Information

30.5A: Meristems - Biology


LEARNING OBJECTIVES

  • Discuss the attributes of meristem tissue and its role in plant development and growth

The adult body of vascular plants is the result of meristematic activity. Plant meristems are centers of mitotic cell division, and are composed of a group of undifferentiated self-renewing stem cells from which most plant structures arise. Meristematic cells are also responsible for keeping the plant growing. The Shoot Apical Meristem (SAM) gives rise to organs like the leaves and flowers, while the Root Apical Meristem (RAM) provides the meristematic cells for the future root growth. The cells of the shoot and root apical meristems divide rapidly and are considered to be indeterminate, which means that they do not possess any defined end fate. In that sense, the meristematic cells are frequently compared to the stem cells in animals, which have an analogous behavior and function.

Meristem tissue and plant development

Meristematic tissues are cells or group of cells that have the ability to divide. These tissues in a plant consist of small, densely packed cells that can keep dividing to form new cells. Meristematic tissue is characterized by small cells, thin cell walls, large cell nuclei, absent or small vacuoles, and no intercellular spaces.

Meristematic tissues are found in many locations, including near the tips of roots and stems (apical meristems), in the buds and nodes of stems, in the cambium between the xylem and phloem in dicotyledonous trees and shrubs, under the epidermis of dicotyledonous trees and shrubs (cork cambium), and in the pericycle of roots, producing branch roots. The two types of meristems are primary meristems and secondary meristems.

Meristem Zones

The apical meristem, also known as the “growing tip,” is an undifferentiated meristematic tissue found in the buds and growing tips of roots in plants. Its main function is to trigger the growth of new cells in young seedlings at the tips of roots and shoots and forming buds. Apical meristems are organized into four zones: (1) the central zone, (2) the peripheral zone, (3) the medullary meristem and (3) the medullary tissue.

The central zone is located at the meristem summit, where a small group of slowly dividing cells can be found. Cells of this zone have a stem cell function and are essential for meristem maintenance. The proliferation and growth rates at the meristem summit usually differ considerably from those at the periphery. Surrounding the central zone is the peripheral zone. The rate of cell division in the peripheral zone is higher than that of the central zone. Peripheral zone cells give rise to cells which contribute to the organs of the plant, including leaves, inflorescence meristems, and floral meristems.

An active apical meristem lays down a growing root or shoot behind itself, pushing itself forward. They are very small compared to the cylinder-shaped lateral meristems, and are composed of several layers, which varies according to plant type. The outermost layer is called the tunica, while the innermost layers are cumulatively called the corpus.

Key Points

  • Mitotic cell division happens in plant meristems, which are composed of a group of self-renewing stem cells from which most plant structures arise.
  • The cells of the shoot and root apical meristems divide rapidly and are “indeterminate”, which means that they are not designed for any specific end goal.
  • The Shoot Apical Meristem (SAM) gives rise to organs like the leaves and flowers, while the Root Apical Meristem (RAM) provides cells for future root growth.
  • Meristematic tissue has a number of defining features, including small cells, thin cell walls, large cell nuclei, absent or small vacuoles, and no intercellular spaces.
  • The apical meristem (the growing tip) functions to trigger the growth of new cells in young seedlings at the tips of roots and shoots and forming buds.
  • The apical meristem is organized into four meristematic zones: (1) central zone, (2) peripheral zone, (3) medullary meristem and (3) medullary tissue.

Key Terms

  • meristem: the plant tissue composed of totipotent cells that allows plant growth
  • undifferentiated: describes tissues where the individual cells have not yet developed mature or distinguishing features, or describes embryonic organisms where the organs cannot be identified
  • apical: situated at the growing tip of the plant or its roots, in comparison with intercalary growth situated between zones of permanent tissue

Biology

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Results

ULTRAPETALA1 negatively regulates the size of the WUS-expressing organizing center during reproductive development

