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Overexpression of HmWOX8 promotes callus proliferation and shoot regeneration by regulating hormone signaling and shoot development-related genes

  • # Authors contributed equally: Xueying Zhao, Along Chen

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  • Shoot regeneration capacity is essential for prosperous genetic transformation. Previous studies have shown that WUSCHEL-related homeobox (WOX) transcription factor plays a crucial role in callus growth, shoot regeneration, and root development. However, the mechanisms and functions of shoot regeneration related to WOX8 remain unclear. In the current research, the HmWOX8 gene was isolated from Hemerocallis middendorffii by RACE (Rapid-amplification of cDNA ends) technology. Overexpression of HmWOX8 improved callus proliferation and shoot regeneration ability of Arabidopsis and rice, whereas silencing HmWOX8 in H. middendorffii resulted in the inverse correlation. Transcriptome analysis revealed that HmWOX8 enhances the efficiency of callus proliferation and shoot regeneration through two different ways of regulation, including hormone signaling pathways and shoot development-related genes. (I) HmWOX8 regulates crosstalk among different hormone signaling pathways by activating and inhibiting the expression of different genes in these pathways, thus ensuring signal integration for efficient callus proliferation and shoot regeneration. (II) HmWOX8 can upregulate the expression level of shoot developmental genes, including WOX5/7, BBM, AIL5/7, PLT1, PIN6, CUC3, and SCR14/30, to regulate shoot emergence and outgrowth. In addition, Yeast two-hybrid assays and Bimolecular fluorescence complementation assay suggested that HmWOX8 directly interacts with HmCUC2, thereby promoting shoot regeneration. The present research improves the understanding of molecular mechanisms for HmWOX8-mediated regeneration and provides valuable gene resources for breeding programs to promote plant regeneration.
  • As a major staple crop, today maize accounts for approximately 40% of total worldwide cereal production (http://faostat.fao.org/). Since its domestication ~9,000 years ago from a subgroup of teosinte (Zea mays ssp. parviglumis) in the tropical lowlands of southwest Mexico[1], its cultivating area has greatly expanded, covering most of the world[2]. Human's breeding and utilization of maize have gone through several stages, from landraces, open-pollinated varieties (OPVs), double-cross hybrids (1930s-1950s) and since the middle 1950s, single-cross hybrids. Nowadays, global maize production is mostly provided by single-cross hybrids, which exhibit higher-yielding and better stress tolerance than OPVs and double-cross hybrids[3].

    Besides its agronomic importance, maize has also been used as a model plant species for genetic studies due to its out-crossing habit, large quantities of seeds produced and the availability of diverse germplasm. The abundant mutants of maize facilitated the development of the first genetic and cytogenetic maps of plants, and made it an ideal plant species to identify regulators of developmental processes[46]. Although initially lagging behind other model plant species (such as Arabidopsis and rice) in multi-omics research, the recent rapid development in sequencing and transformation technologies, and various new tools (such as CRISPR technologies, double haploids etc.) are repositioning maize research at the frontiers of plant research, and surely, it will continue to reveal fundamental insights into plant biology, as well as to accelerate molecular breeding for this vitally important crop[7, 8].

    During domestication from teosinte to maize, a number of distinguishing morphological and physiological changes occurred, including increased apical dominance, reduced glumes, suppression of ear prolificacy, increase in kernel row number, loss of seed shattering, nutritional changes etc.[9] (Fig. 1). At the genomic level, genome-wide genetic diversity was reduced due to a population bottleneck effect, accompanied by directional selection at specific genomic regions underlying agronomically important traits. Over a century ago, Beadle initially proposed that four or five genes or blocks of genes might be responsible for much of the phenotypic changes between maize and teosinte[10,11]. Later studies by Doebley et al. used teosinte–maize F2 populations to dissect several quantitative trait loci (QTL) to the responsible genes (such as tb1 and tga1)[12,13]. On the other hand, based on analysis of single-nucleotide polymorphisms (SNPs) in 774 genes, Wright et al.[14] estimated that 2%−4% of maize genes (~800−1,700 genes genome-wide) were selected during maize domestication and subsequent improvement. Taking advantage of the next-generation sequencing (NGS) technologies, Hufford et al.[15] conducted resequencing analysis of a set of wild relatives, landraces and improved maize varieties, and identified ~500 selective genomic regions during maize domestication. In a recent study, Xu et al.[16] conducted a genome-wide survey of 982 maize inbred lines and 190 teosinte accession. They identified 394 domestication sweeps and 360 adaptation sweeps. Collectively, these studies suggest that maize domestication likely involved hundreds of genomic regions. Nevertheless, much fewer domestication genes have been functionally studied so far.

    Figure 1.  Main traits of maize involved in domestication and improvement.

    During maize domestication, a most profound morphological change is an increase in apical dominance, transforming a multi-branched plant architecture in teosinte to a single stalked plant (terminated by a tassel) in maize. The tillers and long branches of teosinte are terminated by tassels and bear many small ears. Similarly, the single maize stalk bears few ears and is terminated by a tassel[9,12,17]. A series of landmark studies by Doebley et al. elegantly demonstrated that tb1, which encodes a TCP transcription factor, is responsible for this transformation[18, 19]. Later studies showed that insertion of a Hopscotch transposon located ~60 kb upstream of tb1 enhances the expression of tb1 in maize, thereby repressing branch outgrowth[20, 21]. Through ChIP-seq and RNA-seq analyses, Dong et al.[22] demonstrated that tb1 acts to regulate multiple phytohormone signaling pathways (gibberellins, abscisic acid and jasmonic acid) and sugar sensing. Moreover, several other domestication loci, including teosinte glume architecture1 (tga1), prol1.1/grassy tillers1, were identified as its putative targets. Elucidating the precise regulatory mechanisms of these loci and signaling pathways will be an interesting and rewarding area of future research. Also worth noting, studies showed that tb1 and its homologous genes in Arabidopsis (Branched1 or BRC1) and rice (FINE CULM1 or FC1) play a conserved role in repressing the outgrowth of axillary branches in both dicotyledon and monocotyledon plants[23, 24].

    Teosinte ears possess two ranks of fruitcase-enclosed kernels, while maize produces hundreds of naked kernels on the ear[13]. tga1, which encodes a squamosa-promoter binding protein (SBP) transcription factor, underlies this transformation[25]. It has been shown that a de novo mutation occurred during maize domestication, causing a single amino acid substitution (Lys to Asn) in the TGA1 protein, altering its binding activity to its target genes, including a group of MADS-box genes that regulate glume identity[26].

    Prolificacy, the number of ears per plants, is also a domestication trait. It has been shown that grassy tillers 1 (gt1), which encodes an HD-ZIP I transcription factor, suppresses prolificacy by promoting lateral bud dormancy and suppressing elongation of the later ear branches[27]. The expression of gt1 is induced by shading and requires the activity of tb1, suggesting that gt1 acts downstream of tb1 to mediate the suppressed branching activity in response to shade. Later studies mapped a large effect QTL for prolificacy (prol1.1) to a 2.7 kb 'causative region' upstream of the gt1gene[28]. In addition, a recent study identified a new QTL, qEN7 (for ear number on chromosome 7). Zm00001d020683, which encodes a putative INDETERMINATE DOMAIN (IDD) transcription factor, was identified as the likely candidate gene based on its expression pattern and signature of selection during maize improvement[29]. However, its functionality and regulatory relationship with tb1 and gt1 remain to be elucidated.

    Smaller leaf angle and thus more compact plant architecture is a desired trait for modern maize varieties. Tian et al.[30] used a maize-teosinte BC2S3 population and cloned two QTLs (Upright Plant Architecture1 and 2 [UPA1 and UPA2]) that regulate leaf angle. Interestingly, the authors showed that the functional variant of UPA2 is a 2-bp InDel located 9.5 kb upstream of ZmRAVL1, which encodes a B3 domain transcription factor. The 2-bp Indel flanks the binding site of the transcription factor Drooping Leaf1 (DRL1)[31], which represses ZmRAVL1 expression through interacting with Liguleless1 (LG1), a SBP-box transcription factor essential for leaf ligule and auricle development[32]. UPA1 encodes brassinosteroid C-6 oxidase1 (brd1), a key enzyme for biosynthesis of active brassinolide (BR). The teosinte-derived allele of UPA2 binds DRL1 more strongly, leading to lower expression of ZmRAVL1 and thus, lower expression of brd1 and BR levels, and ultimately smaller leaf angle. Notably, the authors demonstrated that the teosinte-derived allele of UPA2 confers enhanced yields under high planting densities when introgressed into modern maize varieties[30, 33].

    Maize plants exhibit salient vegetative phase change, which marks the vegetative transition from the juvenile stage to the adult stage, characterized by several changes in maize leaves produced before and after the transition, such as production of leaf epicuticular wax and epidermal hairs. Previous studies reported that Glossy15 (Gl15), which encodes an AP2-like transcription factor, promotes juvenile leaf identity and suppressing adult leaf identity. Ectopic overexpression of Gl15 causes delayed vegetative phase change and flowering, while loss-of-function gl15 mutant displayed earlier vegetative phase change[34]. In another study, Gl15 was identified as a major QTL (qVT9-1) controlling the difference in the vegetative transition between maize and teosinte. Further, it was shown that a pre-existing low-frequency standing variation, SNP2154-G, was selected during domestication and likely represents the causal variation underlying differential expression of Gl15, and thus the difference in the vegetative transition between maize and teosinte[35].

    A number of studies documented evidence that tassels replace upper ears1 (tru1) is a key regulator of the conversion of the male terminal lateral inflorescence (tassel) in teosinte to a female terminal inflorescence (ear) in maize. tru1 encodes a BTB/POZ ankyrin repeat domain protein, and it is directly targeted by tb1, suggesting their close regulatory relationship[36]. In addition, a number of regulators of maize inflorescence morphology, were also shown as selective targets during maize domestication, including ramosa1 (ra1)[37, 38], which encodes a putative transcription factor repressing inflorescence (the ear and tassel) branching, Zea Agamous-like1 (zagl1)[39], which encodes a MADS-box transcription factor regulating flowering time and ear size, Zea floricaula leafy2 (zfl2, homologue of Arabidopsis Leafy)[40, 41], which likely regulates ear rank number, and barren inflorescence2 (bif2, ortholog of the Arabidopsis serine/threonine kinase PINOID)[42, 43], which regulates the formation of spikelet pair meristems and branch meristems on the tassel. The detailed regulatory networks of these key regulators of maize inflorescence still remain to be further elucidated.

    Kernel row number (KRN) and kernel weight are two important determinants of maize yield. A number of domestication genes modulating KRN and kernel weight have been identified and cloned, including KRN1, KRN2, KRN4 and qHKW1. KRN4 was mapped to a 3-kb regulatory region located ~60 kb downstream of Unbranched3 (UB3), which encodes a SBP transcription factor and negatively regulates KRN through imparting on multiple hormone signaling pathways (cytokinin, auxin and CLV-WUS)[44, 45]. Studies have also shown that a harbinger TE in the intergenic region and a SNP (S35) in the third exon of UB3 act in an additive fashion to regulate the expression level of UB3 and thus KRN[46].

    KRN1 encodes an AP2 transcription factor that pleiotropically affects plant height, spike density and grain size of maize[47], and is allelic to ids1/Ts6 (indeterminate spikelet 1/Tassel seed 6)[48]. Noteworthy, KRN1 is homologous to the wheat domestication gene Q, a major regulator of spike/spikelet morphology and grain threshability in wheat[49].

    KRN2 encodes a WD40 domain protein and it negatively regulates kernel row number[50]. Selection in a ~700-bp upstream region (containing the 5’UTR) of KRN2 during domestication resulted in reduced expression and thus increased kernel row number. Interestingly, its orthologous gene in rice, OsKRN2, was shown also a selected gene during rice domestication to negatively regulate secondary panicle branches and thus grain number. These observations suggest convergent selection of yield-related genes occurred during parallel domestication of cereal crops.

    qHKW1 is a major QTL for hundred-kernel weight (HKW)[51]. It encodes a CLAVATA1 (CLV1)/BARELY ANY MERISTEM (BAM)-related receptor kinase-like protein positively regulating HKW. A 8.9 Kb insertion in its promoter region was find to enhance its expression, leading to enhanced HKW[52]. In addition, Chen et al.[53] reported cloning of a major QTL for kernel morphology, qKM4.08, which encodes ZmVPS29, a retromer complex component. Sequencing and association analysis revealed that ZmVPS29 was a selective target during maize domestication. They authors also identified two significant polymorphic sites in its promoter region significantly associated with the kernel morphology. Moreover, a strong selective signature was detected in ZmSWEET4c during maize domestication. ZmSWEET4c encodes a hexose transporter protein functioning in sugar transport across the basal endosperm transfer cell layer (BETL) during seed filling[54]. The favorable alleles of these genes could serve as valuable targets for genetic improvement of maize yield.

