Table (1) References (56)
  • About this article
    Cite this article
    Han Y, He Y, Yue S, Guo B, Zhu Q, et al. 2023. Floral bud differentiation and mechanism underlying androdioecy of Osmanthus fragrans. Ornamental Plant Research 3:11 doi: 10.48130/OPR-2023-0011
    Han Y, He Y, Yue S, Guo B, Zhu Q, et al. 2023. Floral bud differentiation and mechanism underlying androdioecy of Osmanthus fragrans. Ornamental Plant Research 3:11 doi: 10.48130/OPR-2023-0011

Catalog

  • About this article

/

DownLoad:  Full-Size Img  PowerPoint
Return
Return