Previous analysis of the EMS-induced ult1-1 and ult1-2alleles revealed an increase in inflorescence and floral meristem size,leading to the production of supernumerary flowers and floral organs(Fletcher, 2001)(Fig. 1A-C). To determine the molecular basis of the ult1 meristem enlargement, we performed in situ experiments to analyze the expression pattern of STM as a marker for inflorescence meristem fate. STM is expressed throughout wild-type inflorescence and floral meristems(Long and Barton, 2000 Long et al., 1996), and is absent from the flanking region that corresponds to the incipient flower and floral organ primordia (Fig. 1E). We did not detect any major changes in the overall pattern of STM expression in ult1-1 and ult1-2 inflorescence meristems. STM expression is visible in the central part of the inflorescence and floral meristems and absent from the peripheral region. However, the domain expressing STM is more extensive in ult1mutant meristems than in wild-type meristems(Fig. 1E-G). This result shows that the supernumerary cells present in ult1 mutant meristems(Fletcher, 2001) correspond to meristem cells rather than to lateral organ primordia cells.

Inflorescence and flower phenotypes of ult1 mutants and complementation test. (A-D) Inflorescence meristems of (A) wild-type Ler, (B) ult1-1, (C) ult1-2 and (D) ult1-3plants. (E-G) In situ expression analysis of STM in (E) Ler,(F) ult1-1 and (G) ult1-2 inflorescences. (H) Inflorescence and (inset) flower of ult1-1 plants transformed with the ULT1-214 construct containing a 2.7 kb ULT1 genomic fragment. (I-L) In situ expression analysis of WUS in (I) Ler, (J) ult1-1,(K) ult1-2 and (L) ult1-3 inflorescences. (M-O) Analysis of pSTM::uidA expression in (M) Ler, (N) ult1-1 and(O) ult1-2 inflorescences. (P) Quantification of WUS-expressing cells in Ler (WT) and all three ult1alleles. The mean number of cells was calculated from the most central section of eight individual inflorescences for each genotype. For each section, the maximum number of cells found in one horizontal (width) and one vertical(height) cell file, as well as the total number of cells expressing WUS, was scored. The standard deviation is indicated. Scale bars: 50μm.

Inflorescence and flower phenotypes of ult1 mutants and complementation test. (A-D) Inflorescence meristems of (A) wild-type Ler, (B) ult1-1, (C) ult1-2 and (D) ult1-3plants. (E-G) In situ expression analysis of STM in (E) Ler,(F) ult1-1 and (G) ult1-2 inflorescences. (H) Inflorescence and (inset) flower of ult1-1 plants transformed with the ULT1-214 construct containing a 2.7 kb ULT1 genomic fragment. (I-L) In situ expression analysis of WUS in (I) Ler, (J) ult1-1,(K) ult1-2 and (L) ult1-3 inflorescences. (M-O) Analysis of pSTM::uidA expression in (M) Ler, (N) ult1-1 and(O) ult1-2 inflorescences. (P) Quantification of WUS-expressing cells in Ler (WT) and all three ult1alleles. The mean number of cells was calculated from the most central section of eight individual inflorescences for each genotype. For each section, the maximum number of cells found in one horizontal (width) and one vertical(height) cell file, as well as the total number of cells expressing WUS, was scored. The standard deviation is indicated. Scale bars: 50μm.

To determine whether the ult1 meristem enlargement occurs uniformly across the shoot apex or is confined to one area, we examined the expression patterns of molecular markers for specific meristematic regions in ult1 inflorescences. We have previously reported that the CLV1 expression domain, which corresponds to the most central L3 cells of the SAM, is significantly broader in ult1-1 inflorescence meristems than in wild type (Fletcher,2001). This result suggested a function for ULT1 in restricting cell accumulation in the interior, central region of the meristem. By contrast, the CLV3 expression pattern in the L1 and L2 layers of the central zone appeared to be unchanged in ult1-1 meristems(Fletcher, 2001), either because the CLV3 expression domain is not affected by the mutation,or because its enlargement across such a small group of cells is too slight to be noticed by in situ hybridization.