    In a recent effort to more systematically analyze teosinte alleles that could contribute to yield potential of maize, Wang et al.[55] constructed four backcrossed maize-teosinte recombinant inbred line (RIL) populations and conducted detailed phenotyping of 26 agronomic traits under five environmental conditions. They identified 71 QTL associated with 24 plant architecture and yield related traits through inclusive composite interval mapping. Interestingly, they identified Zm00001eb352570 and Zm00001eb352580, both encode ethylene-responsive transcription factors, as two key candidate genes regulating ear height and the ratio of ear to plant height. Chen et al.[56] constructed a teosinte nested association mapping (TeoNAM) population, and performed joint-linkage mapping and GWAS analyses of 22 domestication and agronomic traits. They identified the maize homologue of PROSTRATE GROWTH1, a rice domestication gene controlling the switch from prostrate to erect growth, is also a QTL associated with tillering in teosinte and maize. Additionally, they also detected multiple QTL for days-to-anthesis (such as ZCN8 and ZmMADS69) and other traits (such as tassel branch number and tillering) that could be exploited for maize improvement. These lines of work highlight again the value of mining the vast amounts of superior alleles hidden in teosinte for future maize genetic improvement.

    Loss of seed shattering was also a key trait of maize domestication, like in other cereals. shattering1 (sh1), which encodes a zinc finger and YABBY domain protein regulating seed shattering. Interesting, sh1 was demonstrated to undergo parallel domestication in several cereals, including rice, maize, sorghum, and foxtail millet[57]. Later studies showed that the foxtail millet sh1 gene represses lignin biosynthesis in the abscission layer, and that an 855-bp Harbinger transposable element insertion in sh1 causes loss of seed shattering in foxtail millet[58].

    In addition to morphological traits, a number of physiological and nutritional related traits have also been selected during maize domestication. Based on survey of the nucleotide diversity, Whitt et al.[59] reported that six genes involved in starch metabolism (ae1, bt2, sh1, sh2, su1 and wx1) are selective targets during maize domestication. Palaisa et al.[60] reported selection of the Y1 gene (encoding a phytoene synthase) for increased nutritional value. Karn et al.[61] identified two, three, and six QTLs for starch, protein and oil respectively and showed that teosinte alleles can be exploited for the improvement of kernel composition traits in modern maize germplasm. Fan et at.[62] reported a strong selection imposed on waxy (wx) in the Chinese waxy maize population. Moreover, a recent exciting study reported the identification of a teosinte-derived allele of teosinte high protein 9 (Thp9) conferring increased protein level and nitrogen utilization efficiency (NUE). It was further shown that Thp9 encodes an asparagine synthetase 4 and that incorrect splicing of Thp9-B73 transcripts in temperate maize varieties is responsible for its diminished expression, and thus reduced NUE and protein content[63].

    Teosintes is known to confer superior disease resistance and adaptation to extreme environments (such as low phosphorus and high salinity). de Lange et al. and Lennon et al.[6466] reported the identification of teosinte-derived QTLs for resistance to gray leaf spot and southern leaf blight in maize. Mano & Omori reported that teosinte-derived QTLs could confer flooding tolerance[67]. Feng et al.[68] identified four teosinte-derived QTL that could improve resistance to Fusarium ear rot (FER) caused by Fusarium verticillioides. Recently, Wang et al.[69] reported a MYB transcription repressor of teosinte origin (ZmMM1) that confers resistance to northern leaf blight (NLB), southern corn rust (SCR) and gray leaf spot (GLS) in maize, while Zhang et al.[70] reported the identification of an elite allele of SNP947-G ZmHKT1 (encoding a sodium transporter) derived from teosinte can effectively improve salt tolerance via exporting Na+ from the above-ground plant parts. Gao et al.[71] reported that ZmSRO1d-R can regulate the balance between crop yield and drought resistance by increasing the guard cells' ROS level, and it underwent selection during maize domestication and breeding. These studies argue for the need of putting more efforts to tapping into the genetic resources hidden in the maize’s wild relatives. The so far cloned genes involved in maize domestication are summarized in Table 1. Notably, the enrichment of transcription factors in the cloned domestication genes highlights a crucial role of transcriptional re-wiring in maize domestication.

    Table 1.  Key domestication genes cloned in maize.
    GenePhenotypeFunctional annotationSelection typeCausative changeReferences
    tb1Plant architectureTCP transcription factorIncreased expression~60 kb upstream of tb1 enhancing expression[1822]
    tga1Hardened fruitcaseSBP-domain transcription factorProtein functionA SNP in exon (K-N)[25, 26]
    gt1Plant architectureHomeodomain leucine zipperIncreased expressionprol1.1 in 2.7 kb upstream of the promoter region increasing expression[27, 28]
    Zm00001d020683Plant architectureINDETERMINATE DOMAIN transcription factorProtein functionUnknown[29]
    UPA1Leaf angleBrassinosteroid C-6 oxidase1Protein functionUnknown[30]
    UPA2Leaf angleB3 domain transcription factorIncreased expressionA 2 bp indel in 9.5 kb upstream of ZmRALV1[30]
    Gl15Vegetative phase changeAP2-like transcription factorAltered expressionSNP2154: a stop codon (G-A)[34, 35]
    tru1Plant architectureBTB/POZ ankyrin repeat proteinIncreased expressionUnknown[36]
    ra1Inflorescence architectureTranscription factorAltered expressionUnknown[37, 38]
    zflPlant architectureTranscription factorAltered expressionUnknown[40, 41]
    UB3Kernel row numberSBP-box transcription factorAltered expressionA TE in the intergenic region;[4446]
    SNP (S35): third exon of UB3
    (A-G) increasing expression of UB3 and KRN
    KRN1/ids1/Ts6Kernel row numberAP2 Transcription factorIncreased expressionUnknown[47, 48]
    KRN2Kernel row numberWD40 domainDecreased expressionUnknown[50]
    qHKW1Kernel row weightCLV1/BAM-related receptor kinase-like proteinIncreased expression8.9 kb insertion upstream of HKW[51, 52]
    ZmVPS29Kernel morphologyA retromer complex componentProtein functionTwo SNPs (S-1830 and S-1558) in the promoter of ZmVPS29[53]
    ZmSWEET4cSeed fillingHexose transporterProtein functionUnknown[54]
    ZmSh1ShatteringA zinc finger and YABBY transcription factorProtein functionUnknown[57, 58]
    Thp9Nutrition qualityAsparagine synthetase 4 enzymeProtein functionA deletion in 10th intron of Thp9 reducing NUE and protein content[63]
    ZmMM1Biotic stressMYB Transcription repressorProtein functionUnknown[69]
    ZmHKT1Abiotic stressA sodium transporterProtein functionSNP947-G: a nonsynonymous variation increasing salt tolerance[70]
    ZmSRO1d-RDrought resistance and productionPolyADP-ribose polymerase and C-terminal RST domainProtein functionThree non-synonymous variants: SNP131 (A44G), SNP134 (V45A) and InDel433[71]
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    After its domestication from its wild progenitor teosinte in southwestern Mexico in the tropics, maize has now become the mostly cultivated crop worldwide owing to its extensive range expansion and adaptation to diverse environmental conditions (such as temperature and day length). A key prerequisite for the spread of maize from tropical to temperate regions is reduced photoperiod sensitivity[72]. It was recently shown that CENTRORADIALIS 8 (ZCN8), an Flowering Locus T (FT) homologue, underlies a major quantitative trait locus (qDTA8) for flowering time[73]. Interestingly, it has been shown that step-wise cis-regulatory changes occurred in ZCN8 during maize domestication and post-domestication expansion. SNP-1245 is a target of selection during early maize domestication for latitudinal adaptation, and after its fixation, selection of InDel-2339 (most likely introgressed from Zea mays ssp. Mexicana) likely contributed to the spread of maize from tropical to temperate regions[74].

    ZCN8 interacts with the basic leucine zipper transcription factor DLF1 (Delayed flowering 1) to form the florigen activation complex (FAC) in maize. Interestingly, DFL1 was found to underlie qLB7-1, a flowering time QTL identified in a BC2S3 population of maize-teosinte. Moreover, it was shown that DLF1 directly activates ZmMADS4 and ZmMADS67 in the shoot apex to promote floral transition[75]. In addition, ZmMADS69 underlies the flowering time QTL qDTA3-2 and encodes a MADS-box transcription factor. It acts to inhibit the expression of ZmRap2.7, thereby relieving its repression on ZCN8 expression and causing earlier flowering. Population genetic analyses showed that DLF1, ZmMADS67 and ZmMADS69 are all targets of artificial selection and likely contributed to the spread of maize from the tropics to temperate zones[75, 76].

    In addition, a few genes regulating the photoperiod pathway and contributing to the acclimation of maize to higher latitudes in North America have been cloned, including Vgt1, ZmCCT (also named ZmCCT10), ZmCCT9 and ZmELF3.1. Vgt1 was shown to act as a cis-regulatory element of ZmRap2.7, and a MITE TE located ~70 kb upstream of Vgt1 was found to be significantly associated with flowering time and was a major target for selection during the expansion of maize to the temperate and high-latitude regions[7779]. ZmCCT is another major flowering-time QTL and it encodes a CCT-domain protein homologous to rice Ghd7[80]. Its causal variation is a 5122-bp CACTA-like TE inserted ~2.5 kb upstream of ZmCCT10[72, 81]. ZmCCT9 was identified a QTL for days to anthesis (qDTA9). A Harbinger-like TE located ~57 kb upstream of ZmCCT9 showed the most significant association with DTA and thus believed to be the causal variation[82]. Notably, the CATCA-like TE of ZmCCT10 and the Harbinger-like TE of ZmCCT9 are not observed in surveyed teosinte accessions, hinting that they are de novo mutations occurred after the initial domestication of maize[72, 82]. ZmELF3.1 was shown to underlie the flowering time QTL qFT3_218. It was demonstrated that ZmELF3.1 and its homolog ZmELF3.2 can form the maize Evening Complex (EC) through physically interacting with ZmELF4.1/ZmELF4.2, and ZmLUX1/ZmLUX2. Knockout mutants of Zmelf3.1 and Zmelf3.1/3.2 double mutant presented delayed flowering under both long-day and short-day conditions. It was further shown that the maize EC promote flowering through repressing the expression of several known flowering suppressor genes (e.g., ZmCCT9, ZmCCT10, ZmCOL3, ZmPRR37a and ZmPRR73), and consequently alleviating their inhibition on several maize florigen genes (ZCN8, ZCN7 and ZCN12). Insertion of two closely linked retrotransposon elements upstream of the ZmELF3.1 coding region increases the expression of ZmELF3.1, thus promoting flowering[83]. The increase frequencies of the causal TEs in Vgt1, ZmCCT10, ZmCCT9 and ZmELF3.1 in temperate maize compared to tropical maize highlight a critical role of these genes during the spread and adaptation of maize to higher latitudinal temperate regions through promoting flowering under long-day conditions[72,8183].

    In addition, Barnes et al.[84] recently showed that the High Phosphatidyl Choline 1 (HPC1) gene, which encodes a phospholipase A1 enzyme, contributed to the spread of the initially domesticated maize from the warm Mexican southwest to the highlands of Mexico and South America by modulating phosphatidylcholine levels. The Mexicana-derived allele harbors a polymorphism and impaired protein function, leading to accelerated flowering and better fitness in highlands.

    Besides the above characterized QTLs and genes, additional genetic elements likely also contributed to the pre-Columbia spreading of maize. Hufford et al.[85] proposed that incorporation of mexicana alleles into maize may helped the expansion of maize to the highlands of central Mexico based on detection of bi-directional gene flow between maize and Mexicana. This proposal was supported by a recent study showing evidence of introgression for over 10% of the maize genome from the mexicana genome[86]. Consistently, Calfee et al.[87] found that sequences of mexicana ancestry increases in high-elevation maize populations, supporting the notion that introgression from mexicana facilitating adaptation of maize to the highland environment. Moreover, a recent study examined the genome-wide genetic diversity of the Zea genus and showed that dozens of flowering-related genes (such as GI, BAS1 and PRR7) are associated with high-latitude adaptation[88]. These studies together demonstrate unequivocally that introgression of genes from Mexicana and selection of genes in the photoperiod pathway contributed to the spread of maize to the temperate regions.

    The so far cloned genes involved in pre-Columbia spread of maize are summarized in Fig. 2 and Table 2.