The WUS gene is expressed in the interior, deeper layers of shoot and floral meristems, overlapping the CLV1 expression domain(Fig. 1I)(Mayer et al., 1998). Mutations in ULT1 result in the lateral expansion of the WUSexpression domain, without altering its layer specificity(Fig. 1J-K). Counting of WUS-expressing cells confirmed that the organizing center is significantly larger in ult1-1 and ult1-2 meristems than in wild-type meristems (Fig. 1P). In wild-type inflorescence meristems, the mean size of the WUS-expressing domain corresponds to 6.12±0.33 cells in width,3±0 cells in height and 14.75±0.97 cells in total. In ult1-1 inflorescence central sections the WUS domain expands to 8.50±1 cells in width, 3.12±0.33 cells in height, and 21.12±2.80 cells in total, while in ult1-2 inflorescence central sections the WUS domain is 7.75±0.66 cells in width,3±0 cells in height and 18±1.66 cells in total. This result shows that the size of the WUS-expressing organizing center is negatively regulated by ULT1 activity. As ult1-1 plants have larger inflorescence and floral meristems than do ult1-2 plants and produce more floral meristems and floral organs(Fletcher, 2001), our results suggest that the size of the WUS-expressing organizing center may directly affect these traits.

Finally, we used a pSTM::uidA(McConnell and Barton, 1998)reporter line as a marker to examine the size of the peripheral zone of the meristem in wild-type and ult1 plants. This reporter construct does not recapitulate the STM expression pattern in the meristem instead,it is expressed at the boundary between the proper inflorescence meristem and the incipient floral primordia (Fig. 1M). The pSTM::uidA expression pattern is unaltered in ult1 inflorescences, indicating that the peripheral region of the mutant meristems is not significantly enlarged(Fig. 1N-O). Altogether, our expression analyses indicate that ULT1 restricts the lateral expansion of CLV1- and WUS- expressing cells in the interior of inflorescence and floral meristems.

Positional cloning of ULT1

To isolate the ULT1 gene, we used CAPS-based mapping(Konieczny and Ausubel, 1993)of recombination breakpoints in 1366 meiotic events among the F2 progeny of ult1-2 (Ler) × wild type (Col-O). We had previously shown that the ult1 mutations mapped between the visible markers ag and ap2 on chromosome 4(Fletcher, 2001). Using the CAPS markers throughout this interval, we established that ult1-2 was flanked by markers PG11 and g8300(www.arabidopsis.org). Thirty-one plants with recombination events between PG11 and g8300 were identified from the mapping population, and used to refine the position of the ult1-2 recombination breakpoints to the ends of BAC F26K10. We sequenced candidate genes annotated on the BAC and identified a single gene(At4g28190) that was mutated in both ult1 alleles.

To confirm the identity of At4g28190 as the ULT1 gene, a genomic clone (ULT1-214) containing the At4g28190 coding region along with 1 kb of upstream and 0.5 kb of downstream sequence was introduced into ult1-1plants, and this clone partially or fully complemented the mutant phenotypes(Fig. 1H). T1 and T2 ult1-1 plants transformed with the ULT1-214 genomic construct produced meristems and flowers similar to those of wild-type plants. In addition, ult1-1 plants carrying the ULT1-214 transgene flowered at the same time as did wild-type plants, while untransformed ult1-1plants flowered 1 week later on average(Fletcher, 2001). These data confirm that At4g28190 encodes the ULT1 gene.

The complete ULT1-coding region was determined by EST and cDNA analysis, RT-PCR and 5′RACE. This region is 714 bp in length, and consists of three exons and two introns(Fig. 2A), encoding a predicted protein of 237 amino acids with a mass of 26.7 kDa. Genomic sequence analysis indicated the presence of a TATA box, a CCAAT box and a GC box, as well as an in frame stop codon upstream of the transcription start site. Ceres cDNA 96705(www.arabidopsis.org)and the sequencing of RT-PCR products support the annotation of this gene. We have identified a missense mutation in the second exon of this gene in the ult1-1 and the ult1-2 alleles(Fig. 2A). The ult1-1mutation is caused by a G to A transition that changes a cysteine residue to a threonine residue at position 173 relative to the translational initiation site (Fig. 2B). The ult1-2 mutation is due to a C to T transition that replaces a serine residue with a phenylalanine residue at position 83.