    Figure 2.  Genes involved in Pre-Columbia spread of maize to higher latitudes and the temperate regions. The production of world maize in 2020 is presented by the green bar in the map from Ritchie et al. (2023). Ritchie H, Rosado P, and Roser M. 2023. "Agricultural Production". Published online at OurWorldInData.org. Retrieved from: 'https:ourowrldindata.org/agricultural-production' [online Resource].
    Table 2.  Flowering time related genes contributing to Pre-Columbia spread of maize.
    GeneFunctional annotationCausative changeReferences
    ZCN8Florigen proteinSNP-1245 and Indel-2339 in promoter[73, 74]
    DLF1Basic leucine zipper transcription factorUnknown[75]
    ZmMADS69MADS-box transcription factorUnknown[76]
    ZmRap2.7AP2-like transcription factorMITE TE inserted ~70 kb upstream[7779]
    ZmCCTCCT-domain protein5122-bp CACTA-like TE inserted ~2.5 kb upstream[72,81]
    ZmCCT9CCT transcription factorA harbinger-like element at 57 kb upstream[82]
    ZmELF3.1Unknownwo retrotransposons in the promote[84]
    HPC1Phospholipase A1 enzymUnknown[83]
    ZmPRR7UnknownUnknown[88]
    ZmCOL9CO-like-transcription factorUnknown[88]
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    Subsequent to domestication ~9,000 years ago, maize has been continuously subject to human selection during the post-domestication breeding process. Through re-sequencing analysis of 35 improved maize lines, 23 traditional landraces and 17 wild relatives, Hufford et al.[15] identified 484 and 695 selective sweeps during maize domestication and improvement, respectively. Moreover, they found that about a quarter (23%) of domestication sweeps (107) were also selected during improvement, indicating that a substantial portion of the domestication loci underwent continuous selection during post-domestication breeding.

    Genetic improvement of maize culminated in the development of high planting density tolerant hybrid maize to increase grain yield per unit land area[89, 90]. To investigate the key morphological traits that have been selected during modern maize breeding, we recently conducted sequencing and phenotypic analyses of 350 elite maize inbred lines widely used in the US and China over the past few decades. We identified four convergently improved morphological traits related to adapting to increased planting density, i.e., reduced leaf angle, reduced tassel branch number (TBN), reduced relative plant height (EH/PH) and accelerated flowering. Genome-wide Association Study (GWAS) identified a total of 166 loci associated with the four selected traits, and found evidence of convergent increases in allele frequency at putatively favorable alleles for the identified loci. Moreover, genome scan using the cross-population composite likelihood ratio approach (XP-CLR) identified a total of 1,888 selective sweeps during modern maize breeding in the US and China. Gene ontology analysis of the 5,356 genes encompassed in the selective sweeps revealed enrichment of genes related to biosynthesis or signaling processes of auxin and other phytohormones, and in responses to light, biotic and abiotic stresses. This study provides a valuable resource for mining genes regulating morphological and physiological traits underlying adaptation to high-density planting[91].

    In another study, Li et al.[92] identified ZmPGP1 (ABCB1 or Br2) as a selected target gene during maize domestication and genetic improvement. ZmPGP1 is involved in auxin polar transport, and has been shown to have a pleiotropic effect on plant height, stalk diameter, leaf length, leaf angle, root development and yield. Sequence and phenotypic analyses of ZmPGP1 identified SNP1473 as the most significant variant for kernel length and ear grain weight and that the SNP1473T allele is selected during both the domestication and improvement processes. Moreover, the authors identified a rare allele of ZmPGP1 carrying a 241-bp deletion in the last exon, which results in significantly reduced plant height and ear height and increased stalk diameter and erected leaves, yet no negative effect on yield[93], highlighting a potential utility in breeding high-density tolerant maize cultivars.

    Shade avoidance syndrome (SAS) is a set of adaptive responses triggered when plants sense a reduction in the red to far-red light (R:FR) ratio under high planting density conditions, commonly manifested by increased plant height (and thus more prone to lodging), suppressed branching, accelerated flowering and reduced resistance to pathogens and pests[94, 95]. High-density planting could also cause extended anthesis-silking interval (ASI), reduced tassel size and smaller ear, and even barrenness[96, 97]. Thus, breeding of maize cultivars of attenuated SAS is a priority for adaptation to increased planting density.

    Extensive studies have been performed in Arabidopsis to dissect the regulatory mechanism of SAS and this topic has been recently extensively reviewed[98]. We recently showed that a major signaling mechanism regulating SAS in Arabidopsis is the phytochrome-PIFs module regulates the miR156-SPL module-mediated aging pathway[99]. We proposed that in maize there might be a similar phytochrome-PIFs-miR156-SPL regulatory pathway regulating SAS and that the maize SPL genes could be exploited as valuable targets for genetic improvement of plant architecture tailored for high-density planting[100].

    In support of this, it has been shown that the ZmphyBs (ZmphyB1 and ZmphyB2), ZmphyCs (ZmphyC1 and ZmphyC2) and ZmPIFs are involved in regulating SAS in maize[101103]. In addition, earlier studies have shown that as direct targets of miR156s, three homologous SPL transcription factors, UB2, UB3 and TSH4, regulate multiple agronomic traits including vegetative tillering, plant height, tassel branch number and kernel row number[44, 104]. Moreover, it has been shown that ZmphyBs[101, 105] and ZmPIF3.1[91], ZmPIF4.1[102] and TSH4[91] are selective targets during modern maize breeding (Table 3).

    Table 3.  Selective genes underpinning genetic improvement during modern maize breeding.
    GenePhenotypeFunctional annotationSelection typeCausative changeReferences
    ZmPIF3.1Plant heightBasic helix-loop-helix transcription factorIncreased expressionUnknown[91]
    TSH4Tassel branch numberTranscription factorAltered expressionUnknown[91]
    ZmPGP1Plant architectureATP binding cassette transporterAltered expressionA 241 bp deletion in the last exon of ZmPGP1[92, 93]
    PhyB2Light signalPhytochrome BAltered expressionA 10 bp deletion in the translation start site[101]
    ZmPIF4.1Light signalBasic helix-loop-helix transcription factorAltered expressionUnknown[102]
    ZmKOB1Grain yieldGlycotransferase-like proteinProtein functionUnknown[121]
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    In a recent study to dissect the signaling process regulating inflorescence development in response to the shade signal, Kong et al.[106] compared the gene expression changes along the male and female inflorescence development under simulated shade treatments and normal light conditions, and identified a large set of genes that are co-regulated by developmental progression and simulated shade treatments. They found that these co-regulated genes are enriched in plant hormone signaling pathways and transcription factors. By network analyses, they found that UB2, UB3 and TSH4 act as a central regulatory node controlling maize inflorescence development in response to shade signal, and their loss-of-function mutants exhibit reduced sensitivity to simulated shade treatments. This study provides a valuable genetic source for mining and manipulating key shading-responsive genes for improved tassel and ear traits under high density planting conditions.

    Nowadays, global maize production is mostly provided by hybrid maize, which exhibits heterosis (or hybrid vigor) in yields and stress tolerance over open-pollinated varieties[3]. Hybrid maize breeding has gone through several stages, from the 'inbred-hybrid method' stage by Shull[107] and East[108] in the early twentieth century, to the 'double-cross hybrids' stage (1930s−1950s) by Jones[109], and then the 'single-cross hybrids' stage since the 1960s. Since its development, single-cross hybrid was quickly adopted globally due to its superior heterosis and easiness of production[3].

    Single-cross maize hybrids are produced from crossing two unrelated parental inbred lines (female × male) belonging to genetically distinct pools of germplasm, called heterotic groups. Heterotic groups allow better exploitation of heterosis, since inter-group hybrids display a higher level of heterosis than intra-group hybrids. A specific pair of female and male heterotic groups expressing pronounced heterosis is termed as a heterotic pattern[110, 111]. Initially, the parental lines were derived from a limited number of key founder inbred lines and empirically classified into different heterotic groups (such as SSS and NSS)[112]. Over time, they have expanded dramatically, accompanied by formation of new 'heterotic groups' (such as Iodent, PA and PB). Nowadays, Stiff Stalk Synthetics (SSS) and PA are generally used as FHGs (female heterotic groups), while Non Stiff Stalk (NSS), PB and Sipingtou (SPT) are generally used as the MHGs (male heterotic groups) in temperate hybrid maize breeding[113].

    With the development of molecular biology, various molecular markers, ranging from RFLPs, SSRs, and more recently high-density genome-wide SNP data have been utilized to assign newly developed inbred lines into various heterotic groups, and to guide crosses between heterotic pools to produce the most productive hybrids[114116]. Multiple studies with molecular markers have suggested that heterotic groups have diverged genetically over time for better heterosis[117120]. However, there has been a lack of a systematic assessment of the effect and contribution of breeding selection on phenotypic improvement and the underlying genomic changes of FHGs and MHGs for different heterotic patterns on a population scale during modern hybrid maize breeding.

    To systematically assess the phenotypic improvement and the underlying genomic changes of FHGs and MHGs during modern hybrid maize breeding, we recently conducted re-sequencing and phenotypic analyses of 21 agronomic traits for a panel of 1,604 modern elite maize lines[121]. Several interesting observations were made: (1) The MHGs experienced more intensive selection than the FMGs during the progression from era I (before the year 2000) to era II (after the year 2000). Significant changes were observed for 18 out of 21 traits in the MHGs, but only 10 of the 21 traits showed significant changes in the FHGs; (2) The MHGs and FHGs experienced both convergent and divergent selection towards different sets of agronomic traits. Both the MHGs and FHGs experienced a decrease in flowering time and an increase in yield and plant architecture related traits, but three traits potentially related to seed dehydration rate were selected in opposite direction in the MHGs and FHGs. GWAS analysis identified 4,329 genes associated with the 21 traits. Consistent with the observed convergent and divergent changes of different traits, we observed convergent increase for the frequencies of favorable alleles for the convergently selected traits in both the MHGs and FHGs, and anti-directional changes for the frequencies of favorable alleles for the oppositely selected traits. These observations highlight a critical contribution of accumulation of favorable alleles to agronomic trait improvement of the parental lines of both FHGs and MHGs during modern maize breeding.

    Moreover, FST statistics showed increased genetic differentiation between the respective MHGs and FHGs of the US_SS × US_NSS and PA × SPT heterotic patterns from era I to era II. Further, we detected significant positive correlations between the number of accumulated heterozygous superior alleles of the differentiated genes with increased grain yield per plant and better parent heterosis, supporting a role of the differentiated genes in promoting maize heterosis. Further, mutational and overexpressional studies demonstrated a role of ZmKOB1, which encodes a putative glycotransferase, in promoting grain yield[121]. While this study complemented earlier studies on maize domestication and variation maps in maize, a pitfall of this study is that variation is limited to SNP polymorphisms. Further exploitation of more variants (Indels, PAVs, CNVs etc.) in the historical maize panel will greatly deepen our understanding of the impact of artificial selection on the maize genome, and identify valuable new targets for genetic improvement of maize.

    The ever-increasing worldwide population and anticipated climate deterioration pose a great challenge to global food security and call for more effective and precise breeding methods for crops. To accommodate the projected population increase in the next 30 years, it is estimated that cereal production needs to increase at least 70% by 2050 (FAO). As a staple cereal crop, breeding of maize cultivars that are not only high-yielding and with superior quality, but also resilient to environmental stresses, is essential to meet this demand. The recent advances in genome sequencing, genotyping and phenotyping technologies, generation of multi-omics data (including genomic, phenomic, epigenomic, transcriptomic, proteomic, and metabolomic data), creation of novel superior alleles by genome editing, development of more efficient double haploid technologies, integrating with machine learning and artificial intelligence are ushering the transition of maize breeding from the Breeding 3.0 stage (biological breeding) into the Breeding 4.0 stage (intelligent breeding)[122, 123]. However, several major challenges remain to be effectively tackled before such a transition could be implemented. First, most agronomic traits of maize are controlled by numerous small-effect QTL and complex genotype-environment interactions (G × E). Thus, elucidating the contribution of the abundant genetic variation in the maize population to phenotypic plasticity remains a major challenge in the post-genomic era of maize genetics and breeding. Secondly, most maize cultivars cultivated nowadays are hybrids that exhibit superior heterosis than their parental lines. Hybrid maize breeding involves the development of elite inbred lines with high general combining ability (GCA) and specific combining ability (SCA) that allows maximal exploitation of heterosis. Despite much effort to dissect the mechanisms of maize heterosis, the molecular basis of maize heterosis is still a debated topic[124126]. Thirdly, only limited maize germplasm is amenable to genetic manipulation (genetic transformation, genome editing etc.), which significantly hinders the efficiency of genetic improvement. Development of efficient genotype-independent transformation procedure will greatly boost maize functional genomic research and breeding. Noteworthy, the Smart Corn System recently launched by Bayer is promised to revolutionize global corn production in the coming years. At the heart of the new system is short stature hybrid corn (~30%−40% shorter than traditional hybrids), which offers several advantages: sturdier stems and exceptional lodging resistance under higher planting densities (grow 20%−30% more plants per hectare), higher and more stable yield production per unit land area, easier management and application of plant protection products, better use of solar energy, water and other natural resources, and improved greenhouse gas footprint[127]. Indeed, a new age of maize green revolution is yet to come!

    This work was supported by grants from the Key Research and Development Program of Guangdong Province (2022B0202060005), National Natural Science Foundation of China (32130077) and Hainan Yazhou Bay Seed Lab (B21HJ8101). We thank Professors Hai Wang (China Agricultural University) and Jinshun Zhong (South China Agricultural University) for valuable comments and helpful discussion on the manuscript. We apologize to authors whose excellent work could not be cited due to space limitations.

  • The authors declare that they have no conflict of interest. Haiyang Wang is an Editorial Board member of Seed Biology who was blinded from reviewing or making decisions on the manuscript. The article was subject to the journal's standard procedures, with peer-review handled independently of this Editorial Board member and his research groups.

  • Supplemental Table S1 qRT-PCR primer sequences.
    Supplemental Table S2 qRT-PCR primer sequences.
    Supplemental Fig. S1 PCR identification of Arabidopsis transformed with HmWOX8 gene.
    Supplemental Fig. S2 GUS staining verification, and HmWOX8 gene relative expression in WT and OE of rice.
    Supplemental Fig. S3 GO enrichment analysis of differentially expressed genes (DEGs) between HmWOX8-OE and WT
    Supplemental Fig. S4 qRT-PCR validation of DEGs in WT and HmWOX8-OE. The bars represented standard deviation.
  • [1]

    Bano MA, Khan J. 2022. The effect of Pseudomonas putida and spermine on growth and bioactive metabolites of Hemerocallis fulva L. leaves. Russian Journal of Plant Physiology 69:132

    doi: 10.1134/S1021443722060024

    CrossRef   Google Scholar

    [2]

    Rodriguez-Enriquez MJ, Grant-Downton RT. 2013. A new day dawning: Hemerocallis (daylily) as a future model organism. AoB Plants 5:pls055

    doi: 10.1093/aobpla/pls055

    CrossRef   Google Scholar

    [3]

    Sun X, Wu R. 2016. Recent advances in Hemerocallis. Journal of Henan Agricultural Sciences 45:7−11,18

    doi: 10.15933/j.cnki.1004-3268.2016.04.002

    CrossRef   Google Scholar

    [4]

    Duan L, Li Y, Liu X, Dong Y, Yu S, et al. 2023. Research progress on breeding of new Hemerocallis varieties at home and abroad. Journal of Nuclear Agricultural Sciences 37:730−39

    doi: 10.11869/j.issn.1000-8551.2023.04.0730

    CrossRef   Google Scholar

    [5]

    Ikeuchi M, Favero DS, Sakamoto Y, Iwase A, Coleman D, et al. 2019. Molecular mechanisms of plant regeneration. Annual Review of Plant Biology 70:377−406

    doi: 10.1146/annurev-arplant-050718-100434

    CrossRef   Google Scholar

    [6]

    Xu L, Huang H. 2014. Genetic and epigenetic controls of plant regeneration. Current Topics in Developmental Biology 108:1−33

    doi: 10.1016/B978-0-12-391498-9.00009-7

    CrossRef   Google Scholar

    [7]

    Ikeuchi M, Ogawa Y, Iwase A, Sugimoto K. 2016. Plant regeneration: cellular origins and molecular mechanisms. Development 143:1442−51

    doi: 10.1242/dev.134668

    CrossRef   Google Scholar

    [8]

    Sang Y, Cheng Z, Zhang X. 2018. Plant stem cells and de novo organogenesis. New Phytologist 218:1334−39

    doi: 10.1111/nph.15106

    CrossRef   Google Scholar

    [9]

    Shin J, Bae S, Seo PJ. 2020. De novo shoot organogenesis during plant regeneration. Journal of Experimental Botany 71:63−72

    doi: 10.1093/jxb/erz395

    CrossRef   Google Scholar

    [10]

    Lee HG, Jang SY, Jie EY, Choi SH, Park OS, et al. 2023. Adenosine monophosphate enhances callus regeneration competence for de novo plant organogenesis. Molecular Plant 16:1867−70

    doi: 10.1016/j.molp.2023.10.004

    CrossRef   Google Scholar

    [11]

    Cheng Z, Wang L, Sun W, Zhang Y, Zhou C, et al. 2012. Pattern of auxin and cytokinin responses for shoot meristem induction results from the regulation of cytokinin biosynthesis by AUXIN RESPONSE FACTOR3. Plant Physiology 161:240−51

    doi: 10.1104/pp.112.203166

    CrossRef   Google Scholar

    [12]

    Ckurshumova W, Smirnova T, Marcos D, Zayed Y, Berleth T. 2014. Irrepressible MONOPTEROS/ARF5 promotes de novo shoot formation. New Phytologist 204:556−66

    doi: 10.1111/nph.13014

    CrossRef   Google Scholar

    [13]

    Liu K, Yang A, Yan J, Liang Z, Yuan G, et al. 2023. MdAIL5 overexpression promotes apple adventitious shoot regeneration by regulating hormone signaling and activating the expression of shoot development-related genes. Horticulture Research 10:uhad198

    doi: 10.1093/hr/uhad198

    CrossRef   Google Scholar

    [14]

    Fan M, Xu C, Xu K, Hu Y. 2012. LATERAL ORGAN BOUNDARIES DOMAIN transcription factors direct callus formation in Arabidopsis regeneration. Cell Research 22:1169−80

    doi: 10.1038/cr.2012.63

    CrossRef   Google Scholar

    [15]

    Zhao D, Wang Y, Feng C, Wei Y, Peng X, et al. 2020. Overexpression of MsGH3.5 inhibits shoot and root development through the auxin and cytokinin pathways in apple plants. The Plant Journal 103:166−83

    doi: 10.1111/tpj.14717

    CrossRef   Google Scholar

    [16]

    Mao J, Ma D, Niu C, Ma X, Li K, et al. 2022. Transcriptome analysis reveals the regulatory mechanism by which MdWOX11 suppresses adventitious shoot formation in apple. Horticulture Research 9:uhac080

    doi: 10.1093/hr/uhac080

    CrossRef   Google Scholar

    [17]

    Wang K, Shi L, Liang X, Zhao P, Wang W, et al. 2022. The gene TaWOX5 overcomes genotype dependency in wheat genetic transformation. Nature Plants 8:110−17

    doi: 10.1038/s41477-021-01085-8

    CrossRef   Google Scholar

    [18]

    Werner T, Motyka V, Laucou V, Smets R, Van Onckelen H, et al. 2003. Cytokinin-deficient transgenic Arabidopsis plants show multiple developmental alterations indicating opposite functions of cytokinins in the regulation of shoot and root meristem activity. The Plant Cell 15:2532−50

    doi: 10.1105/tpc.014928

    CrossRef   Google Scholar

    [19]

    Zhang T, Lian H, Zhou C, Xu L, Jiao Y, et al. 2017. A two-step model for de novo activation of WUSCHEL during plant shoot regeneration. The Plant Cell 29:1073−87

    doi: 10.1105/tpc.16.00863

    CrossRef   Google Scholar

    [20]

    Buechel S, Leibfried A, To JPC, Zhao Z, Andersen SU, et al. 2010. Role of A-type ARABIDOPSIS RESPONSE REGULATORS in meristem maintenance and regeneration. European Journal of Cell Biology 89:279−84

    doi: 10.1016/j.ejcb.2009.11.016

    CrossRef   Google Scholar

    [21]

    Chatfield SP, Raizada MN. 2008. Ethylene and shoot regeneration: hookless1 modulates de novo shoot organogenesis in Arabidopsis thaliana. Plant Cell Reports 27:655−66

    doi: 10.1007/s00299-007-0496-3

    CrossRef   Google Scholar

    [22]

    Chen J, Tomes S, Gleav AP, Hall W, Luo Z, et al. 2022. Significant improvement of apple (Malus domestica Borkh.) transgenic plant production by pre-transformation with a Baby boom transcription factor. Horticulture Research 9:uhab014

    doi: 10.1093/hr/uhab014

    CrossRef   Google Scholar

    [23]

    Druege U, Franken P, Hajirezaei MR. 2016. Plant hormone homeostasis, signaling, and function during adventitious root formation in cuttings. Frontiers in Plant Science 7:381

    doi: 10.3389/fpls.2016.00381

    CrossRef   Google Scholar

    [24]

    Wang X, Ma H, Cao D. 2014. Establishment of regeneration system of Hemeroallis middendorfii Trautv. et Mey 'Sweet Treasure'. Journal of Gansu Agricultural University 49:136−42

    doi: 10.3969/j.issn.1003-4315.2014.04.024

    CrossRef   Google Scholar

    [25]

    Zuo G, Cheng X, Yu J, Yin L, Hou F, et al. 2022. Callus induction and plant regeneration from the scape of Hemerocallis citrina. Journal of Agricultural University of Hebei 45:37−42

    doi: 10.13320/j.cnki.jauh.2022.0058

    CrossRef   Google Scholar

    [26]

    Zuo G, Li K, Guo Y, Niu X, Yin L, et al. 2024. Development and optimization of a rapid in vitro micropropagation system for the perennial vegetable night lily, Hemerocallis citrina Baroni. Agronomy 14:244

    doi: 10.3390/agronomy14020244

    CrossRef   Google Scholar

    [27]

    Lutz KA, Martin C, Khairzada S, Maliga P. 2015. Steroid-inducible BABY BOOM system for development of fertile Arabidopsis thaliana plants after prolonged tissue culture. Plant Cell Reports 34:1849−56

    doi: 10.1007/s00299-015-1832-7

    CrossRef   Google Scholar

    [28]

    Srinivasan C, Liu Z, Heidmann I, Supena EDJ, Fukuoka H, et al. 2007. Heterologous expression of the BABY BOOM AP2/ERF transcription factor enhances the regeneration capacity of tobacco (Nicotiana tabacum L.). Planta 225:341−51

    doi: 10.1007/s00425-006-0358-1

    CrossRef   Google Scholar

    [29]

    Yang HF, Kou YP, Gao, B, Soliman TMA, Xu KD, et al. 2014. Identification and functional analysis of BABY BOOM genes from Rosa canina. Biologia plantarum 58:427−35

    doi: 10.1007/s10535-014-0420-y

    CrossRef   Google Scholar

    [30]

    Kareem A, Durgaprasad K, Sugimoto K, Du Y, Pulianmackal AJ, et al. 2015. PLETHORA genes control regeneration by a two-step mechanism. Current Biology 25:1017−30

    doi: 10.1016/j.cub.2015.02.022

    CrossRef   Google Scholar

    [31]

    Daimon Y, Takabe K, Tasaka M. 2003. The CUP-SHAPED COTYLEDON genes promote adventitious shoot formation on calli. Plant and Cell Physiology 44:113−21

    doi: 10.1093/pcp/pcg038

    CrossRef   Google Scholar

    [32]

    Gordon SP, Heisler MG, Reddy GV, Ohno C, Das P, et al. 2007. Pattern formation during de novo assembly of the Arabidopsis shoot meristem. Development 134:3539−48

    doi: 10.1242/dev.010298

    CrossRef   Google Scholar

    [33]

    Lian G, Ding Z, Wang Q, Zhang D, Xu J. 2014. Origins and evolution of WUSCHEL-related homeobox protein family in plant kingdom. The Scientific World Journal 2014:534140

    doi: 10.1155/2014/534140

    CrossRef   Google Scholar

    [34]

    Wang Y, He S, Long Y, Zhang X, Zhang X, et al. 2022. Genetic variations in ZmSAUR15 contribute to the formation of immature embryo-derived embryonic calluses in maize. The Plant Journal 109:980−91

    doi: 10.1111/tpj.15609

    CrossRef   Google Scholar

    [35]

    Ikeuchi M, Iwase A, It T, Tanaka H, Favero DS, et al. 2022. Wound-inducible WUSCHEL-RELATED HOMEOBOX 13 is required for callus growth and organ reconnection. Plant Physiology 188:425−41

    doi: 10.1093/plphys/kiab510

    CrossRef   Google Scholar

    [36]

    Kim JY, Yang W, Forner J, Lohmann JU, Noh B, et al. 2018. Epigenetic reprogramming by histone acetyltransferase HAG1/AtGCN5 is required for pluripotency acquisition in Arabidopsis. The EMBO Journal 37:e98726

    doi: 10.15252/embj.201798726

    CrossRef   Google Scholar

    [37]

    Wang J, Tan M, Wang X, Jia L, Wang M, et al. 2023. WUS-RELATED HOMEOBOX 14 boosts de novo plant shoot regeneration. Plant Physiology 192:748−52

    doi: 10.1093/plphys/kiad125

    CrossRef   Google Scholar

    [38]