ULTRAPETALA1 cloning and sequence analysis. (A) Schematic of the positional cloning of the ULT1 locus and the structure of the ULT1 gene. The region of chromosome 4 containing BACs T29A15 to F19B15 is represented. The CAPS markers designed for mapping ult1-2are shown in black boxes and the frequency of recombinant chromosomes is indicated for each marker. The exon/intron structure of the ULT1 gene is shown along with the positions of the ult1-1 and ult1-2mutations. (B) Alignment of the conceptual translation products of the Arabidopsis ULT1 and ULT2 genomic sequences with conceptually translated consensus EST sequences from four other plant species. The sequences compared are from Arabidopsis thaliana ULT1 (AtULT1,At4g28190), Arabidopsis thaliana ULT2 (AtULT2, At2g20825), Glycine max (GmULT, BM524875.1), Lycopersicon esculentum(LeULT, EST357945), Oryza sativa (OsULT, CA763280.1) and Triticum aestivum (TaULT, BG604592). Identical amino acids are boxed and blocks of similar amino acid residues are shaded. The positions of the mutations in ULT1 and ULT2 are shown above the sequences, the SAND domain is boxed in red and the B box-like motif is boxed in green. Stars indicate the amino acid substitution in the ult1-1 (C173T) and ult1-2 (S83F)alleles. Arrowheads denote the position of the T-DNA insertion in the ult1-3 and the ult2-1 allele. Arginine/lysine rich nuclear localization signal (NLS) candidate polypeptides are underlined in white. (C)Multiple sequence alignment of AtULT1, AtULT2 and animal SAND domains from CeC44F1.2 (Caenorhabditis elegans, Z49067), CeC25G4.4(Caenorhabditis elegans, Z70680), HsSp100b (Homo sapiens,U36501), HsNucP41 (Homo sapiens, Q14976), HsNUDR (Homo sapiens, AF049459), DmDEAF-1 (Drosophila melanogaster,AAC47040), HsGMEB2 (Homo sapiens, NM031803), HsGMEB1 (Homo sapiens, NM006582), AIRE-1 (Homo sapiens, AB006682). The alignment was obtained with the ClustalW 1.82 program and manually refined using the calculated two-dimensional structure. Secondary structure elements are shown above the multiple alignment. Period, semicolon and asterisk mark partial to full residue conservation. Color-coding reflects the conservation of amino acid types. Background colors reveal their physiochemical properties(green: hydrophobic red: positively charged residues blue: negatively charged residues), while foreground colors mark identical (red) and similar(blue) amino acids. The amino acid corresponding to the position of ult1-2 mutation is underlined. (D) Alignment of the AtULT1 and AtULT2 B box-like domains with B-box proteins from animals: CeLIN-41(Caenorhabditis elegans, NP492488), CeNCL-1 (Caenorhabditis elegans, P34611), DmDAPPLED (Drosophila melanogaster, Q9V4M2),HsTIF-1 α (Homo sapiens, NP003843), HsPML (Homo sapiens, P29590). The conserved cysteine/histidine residues are boxed. Below the sequence alignment, the conserved spacing of the B2 B-box consensus(Torok and Etkin, 2000) and the ULT B-box consensus are compared.