    Wu X, Chory J, Weigel D. 2007. Combinations of WOX activities regulate tissue proliferation during Arabidopsis embryonic development. Developmental Biology 309:306−16

    doi: 10.1016/j.ydbio.2007.07.019

    CrossRef   Google Scholar

    [39]

    Palovaara J, Hallberg H, Stasolla C, Hakman I. 2010. Comparative expression pattern analysis of WUSCHEL related homeobox2 (WOX2) and WOX8/9 in developing seeds and somatic embryos of the gymnosperm Picea abies. New Phytologist 188:122−35

    doi: 10.1111/j.1469-8137.2010.03336.x

    CrossRef   Google Scholar

    [40]

    Jiang F, Wei G, Sun X, Song X, Wen H, et al. 2018. Cloning and characterization of QtWOX8 gene from Ornithogalum thyrsoide. Molecular Plant Breeding 16:4600−06

    doi: 10.13271/j.mpb.016.004600

    CrossRef   Google Scholar

    [41]

    Shi L, Wang K, Du L, Song Y, Li H, et al. 2021. Genome-wide identification and expression profiling analysis of WOX family protein-encoded genes in Triticeae species. International Journal of Mole cular Sciences 22:9325

    doi: 10.3390/ijms22179325

    CrossRef   Google Scholar

    [42]

    Breuninger H, Rikirsch E, Hermann M, Ueda M, Laux T. 2008. Differential expression of WOX genes mediates apical-Basal axis formation in the Arabidopsis embryo. Developmental Cell 14:867−76

    doi: 10.1016/j.devcel.2008.03.008

    CrossRef   Google Scholar

    [43]

    Emanuelsson O, Nielsen H, Brunak S, Von Heijne G. 2000. Predicting subcellular localization of proteins based on their N-terminal amino acid sequence. Journal of Molecular Biology 300:1005−16

    doi: 10.1006/jmbi.2000.3903

    CrossRef   Google Scholar

    [44]

    Harrison SJ, Mott EK, Parsley K, Aspinall S, Gray JC, et al. 2006. A rapid and robust method of identifying transformed Arabidopsis thaliana seedlings following floral dip transformation. Plant Methods 2:19

    doi: 10.1186/1746-4811-2-19

    CrossRef   Google Scholar

    [45]

    Mortazavi A, Williams BA, McCue K, Schaeffer L, Wold B. 2008. Mapping and quantifying mammalian transcriptomes by RNA-Seq. Nature Methods 5:621−28

    doi: 10.1038/nmeth.1226

    CrossRef   Google Scholar

    [46]

    Kenneth JL, Thomas DS. 2001. Analysis of relative gene expression data using real-time quantitative PCR and the 2−ΔΔCᴛ method. Methods 25:402−08

    doi: 10.1006/meth.2001.1262

    CrossRef   Google Scholar

    [47]

    Wang XD, Nolan KE, Irwanto RR, Sheahan MB, Rose RJ. 2011. Ontogeny of embryogenic callus in Medicago truncatula: the fate of the pluripotent and totipotent stem cells. Annals of Botany 107:599−609

    doi: 10.1093/aob/mcq269

    CrossRef   Google Scholar

    [48]

    Duclercq J, Sangwan-Norreel B, Catterou M, Sangwan RS. 2011. De novo shoot organogenesis: from art to science. Trends in Plant Science 16:597−606

    doi: 10.1016/j.tplants.2011.08.004

    CrossRef   Google Scholar

    [49]

    Dolzblasz A, Nardmann J, Clerici E, Causier B, van der Graaff E, et al. 2016. Stem cell regulation by Arabidopsis WOX genes. Molecular Plant 9:1028−39

    doi: 10.1016/j.molp.2016.04.007

    CrossRef   Google Scholar

    [50]

    Chu H, Liang W, Li J, Hong F, Wu Y, et al. 2013. A CLE-WOX signalling module regulates root meristem maintenance and vascular tissue development in rice. Journal of Experimental Botany 64:5359−69

    doi: 10.1093/jxb/ert301

    CrossRef   Google Scholar

    [51]

    Kadri A, Grenier De March G, Guerineau F, Cosson V, Ratet P. 2021. WUSCHEL overexpression promotes callogenesis and somatic embryogenesis in Medicago truncatula Gaertn. Plants 10:715

    doi: 10.3390/plants10040715

    CrossRef   Google Scholar

    [52]

    Long X, Zhang J, Wang D, Weng Y, Liu S, et al. 2023. Expression dynamics of WOX homeodomain transcription factors during somatic embryogenesis in Liriodendron hybrids. Forestry Research 3:15

    doi: 10.48130/FR-2023-0015

    CrossRef   Google Scholar

    [53]

    Sarkar AK, Luijten M, Miyashima S, Lenhard M, Hashimoto T, et al. 2007. Conserved factors regulate signalling in Arabidopsis thaliana shoot and root stem cell organizers. Nature 446:811−14

    doi: 10.1038/nature05703

    CrossRef   Google Scholar

    [54]

    Mason MG, Mathews DE, Argyros DA, Maxwell BB, Kieber JJ, et al. 2005. Multiple type-B response regulators mediate cytokinin signal transduction in Arabidopsis. The Plant Cell 17:3007−18

    doi: 10.1105/tpc.105.035451

    CrossRef   Google Scholar

    [55]

    Dai X, Liu Z, Qiao M, Li J, Li S, et al. 2017. ARR12 promotes de novo shoot regeneration in Arabidopsis thaliana via activation of WUSCHEL expression. Journal of Integrative Plant Biology 59:747−58

    doi: 10.1111/jipb.12567

    CrossRef   Google Scholar

    [56]

    Park OS, Bae SH, Kim SG, Seo PJ. 2019. JA-pretreated hypocotyl explants potentiate de novo shoot regeneration in Arabidopsis. Plant Signaling & Behavior 14:1618180

    doi: 10.1080/15592324.2019.1618180

    CrossRef   Google Scholar

    [57]

    Wang J, Su Y, Kong X, Ding Z, Zhang XS. 2020. Initiation and maintenance of plant stem cells in root and shoot apical meristems. aBIOTECH 1:194−204

    doi: 10.1007/s42994-020-00020-3

    CrossRef   Google Scholar

    [58]

    Liu J, Hu X, Qin P, Prasad K, Hu Y, et al. 2018. The WOX11-LBD16 pathway promotes pluripotency acquisition in callus cells during de novo shoot regeneration in tissue culture. Plant and Cell Physiology 59:739−48

    doi: 10.1093/pcp/pcy010

    CrossRef   Google Scholar

  • Cite this article

    Zhao X, Chen A, Gao Z, Hou F, Chen Y, et al. 2024. Overexpression of HmWOX8 promotes callus proliferation and shoot regeneration by regulating hormone signaling and shoot development-related genes. Ornamental Plant Research 4: e026 doi: 10.48130/opr-0024-0024
    Zhao X, Chen A, Gao Z, Hou F, Chen Y, et al. 2024. Overexpression of HmWOX8 promotes callus proliferation and shoot regeneration by regulating hormone signaling and shoot development-related genes. Ornamental Plant Research 4: e026 doi: 10.48130/opr-0024-0024

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Overexpression of HmWOX8 promotes callus proliferation and shoot regeneration by regulating hormone signaling and shoot development-related genes

Ornamental Plant Research  4 Article number: e026  (2024)  |  Cite this article

Abstract: Shoot regeneration capacity is essential for prosperous genetic transformation. Previous studies have shown that WUSCHEL-related homeobox (WOX) transcription factor plays a crucial role in callus growth, shoot regeneration, and root development. However, the mechanisms and functions of shoot regeneration related to WOX8 remain unclear. In the current research, the HmWOX8 gene was isolated from Hemerocallis middendorffii by RACE (Rapid-amplification of cDNA ends) technology. Overexpression of HmWOX8 improved callus proliferation and shoot regeneration ability of Arabidopsis and rice, whereas silencing HmWOX8 in H. middendorffii resulted in the inverse correlation. Transcriptome analysis revealed that HmWOX8 enhances the efficiency of callus proliferation and shoot regeneration through two different ways of regulation, including hormone signaling pathways and shoot development-related genes. (I) HmWOX8 regulates crosstalk among different hormone signaling pathways by activating and inhibiting the expression of different genes in these pathways, thus ensuring signal integration for efficient callus proliferation and shoot regeneration. (II) HmWOX8 can upregulate the expression level of shoot developmental genes, including WOX5/7, BBM, AIL5/7, PLT1, PIN6, CUC3, and SCR14/30, to regulate shoot emergence and outgrowth. In addition, Yeast two-hybrid assays and Bimolecular fluorescence complementation assay suggested that HmWOX8 directly interacts with HmCUC2, thereby promoting shoot regeneration. The present research improves the understanding of molecular mechanisms for HmWOX8-mediated regeneration and provides valuable gene resources for breeding programs to promote plant regeneration.

    • Hemerocallis middendorffii (H. middendorffii) belongs to Hemerocallis of Aphoriaceae, which is an excellent open-field perennial flower with high economic value[1,2]. In recent years, with the promotion of eco-friendly and economical landscaping concepts, perennial flowers have gained widespread recognition in garden landscaping, and there has been an increasing demand for new cultivars of Hemerocallis. Therefore, breeding work for Hemerocallis has broad application prospects and high economic value. Currently, traditional field hybrid breeding is the main mode of obtaining excellent new cultivars of Hemerocallis. However, this method is time-consuming and has a long breeding cycle[3]. The use of transgenic technology can overcome the shortcomings of traditional breeding and has significant implications for improving Hemerocallis cultivars[4]. In monocotyledonous plants, only some undifferentiated tissues such as immature embryos can be induced to form calli[5]. H. middendorffii is a typical monocotyledonous plant, and there existed some problems of callus induction difficulty and low proliferation efficiency in genetic transformation in our previous study. It is an important trend to explore and improve the differentiation and transformation efficiency of H. middendorffii to speed up the breeding of new cultivars.

      Remarkable regeneration capacity is an important mean for plants to survive in complex environments. Plant regeneration can be divided into three mechanisms, tissue repair, de novo organogenesis, and somatic embryogenesis[6]. De novo organogenesis refers to the process of regenerating adventitious roots or shoots from detached or wounded organs[7,8]. Shoot regeneration is an essential process involving massive cell fate transition in callus cells and spatial reorganization of cell identities[9]. Experimental evidence has shown that callus proliferation is required for successful shoot outgrowth. Shoot regeneration is a two-step process: Firstly, callus formation was promoted by culturing on auxin-rich callus-inducing medium (CIM), and then the callus was transferred to cytokinin-rich shoot induction medium (SIM) to produce adventitious shoots (AS)[10].

      It is well known that plant hormones are one of the key factors affecting shoot regeneration, and the balance and interaction between auxin and cytokinin (CK) determine cell fate transitions during this process[11]. Auxin response factors (ARFs), the key mediators of auxin signaling, are mainly involved in plant regeneration[12,13]. IAA-related genes are also involved in AS regeneration, such as AUX1, SAUR, and GH3[1417]. YUC-mediated auxin biosynthesis is required for efficient shoot regeneration[11]. CK is another major hormone affecting de novo shoot organogenesis. A high CK concentration can make cells lose the characteristics of roots, destroy root primordia, and promote the regeneration of plant shoots[18]. Consistently, many genes related to CK biosynthesis and signal pathways are considered to be key drivers of shoot regeneration. The type A-ARR and B-ARR genes play negative and positive regulatory roles in the CK signaling pathway, respectively[19]. Overexpression of type A-ARR (ARR7 and ARR15) results in the suppression of shoot regeneration, while high-order mutants of A-ARRs display enhanced shoot regeneration[20]. Additionally, other phytohormones, including abscisic acid (ABA), ethylene (ETH), brassinolide (BR), jasmonic acid (JA), and salicylic acid (SA), which also regulate the process of shoot regeneration[2123]. Previous studies mainly focused on the effects of different hormones and their concentrations on callus induction, proliferation, and adventitious shoots of Hemerocallis[2426]. However, there is no research to reveal the mechanism of plant hormones and their pathways on the regeneration of H. middendorffii at the molecular level.

      Many studies have explored the molecular mechanisms related to shoot regeneration and identified several regeneration-promoting key genes, thereby significantly boosting the regeneration efficiency and genetic transformation of plants. For instance, AINTEGUMENTA-LIKE 5 (AIL5) overexpression increases adventitious shoot regeneration efficiency through activating the expression of shoot development-related genes[13]. Overexpression of the Baby Boom (BBM) gene can enhance bud regeneration ability and genetic transformation efficiency[22,2729]. It has been documented that lateral organ boundary regulator gene CUP-SHAPED COTYLEDON CUC2 (CUC2) plays a major role in de novo shoot regeneration[30]. Ectopic overexpression of CUC1 or CUC2 can enhance de novo shoot formation and the corresponding double mutant cuc1;cuc2 displays reduced shoot regeneration[31,32].

      The WUSCHEL-related homeobox (WOX) gene family is one kind of unique transcription factor in plants, many studies have showed that the WOX gene family members have crucial functions in plant regeneration[33]. Overexpression of the TaWOX5 gene dramatically increases transformation efficiency with less genotype dependency in wheat (Triticum aestivum L.)[34]. In Arabidopsis (Arabidopsis thaliana) and rice (Oryza sativa L.), WOX11 has been identified as the key gene involved in promotion of adventitious rooting. Arabidopsis WOX13 gene facilitates efficient callus formation and organ reconnection by modifying cell wall properties[35]. AtWOX14 can enhance the regeneration of adventitious shoots by affecting the pluripotency of callus cells[36], and its putative rice ortholog OsWOX13 significantly promote shoot regeneration capacity[37]. To date, the function of WOX8 gene has only been reported in a few plant species, including Arabidopsis, Picea abies, Ornithogalum thyrsoides, and Triticum aestivum[3841]. In Arabidopsis, STIMPY-LIKE (STPL)/WOX8 positively regulates early embryonic growth[38]. In addition, it has been shown that WOX8 and its close homolog WOX9 regulate the development of the basal embryo lineage and also of the apical embryo lineage via noncell autonomous activation of WOX2[42]. PaWOX2 and PaWOX8/9 of Picea abies are expressed at high levels in the early growth stages of zygotic and somatic embryos[39]. TaWOX8 genes could promote immature callus proliferation in Triticum aestivum embryos[41]. Although the roles of the WOX8 gene have been studied, the molecular mechanism of the WOX8 gene and cooperative network with other key genes in regulating callus proliferation and shoot regeneration are largely unknown.

      In the present study, the HmWOX8 gene from H. middendorffii was separated using RACE. Through overexpression in Arabidopsis, rice and gene silencing in H. middendorffii, the positive effects of HmWOX8 on callus proliferation and shoot regeneration were identified. In addition, to explore the key candidate genes associated with callus proliferation and shoot regeneration, transcriptome analyses was performed between HmWOX8-overexpression lines (HmWOX8-OE) and wild-type (WT) in rice. Many genes related to hormone signaling pathways and shoot development were significantly and differentially expressed in HmWOX8-OE, and WT. Yeast two-hybrid assays and Bimolecular fluorescence complementation assay confirmed that HmWOX8 could interact with HmCUC2 to promote AS formation. The present findings clarify the positive roles of the WOX8 gene in callus proliferation and shoot regeneration, and provide an important theoretical foundation for further perfecting the TF transcriptional regulation of plant regeneration.

    • H. middendorffii was field grown at the Horticulture Experimental Station of Northeast Agricultural University (Harbin, China; 126.7° E, 45.7° N). The growth conditions in the greenhouse were as follows: relative humidity of 65%−75%, temperature of 20−28 °C, and 12 h of illumination (700 mmol·m−2·s−1) per day. Arabidopsis ecotype Columbia (Col) and transgenic Arabidopsis plants were grown in a climate incubator (4000 lx) at 24 °C under long-day conditions (16 h-light/8 h-dark). WT rice plants (Oryza sativa L. ssp. japonica cv. Nipponbare) and transgenic rice were cultured in climate chamber under a 16-h light (28 °C) : 8-h dark (26 °C) photoperiod.

    • RNA was extracted using TRIzol reagent (Invitrogen, Carlsbad, CA, USA) from H. middendorffii leaves, and reverse transcription was performed to obtain cDNA. The full-length cDNA of HmWOX8 were cloned by applying rapid amplification of cDNA ends (RACE) technology. The primers for 5'- and 3'-RACE cDNA were designed based on the transcriptome of H. middendorfii. Following the manufacturer's instructions for the SMARTer RACE 5'/3' Kit (Clontech, Shiga, Japan), the full-length cDNAs were obtained (GenBank ON303271.2). The target fragment was ligated to the pMD19-T cloning vector (Takara Bio, Japan) and subjected to blue-white spot screening. The positive bacteria were sent to Kumei Biotech (Jilin, China) for sequencing. For construction of the overexpression plasmids HmWOX8. After XbaI enzyme digestion, the full-length coding sequences (without a stop codon) were inserted into pCAMBIA1300-35S::GFP vector using ClonExpress II One Step Cloning Kit (Vazyme Biotech Co., Ltd., Nanjing, China).

    • Sequence alignments were performed using the clustalw (www.genome.jp/tools-bin/clustalw), and the figure of multiple sequence alignment was plotted by SnapGene software. The conserved domain of HmWOX8 and WOX8 proteins from other plant species were identified using MEME (https://meme-suite.org/meme/tools/meme). A phylogenetic analysis was constructed using the Neighbor-Joining method with 1,000 bootstrap replicates in MEGA 7 software.

    • The empty plasmid without HmWOX8 (35S::GFP) was used as a control. The empty plasmid and pCAMBIA1300-35S::HmWOX8-GFP (pC1300-HmWOX8-GFP) fusion plasmid were transformed into rice protoplasts with nuclear marker, respectively. After 48 h of dark culture, the expression of HmWOX8 in rice protoplasts was observed under a laser copolymerization microscope[43].

    • The pC1300-HmWOX8-GFP fusion plasmid was transformed into Agrobacterium GV3101 competent cells using a freeze-thaw technique. The wild-type and wox8 mutant of Arabidopsis thaliana were used for transformation by the floral dipping method[44]. The resistant seedlings were screened using 50 mg·L−1 kanamycin-labeled and identified by PCR of HmWOX8 specific primers. Total RNA of from T1 leaves was extracted using TRIzol reagent and reverse-transcribed with the PrimeScript RT reagent kit with gDNA Eraser (TaKaRa). qPCR was performed with the SYBR Premix Ex Taq kit (TaKaRa) using the QuantStudio 1 (Thermo Fisher Scientific Inc, USA). The Arabidopsis thaliana Actin1 gene was used as an internal control to normalize the different samples. Three biological and three technical replicates were performed for each sample. The primers used here are listed in Supplemental Table S1. The transgenic plants were transplanted into sterilized soil until harvest of T3 generation seeds.

      The wild A. thaliana and T3-generation HmWOX8 gene overexpressed and recovered A. thaliana stems were used as explants. After cutting them, they were inoculated on Murashige and Skoog (MS) medium containing 0.50 mg·L−1 6-BA + 0.10 mg·L−1 NAA + 30 g·L−1 sucrose to induce callus. Callus images were taken at 20 d after cutting and callus was photographed using a Stereotype microscope (1.25×). A ruler was used to measure the size of the callus and calculate their area. Semi-thin sections were observed using the digital slice scanning system (Thermo Fisher Scientific Inc, USA).

    • PROKII-ALCR-HmWOX8-GUS recombinant plasmid was inserted into WT rice plants. Transgenic rice seedlings were cultivated, after which total RNA was isolated and cDNA was synthesized. Transgenic rice plants were verified by RT-qPCR. GUS staining was used to verify the establishment of ethanol-induced startup subsystem in rice.

      Rice seeds of WT and HmWOX8-OE were sterilized with 75% ethanol for 1 min and 10% sodium hypochlorite for 20 min in sequence, followed by five rinses with sterile distilled water. Then the seeds were placed on MS medium with 0.3 g·L−1 proline + 0.6 g·L−1 hydrolyzed casein + 2 mg·L−1 2,4-D + 30 g·L−1 sucrose + 3 g·L−1 phytagel (pH = 5.9). The growth conditions were 26 °C dark (24 h) with 60%−70% relative humidity. On the second day, rice seeds were sprayed with 2% ethanol. Callus were observed, sampled, and photographed 5, 15, and 25 d after callus induction. Materials at 25 d were photographed with a scanning electron microscope (S3400) for morphological observation. The proliferation-induced callus of WT and HmWOX8-OE were cut into 0.5 cm3 pieces and placed in shoot regeneration medium (N6 + 2 mg·L−1 6-BA + 0.2 mg·L−1 NAA + 30 g·L−1 sucrose + 3 g·L−1 phytagel). After 15 d of culture, the shoot regeneration was observed.

    • Total RNA was extracted using TRIzol reagent from the 15 d of WT and HmWOX8-OE lines rice in shoot culture medium. The quality and purity of the RNA were checked using a NanoDrop ND-8000 spectrophotometer (Thermo Fisher Scientific, Pittsburgh, PA, USA), and RNA sequencing was performed in Meiji (Shanghai, China). A total of six libraries (three biological replicates) were sequenced on the Illumina Novaseq 6000 platform. Raw reads were cleaned and filtered. Gene expression levels were then estimated with FPKM[45]. DESeq was used to detect DEGs between the OE and WT lines with the following criteria: p-value < 0.05 and |log2(FoldChange)| > 1. All identified DEGs were mapped to gene ontology (GO) and Tokyo Encyclopedia of Genes and Genomes (KEGG) databases. The significantly enriched biochemical pathways were obtained using KOBAS with corrected p-value < 0.05.

      For the expression validation, 14 DEGs were randomly selected from RNA-seq data to analyze the relative gene expression levels. Total RNA from the callus was isolated according to the manufacturer's protocol. The qRT-PCR was performed according to a previously described method. All the experiments were repeated three times. The relative expression of genes was calculated using the 2−ΔΔCᴛ method[46]. The primers used for qRT-PCR are shown in Supplemental Table S2.

    • The pTRV2-HmWOX8 vector was constructed using a 205 bp fragment of HmWOX8 with BamH I restriction sites. After BamH I enzyme digestion, the 205 bp fragment of HmWOX8 was inserted into the pTRV2 vector using ClonExpress II One Step Cloning Kit, and then transformed into E. coli DH5α. pTRV2-HmWOX8 was introduced into the Agrobacterium EHA105 competent state using the freeze–thaw method. Single colonies were cultured in the corresponding liquid LB media until the cells reached an OD600 of 1.0−1.3. The Agrobacterium cells were centrifuged at 6,000 rpm for 6 min. The bacterial fluid was collected and suspended in 4.74 g·L−1 of MS, 400 μmol·L−1 of acetosyringone, 10 mmol·L−1 of MgCl2, 10 mmol·L−1 MES, 400 mg·L−1 cysteine, and 5 ml·L−1 Twain 20, (pH = 5.6). The OD600 was adjusted to 1.2. pTRV1 and mixed with pTRV2-HmWOX8 bacterial liquid 1:1, and the two bacterial liquids were fully mixed at room temperature. Callus induced by proliferation were selected (differentiated and etiolated tissues were removed), and cut into small pieces of 0.3 cm3. After 7 d, infection liquid was injected into the callus with a 1 mL sterile syringe with a needle. Once every 7 d, this was repeated twice. The infected callus and wild-type callus were treated in the dark for 24 h, and then grown at 22 °C in a greenhouse with a 16 h/8 h light/dark cycle. AS regeneration ability was quantified by estimating the AS increment coefficient, number of ASs per explant, and AS regenerative efficiency. The results of three experiments were analyzed, with each experiment conducted using 32 explants. Quantification parameters were as follows:

      AS increment coefficient = number of ASs/number of explants; number of ASs per explant = number of ASs/number of explants that regenerated ASs; and AS regenerative efficiency (%) = (number of explants that regenerated ASs/number of explants) × 100% (for the calculation, only shoots longer than 1 cm were considered).

    • The full-length coding region of HmWOX8 was cloned into the pGADT7 vector as prey, and HmCUC2 was inserted into the pGBKT7 vector as bait, respectively. These vectors were transformed into the Y2H gold strain. First, transformants were inoculated on synthetic defined plates SD/-Leu/-Trp (DDO) using the daubing method and incubated at 28 °C for 3−5 d. When colonies on the plates grew to 2−3 mm, the co-transformed colonies were transferred to SD/-Leu/-Trp/-His/-Ade/X-a-gal plates and incubated at 28 °C for 3−5 d to observe their growth. The interaction between two fusion proteins were identified by detecting the blue color generated by yeast cells on plates.

      For the BiFC assay, the full-length coding regions of HmWOX8 without stop codons was inserted into pCAMBIA1300-35S-N-GFPN vector, HmCUC2 without stop codons was inserted into pCAMBIA1300-35S-C-GFPC vector, respectively. For transient expression, Agrobacterium tumefaciens strain GV3101 carrying those constructs was infiltrated with p19 (1:1:1 ratio; OD600 = 0.5) into the abaxial side of leaves from tobacco. After 3 d of agroinfiltration, fluorescent signals were analyzed with a confocal microscope.

    • Data were analyzed using one-way analyses of variance with SPSS v10.0 software (SPSS, Inc., Chicago, IL, USA). The mean values were compared via the least significant difference test at the 0.05 probability level. The GraphPad Prism v8 (Graphpad, USA) was used for plotting.

    • The open reading frame of HmWOX8 consisted of 777 bp and was predicted to encode a protein of 258 amino acids (GenBank: ON303271.2). The theoretical isoelectric point (pI) of HmWOX8 was 4.80. The calculated molecular weight of HmWOX8 was 29.25 kD. Sequence alignment analysis showed HmWOX8 and WOX8 proteins from other plant species contained a homeodomain (HD) conserved domain (Fig. 1a, b). The neighbor-joining method (NJ) was used for multiple sequence alignment analysis of the amino acid sequence of HmWOX8 to build a phylogenetic tree. Results showed that the HmWOX8 was highly homologous with AoWOX8 (Fig. 1c).

      Figure 1. 

      Multiple sequence alignment, phylogenetic tree analysis and subcellular localization. (a) Multiple sequence alignment. (b) Conserved domain of HmWOX8 and five orthologous WOX8 proteins. The orthologous proteins were Cocos nucifera (CnWOX8), Elaeis guineensis (EgWOX8), Phoenix dactylifera (PdWOX8), Telopea speciosissima (TsWOX8), Asparagus officinalis (AoWOX8). (c) The Neighbor-Joining phylogenetic tree analysis of the amino acid sequence alignment of the HmWOX8 protein and WOX8 proteins from other plant species. The N-J phylogenetic tree was constructed using MEGA11. AoWOX8 (Asparagus officinalis, XP_0202427), OsWOX8 (Oryza sativa, NP_001393270.1), LrWOX8 (Lolium rigidum, XP_047055505.1), LpWOX8 (Lolium perenne, XP_051227103.1), RpWOX8 (Rhynchospora pubera, KAJ4775450.1), CnWOX8 (Cocos nucifera, KAG1361082.1), EgWOX8 (Elaeis guineensis, XP_010935303.1), PdWOX8 (Phoenix dactylifera, XP_008781303.1), CmWOX8 (Cinnamomum micranthum f. kanehirae, RWR92943.1), TsWOX8 (Telopea speciosissima, XP_043714174.1), MiWOX8 (Macadamia integrifolia, XP_042513605.1), DlWOX8 (Diospyros lotus, XP_052202296.1), BvWOX8 (Beta vulgaris subsp. Vulgaris, XP_010680605.2), CqWOX8 (Chenopodium quinoa, XP_021714114.1), SoWOX8 (Spinacia oleracea, XP_021835543.1), AtWOX8 (Arabidopsis thaliana, AED95323.1). (d) Subcellular localization of HmWOX8 protein in rice protoplasts. Scale bars = 10 μm.

      To determine the subcellular localization of HmWOX8, the pCAMBIA1300-35S::HmWOX8-GFP (pC1300-HmWOX8-GFP) fusion plasmid were introduced into rice protoplasts with the nuclear marker. Under a laser confocal microscope, the fluorescence of HmWOX8 coincided with the fluorescence of a nuclear marker located in the nucleus (Fig. 1d).

    • To confirm the functions of the HmWOX8 gene in callus proliferation, expression vector pCAMBIA1300-HmWOX8-GFP was constructed and transformed into wild-type Arabidopsis and wox8 mutant. Six independent T3 generation overexpressed lines and five independent T3 generation recovery lines of the HmWOX8 gene were examined for kanamycin resistance, and the result was subsequently confirmed with PCR identification (Supplemental Fig. S1a, b). These lines were screened using the selection method described above until T3 generation overexpressed and recovery seeds were obtained. The expression level of HmWOX8 significantly increased in the transformed lines according to qRT-PCR assays (Fig. 2a, b). While WT explants form well-developed calli from the petiole cut end, HmWOX8 overexpression explants generate compact calli with a significant increase in the projection area. Phenotype analysis revealed that callus areas were bigger in HmWOX8 overexpression lines compared to the wild-type (Fig. 2c, d, i). Relative to WT, the areas of callus were increased by restoring HmWOX8 to wox8 mutant (Fig. 2c, e, i). To further identify the developmental function of HmWOX8 in callus proliferation, whether the reduction of callus areas in WT was due to the reduction in cell number or cell size was next asked. Therefore, the semi-thin sections at the median plane of the calli were performed. All the cells contained in the sections were calculated manually, found that WT calli have 22 large cells (n = 3, big cells > 2,500 μm2) (Fig. 2f), whereas HmWOX8 overexpression calli have 128 large cells (n = 3, big cells > 2,500 μm2) for each section on average (Fig. 2g), implicating that cell proliferation was increased in the HmWOX8 overexpression calli. The wox8 mutant calli have 17 big cells (n = 3, big cells > 2,500 μm2). Although there are only 17 large cells in the wox8 mutant callus (the number of large cells is less than that of WT, the callus area was significantly larger than that of WT (Fig. 2i). This is because there are more intermediate cells in the callus of wox8 mutation than WT (400 μm2 < intermediate cells < 2,500 μm2) (Fig. 2g).

      Figure 2. 

      Functional analysis of HmWOX8 transgenic A. thaliana. (a), (b) Identification of HmWOX8 gene expression in transformed Arabidopsis positive plants and wox8 mutant positive plants. (c)−(e) Callus phenotype of wild-type, HmWOX8-overexpression and recovery T3 transgenic plants. Scale bars = 1 mm. (f)−(h) Semi-thin cross sections of wild-type, HmWOX8-overexpression and recovery T3 transgenic plants, Scale bars = 200 μm. (i) Comparison of Wild-type (WT), HmWOX8 overexpression (OE) and recovery (RE) of callus areas in Arabidopsis. ** indicate significance at p < 0.01. The different lowercase letters indicate significant differences (p < 0.05).

    • The exact role and underlying mechanism of HmWOX8 in plant regeneration are still unclear. To verify whether the overexpression of HmWOX8 (HmWOX8-OE) affects callus proliferation, PROKII-ALCR-HmWOX8-GUS recombinant plasmid was inserted into wild-type rice. The GUS staining method was used to verify the establishment of ethanol-induced startup subsystem in transgenic rice (Supplemental Fig. S2ad). The expression level of the HmWOX8 gene in overexpressing transgenic callus was significantly higher than that in wild-type (WT) callus (Supplemental Fig. S2e). Considering the same genetic background and cultivation environment, all phenotypic differences between HmWOX8-OE and WT callus were due to the overexpression of HmWOX8. The callus-associated phenotypes of HmWOX8-OE transgenic and WT callus were observed. The callus phenotype after continuous culture for 5, 15, 25 d are shown in Fig. 3a, transgenic callus areas were significantly higher than that of WT callus areas. Compared with WT, the callus area of HmWOX8-OE has exhibited an increase by 2.52-fold at early stages of callus formation (5d) (Fig. 3b). After continuous culture for 15 d, overexpressing HmWOX8 callus areas reached 39.62 mm2, which was 1.83-fold that of WT (Fig. 3b). Consistent with 5 and 15 d, the 25th day of callus induction also showed an increase of callus density in overexpressing HmWOX8 lines, compared to that in the WT (1.71-fold) (Fig. 3b). Collectively, these results indicated that the overexpression of HmWOX8 could significantly improve the proliferation efficiency of callus. The HmWOX8-OE and WT rice calli cultured for 25 d were observed by scanning electron microscope. The results showed that the surface and length-cutting section of WT were loose and irregular, while HmWOX8-OE have a compact and regular calli with relatively large volume (Fig. 3c). In addition, cell clusters of HmWOX8-OE calli had formed protrusions (Fig. 3c).

      Figure 3. 

      Effects of HmWOX8 overexpression on callus proliferation and shoot regeneration in rice. (a) The callus phenotype of WT and HmWOX8-OE after continuous culture for 5 d, 15 d, and 25 d. Scale bars = 1 cm. (b) Comparison of callus area between WT and HmWOX8-OE after continuous culture for 5 d, 15 d, and 25 d. (c) The scanning electron microscope of WT and HmWOX8-OE. Scale bars = 1 mm. (d) The shoot phenotype of WT and HmWOX8-OE after 15 d in regeneration medium. Scale bars = 1 cm. ** indicate significance at p < 0.01.

      To test the potential role of HmWOX8 in adventitious shoot regeneration, the HmWOX8-OE and WT rice callus were cut into 0.5 cm3 pieces and placed in the shoot regeneration medium. After 15 d, the callus of WT rice proliferated normally, and there were no regenerated green buds (Fig. 3d). Nevertheless, the callus area of over-expressed HmWOX8 increased, and green buds appeared (Fig. 3d). The visual evaluation indicated that the AS regeneration ability of transgenic lines was significantly improved. Therefore, HmWOX8 was speculated to play a crucial role in AS regeneration.

    • To identify the regulatory genes in HmWOX8-overexpression lines, transcriptome sequencing of HmWOX8-OE and WT were performed. Compared to WT, a total of 2,373 DEGs were identified, including 1,037 upregulated and 1,336 downregulated genes in HmWOX8-OE (Fig. 4a). In the KEGG analysis, these DEGs were enriched in the pathways of 'Plant hormone signal transduction' (map04075; p-value = 3.9e-03), 'Phenylpropanoid biosynthesis' (map00940; p-value = 1.9e-03), and 'Starch and sucrose metabolism' (map00500; p-value = 2.1e-02) in HmWOX8-OE (Fig. 4b). Furthermore, GO enrichment patterns were generally consistent with KEGG analysis. The pathway 'carbohydrate metabolic process' in the biological process category (GO:0005975; p-value = 8.18e-06) was significantly enriched in HmWOX8-OE compared to WT (Supplemental Fig. S3). In the molecular function category, 'glycosyltransferase activity' (GO:0016757; p-value = 2.67e-06) was the most abundant pathway (Supplemental Fig. S3). In addition, the dominant subcategory was 'extracellular region' (GO:0005576; p-value =8.90e-06) in the cellular component (Supplemental Fig. S3).

      Figure 4. 

      Volcano map and KEGG enrichment analysis. (a) Volcano map analysis. (b) KEGG enrichment analysis of differentially expressed genes (DEGs) between HmWOX8-OE and WT.

    • The DEGs related to the plant hormone signal transduction pathway were analyzed based on the KEGG enrichment analysis. A total of 56 enriched DEGs were associated with seven sub-pathways of the plant hormone signal transduction pathway (Fig. 5a). This includes 30 DEGs from the auxin sub-pathway, four from the CK sub-pathway, and nine DEGs from the ABA sub-pathway. It also includes three DEGs from the ETH sub-pathway, two from the BR sub-pathway, two from the JA sub-pathway, and six DEGs from the SA sub-pathway (Fig. 5a). In the auxin sub-pathway, four upregulated ARF genes and six downregulated GH3 genes were detected in HmWOX8-OE vs WT. The expression levels of other auxin signaling-related genes, including AUX1, AUX/IAA, and SAUR, were changed between the HmWOX8-OE lines and WT plants (Fig. 5b). When considering the CK signal transduction pathway, upregulated ARR2 (B-ARR) was detected, whereas three differentially expressed A-ARR (ARR4, ARR5, and ARR9) were downregulated between the HmWOX8-OE lines and WT plants (Fig. 5b). The transcriptome analysis revealed that PYL/PYL and PP2C genes were differentially expressed, with five DEGs downregulated and three upregulated in HmWOX8-OE. One gene was upregulated, which encodes SnRK2 (Fig. 5b). Regarding the ETH signal transduction pathway, CTR1 and EIN2 were downregulated, whereas the ERF1/2 gene was upregulated in HmWOX8-OE lines (Fig. 5b). As for BR signal, BRI1 gene was downregulated, whereas BKI1 gene was upregulated (Fig. 5b). Compared with WT, JAR1 and MYC2 genes were downregulated in HmWOX8-OE of JA sub-pathway (Fig. 5b). Several DEGs involved in the SA sub-pathway were also observed. Three DEGs were annotated as NPR1 genes, one was upregulated and the other two were downregulated in HmWOX8-OE vs WT. One downregulated TGA gene was detected in the HmWOX8-OE lines (Fig. 5b).

      Figure 5. 

      DEGs related to phytohormone signaling transduction pathway and shoot development-related genes. (a) Expression of the phytohormone signal pathway. The color of the background frame indicates the pattern of gene expression: red for upregulation, green for downregulation, and yellow for both upregulation and downregulation. The bars represent standard deviation. (b) Heatmap of the DEGs related to plant hormone signal transduction pathway. (c) Relative expression of shoot development-related genes. Error bars = standard deviation. * p < 0.05, ** p < 0.01 (Student's t-test).

      Based on RNA-seq data, shoot development-related genes were analyzed in the HmWOX8 transgenic plants during AS formation. The expression levels of WOX5, 7, and AIL5, 7 were upregulated in HmWOX8-OE lines. Expression levels of the BBM gene were higher in HmWOX8-OE plants than in WT. Compared with the WT plants, PLT1 PIN6, and CUC3 genes were significantly upregulated in the HmWOX8-OE lines. Moreover, SCR14, and SCR30 genes was upregulated in HmWOX8-OE lines (Fig. 5c).

    • To verify the accuracy and reproducibility of the transcriptome sequencing data, the transcript abundance of 14 selected DEGs was analyzed by qRT-PCR. The expression patterns of these genes are basically consistent with the transcriptome data (Supplemental Fig. S4). The results showed the high reliability of RNA-seq data.

    • To further investigate the function of HmWOX8 in H. middendorffii, agrobacterium cultures harboring pTRV2-empty, and pTRV2-HmWOX8 were used to infiltrate the callus of the same size (Fig. 6a). When the callus was infected twice with the recombinant TRV construct (15 d), a morphological difference was observed between the WT, and VIGS-HmWOX8 plants (Fig. 6b). The callus area of the silent HmWOX8 gene was smaller than that of the empty control and WT callus (Fig. 6b). To verify whether HmWOX8 was silent, the expression level of HmWOX8 in the callus infected with pTRV1 + pTRV2-HmWOX8 was measured using the qRT-PCR. Compared with WT and empty, the content of the HmWOX8 gene was significantly reduced in VIGS-HmWOX8 lines (Fig. 6c). After 15 d in vitro culture on shoot regeneration medium, the empty control and WT callus have begun to sprout, while the callus of silent HmWOX8 gene grows slowly (Fig. 6d). Compared to that in the WT and empty, transgenic callus areas were significantly lower. AS increment coefficient and average number of regenerated AS per callus for WT and empty plants were clearly higher than those for VIGS-HmWOX8 (Fig. 6e). After 30 d, WT and empty have differentiated into green leaves, while the callus of silent HmWOX8 has just sprouted. AS regenerative efficiency in WT and empty were significantly higher than that of VIGS-HmWOX8, and no significant difference in AS regenerative efficiency was noted between WT and empty (Fig. 6g). AS increment coefficient and average number of regenerated AS per callus in VIGS-HmWOX8 callus was lower than that in WT and empty plants (Fig. 6g). These results indicated that HmWOX8 promotes AS formation.

      Figure 6. 

      TRV-mediated VIGS of the HmWOX8 gene in H. middendorffii. (a) Callus morphology of VIGS-HmWOX8 lines cultured on induction medium for 1 d compared with WT (uninfected callus) and empty (infection with pTRV1+pTRV2 agrobacterium). Scale bar = 1 cm. (b) Callus morphology of VIGS-HmWOX8 lines cultured on induction medium for 15 d compared with WT and empty. Scale bar = 1 cm. (c) HmWOX8 expression levels in WT, empty and VIGS-HmWOX8 lines. (d) Morphology of callus and AS regeneration in WT, empty, and VIGS-HmWOX8 in 15 d. Scale bar = 1 cm. (e) Callus areas, AS increment coefficient and the average number of per explant of WT, empty, and VIGS-HmWOX8 lines in 15 d. (f) Morphology of AS regenerated from callus in WT, empty, and VIGS-HmWOX8 for 45 d. Scale bar = 1 cm. (g) AS regenerative efficiency, AS increment coefficient and average number of AS per explant of WT, empty, and VIGS-HmWOX8 lines in 45 d. Error bars = standard deviation. *p <0.05, **p <0.01 (Student's t-test).

    • Through the String database, the interaction between OsWOX8 and OsCUCs was predicted in rice. At the same time, it was found that the CUC3 gene was upregulated 7.85-fold in HmWOX8-OE plants based on rice transcriptome analysis, and its expression pattern is similar to HmWOX8. To test this possibility, the homologous gene CUC2 from H. middendorffii was cloned. Then, a yeast two-hybrid (Y2H) assay was conducted using full-length HmCUC2 as bait, and HmWOX8 as prey. The yeast color test with X-α-gal showed that the combinations of yeast co-transformed with AD and BK plasmids were blue (Fig. 7a), confirming the strength of interactions between the HmWOX8 protein and HmCUC2 protein. To confirm the interaction between HmWOX8 and HmCUC2 in living plant cells, the BiFC assays were conducted using the split GFP system. The results indicated that HmWOX8 and HmCUC2 interacted in the nucleus as evidenced by green fluorescence, while negative controls showed no signals in plant cells (Fig. 7b).

      Figure 7. 

      Direct interaction between HmWOX8 and HmCUC2. (a) Yeast two-hybrid assay. pGBKT7 + pGADT7-T was used as a positive control, and pGBKT7-lam + pGADT7-T was used as a negative control. (b) Bimolecular fluorescence complementation assay. GFP was used as a positive control, GFPN-HmWOX8 + GFPC and GFPN + GFPC-HmCUC2 were used as a negative control. Scale bar = 30 μm.

    • Understanding the fate, dynamics and regulatory mechanisms of cells during callus proliferation and shoot regeneration is essential for plant developmental biology[47]. An efficient shoot regeneration system is the main factor leading to successful genetic transformation[21,48]. Little work has been done on improving the shoot regeneration ability in H. middendorffii. WOX proteins are key regulators implicated in stem cell proliferation and maintenance in different types of meristems, thereby serving as promising developmental regulators for plant regeneration[49]. In this study, a crucial role for HmWOX8 in the regulation of callus proliferation and shoot regeneration was uncovered.

      WOX gene family is highly conserved in plants. According to the phylogenetic analysis, HmWOX8 gene is similar to other WOX8 genes from different species. Multiple sequence alignment and conserved motif analyses further validated that HmWOX8 proteins contained a highly conserved DNA-binding HOX domain, and WOX8 genes were conserved in plants. The subcellular localization of HmWOX8 was determined in rice protoplasts, indicating that the protein is located in the nucleus. Previous studies have shown that the WOX gene family members have crucial functions in many growth and development processes, such as stem cell maintenance, somatic embryogenesis, and organ formation in plants[5053]. Studies on WOX8 have focused on its function in the regulation of early embryonic development[38,41], and little is known of its role in callus proliferation and shoot regeneration. To explore the developmental function of HmWOX8, the HmWOX8 overexpression vector was transformed into Arabidopsis and rice. The results revealed that HmWOX8 promotes callus proliferation in vitro by increasing callus areas. In addition, callus areas and shoot regenerative efficiency of VIGS-HmWOX8 lines were lower than that in WT and empty in H. middendorffii. In summary, HmWOX8 was positively correlated with callus proliferation and shoot regeneration.

      It is well known that shoot regeneration is dependent on cell division and cell response to plant hormones, especially auxin and cytokinin responses. A high ratio of CK to auxin promotes shoot regeneration, whereas a high ratio of auxin to CK promotes root regeneration[11]. Transcriptome analysis uncovered that HmWOX8 overexpression changed the expression of hormone signaling pathway genes (Fig. 5a). ARF is an important regulatory factor in the auxin signaling pathway, which connects auxin with the target gene of downstream response. It is worth noting that the four ARF genes were upregulated in the HmWOX8-OE lines (Fig. 5b), suggesting there may be downstream genes of HmWOX8 that promote AS regeneration. Evidence has shown that MdARF9 is the downstream target gene of MdAIL5 in apple, and positively affects AS regeneration[13]. CK response is critical for de novo stem cell initiation and shoot meristem establishment. Type-B ARRs are transcriptional activators of cytokinin signaling, which maintains the signaling homeostasis by directly regulating type-A ARRs[19]. Correspondingly, type-A ARRs negatively regulate cytokinin signaling[54]. Previous research revealed that overexpression of ARR12 (B-ARRs) increases shoot regeneration, ARR7 and ARR15 (A-ARRs) overexpression results in the suppression of shoot regeneration[20,55]. In the current study, the expression level of B-ARRs was clearly increased, and multiple A-ARRs expression levels were decreased in the HmWOX8-OE lines (Fig. 5b), indicating that HmWOX8 can enhance the regenerative ability of explants by regulating the negative feedback loop in CK signaling pathway. Overall, the increased auxin and CK biosynthesis and sensitivity may play an indispensable influence in promoting AS regeneration of HmWOX8-OE lines. Previous studies found that ethylene exerts both positive and negative impacts on shoot regeneration, depending on the sensitivity of explants to ethylene signaling[21]. The JA-signaling pathway is involved in de novo shoot regeneration[56]. However, the molecular mechanism underlying the role of ABA, BR and SA signaling pathways in shoot regeneration is still largely unclear.

      Shoot and root regeneration depend on the activity of stem cells at the stem cell niche to establish apical meristem primordium, a process which is stimulated and regulated by a number of specific regulators[57]. Studies have shown that the regulators of root meristem formation such as WOX5/7/11, LBD16, as well as SCARECROW (SCR) are involved in the acquisition of competency for shoot regeneration[36,58]. In the current study, the expression levels of WOX5/7 and SCR14/30 in the HmWOX8-OE lines were significantly increased (Fig. 5c), indicating that HmWOX8 could enhance shoot regeneration by upregulating these pluripotent factors. BBM is a member of the APETALA2/ETHYLENE RESPONSE FACTOR (AP2/ERF) family and its expression has been shown to improve plant transformation and regeneration[22]. One upregulated BBM gene was detected in transcriptome data (Fig. 5c). AILs, PLT1, and PLT2 promote shoot regeneration through the activation of CUC genes[30]. The CUC proteins are indispensable for the establishment of shoot promeristem, the CUC3 gene were clearly upregulated in the HmWOX8-OE lines in this study (Fig. 5). The CUC2 activity is required once shoot progenitors are regenerated and it is essential to initiate the regeneration of lateral organs at the periphery of shoot progenitors[31]. The Y2H and BiFC assays showed that HmWOX8 can interact with HmCUC2 protein (Fig. 7).

      Based on these results, it is speculated that HmWOX8 promotes callus proliferation and shoot regeneration through two pathways (Fig. 8). (I) HmWOX8 activates the expression of ARF, B-ARR, SnRK2, ETR1/2, BKI1, and inhibits GH3, A-ARR, CTR1, EIN2, BRI1, JAR1, MYC2, TGA expression to regulate crossing among different hormone signaling pathways, thus ensuring signal integration for efficient callus proliferation and AS regeneration. (II) HmWOX8 can upregulate the expression of some shoot development-related genes, including, WOX5/7, BBM, AIL5/7, PLT1, PIN6, CUC3 and SCR14/30, to regulate AS formation. In addition, studies have found that HmWOX8 can directly interact with HmCUC2. Therefore, it is suspected that HmCUC2 may be a key regulator of AS regeneration in plants, and this needs to be proven in subsequent studies. This study provides a theoretical foundation for clarifying the mechanism of HmWOX8-mediated promotion of callus proliferation and AS regeneration.

      Figure 8. 

      Hypothetical model of HmWOX8 regulating callus proliferation and shoot regeneration.

    • In the current study, the HmWOX8 gene was isolated from H. middendorffii, and its function characterization explored using transient overexpression and VIGS analysis. The results indicated that HmWOX8 positively regulates callus proliferation and shoot regeneration in Arabidopsis and rice. Additionally, silencing the HmWOX8 gene in H. middendorffii exhibits lower callus proliferation efficiency and shoot regeneration ability. The transcriptome analyses in HmWOX8-OE and WT rice were conducted to elucidate the potential mechanisms involved in plant regeneration. The results revealed that HmWOX8 enhances the efficiency of callus proliferation and shoot regeneration through two different regulation paths, including hormone signaling pathways and shoot development-related genes. This study provides key insights into the functional diversification of the WOX gene family during plant regeneration. However, more work would be needed to further excavate the HmWOX8 gene of upstream regulatory factors and downstream target genes in shoot regeneration process.

    • The authors confirm contribution to the paper as follows: study conception and design: Liu Y, Chen Y; data collection: Gao Z, Hou F; analysis and interpretation of results: Chen A; helpful discussion provided: Zhao X; manuscript review and editing: Zhao X. All authors reviewed the results and approved the final version of the manuscript.

    • All data generated or analyzed during this study are included in this published article and its supplementary information files.

      • This research was funded by the National Natural Science Foundation of China (No. 32102420).

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

      • # Authors contributed equally: Xueying Zhao, Along Chen

      • Copyright: © 2024 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/.
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    Zhao X, Chen A, Gao Z, Hou F, Chen Y, et al. 2024. Overexpression of HmWOX8 promotes callus proliferation and shoot regeneration by regulating hormone signaling and shoot development-related genes. Ornamental Plant Research 4: e026 doi: 10.48130/opr-0024-0024
    Zhao X, Chen A, Gao Z, Hou F, Chen Y, et al. 2024. Overexpression of HmWOX8 promotes callus proliferation and shoot regeneration by regulating hormone signaling and shoot development-related genes. Ornamental Plant Research 4: e026 doi: 10.48130/opr-0024-0024

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