ULTRAPETALA1 cloning and sequence analysis. (A) Schematic of the positional cloning of the ULT1 locus and the structure of the ULT1 gene. The region of chromosome 4 containing BACs T29A15 to F19B15 is represented. The CAPS markers designed for mapping ult1-2are shown in black boxes and the frequency of recombinant chromosomes is indicated for each marker. The exon/intron structure of the ULT1 gene is shown along with the positions of the ult1-1 and ult1-2mutations. (B) Alignment of the conceptual translation products of the Arabidopsis ULT1 and ULT2 genomic sequences with conceptually translated consensus EST sequences from four other plant species. The sequences compared are from Arabidopsis thaliana ULT1 (AtULT1,At4g28190), Arabidopsis thaliana ULT2 (AtULT2, At2g20825), Glycine max (GmULT, BM524875.1), Lycopersicon esculentum(LeULT, EST357945), Oryza sativa (OsULT, CA763280.1) and Triticum aestivum (TaULT, BG604592). Identical amino acids are boxed and blocks of similar amino acid residues are shaded. The positions of the mutations in ULT1 and ULT2 are shown above the sequences, the SAND domain is boxed in red and the B box-like motif is boxed in green. Stars indicate the amino acid substitution in the ult1-1 (C173T) and ult1-2 (S83F)alleles. Arrowheads denote the position of the T-DNA insertion in the ult1-3 and the ult2-1 allele. Arginine/lysine rich nuclear localization signal (NLS) candidate polypeptides are underlined in white. (C)Multiple sequence alignment of AtULT1, AtULT2 and animal SAND domains from CeC44F1.2 (Caenorhabditis elegans, Z49067), CeC25G4.4(Caenorhabditis elegans, Z70680), HsSp100b (Homo sapiens,U36501), HsNucP41 (Homo sapiens, Q14976), HsNUDR (Homo sapiens, AF049459), DmDEAF-1 (Drosophila melanogaster,AAC47040), HsGMEB2 (Homo sapiens, NM031803), HsGMEB1 (Homo sapiens, NM006582), AIRE-1 (Homo sapiens, AB006682). The alignment was obtained with the ClustalW 1.82 program and manually refined using the calculated two-dimensional structure. Secondary structure elements are shown above the multiple alignment. Period, semicolon and asterisk mark partial to full residue conservation. Color-coding reflects the conservation of amino acid types. Background colors reveal their physiochemical properties(green: hydrophobic red: positively charged residues blue: negatively charged residues), while foreground colors mark identical (red) and similar(blue) amino acids. The amino acid corresponding to the position of ult1-2 mutation is underlined. (D) Alignment of the AtULT1 and AtULT2 B box-like domains with B-box proteins from animals: CeLIN-41(Caenorhabditis elegans, NP492488), CeNCL-1 (Caenorhabditis elegans, P34611), DmDAPPLED (Drosophila melanogaster, Q9V4M2),HsTIF-1 α (Homo sapiens, NP003843), HsPML (Homo sapiens, P29590). The conserved cysteine/histidine residues are boxed. Below the sequence alignment, the conserved spacing of the B2 B-box consensus(Torok and Etkin, 2000) and the ULT B-box consensus are compared.

Database searches revealed the presence of a sequence on Arabidopsis chromosome 2 that is highly similar to ULT1 at the nucleotide level. This paralogous gene, At2g20825, consists of two exons and a single intron. Because overexpression of this gene can rescue the ult1-1 mutant phenotype (see below), we refer to this locus as ULT2. Conceptual translation of ULT2 gives a putative protein of 226 amino acids (26.1 kDa) with 81% identity and 86% similarity to ULT1 over the full-length of the proteins(Fig. 2B). Twenty-one of the 23 cysteine residues present in ULT1, including C173 which is mutated in the ult1-1 allele, are conserved in the ULT2 protein. The serine residue(S83) that is mutated in the ult1-2 allele is also conserved in ULT2. Notably, ULT1 contains five amino acids at the N terminus (residues 2-5) and six amino acids in the middle of the protein (residues 121-126) that are not present in the putative ULT2 protein.

We have identified sequences corresponding to ULT1- and ULT2-like genes in a number of other plant species, including tomato,maize, cotton, rice, soybean and wheat. So far, only a single ULT-like gene has been identified in these species, compared with two in Arabidopsis. An amino acid alignment of the putative ULT-like proteins for which full-length or nearly full-length genomic and/or EST sequences are available, is shown in Fig. 2B. The overall identity between the proteins ranges from 59% to 72% across the length of the protein. No functions have yet been assigned to any of these ULT-like proteins. The ult1-1 and ult1-2mutations both occur in amino acids that are invariant among all nine of the plant species examined, suggesting that these residues are crucial for protein function.

Sequence analysis of the ULT1 and ULT2 proteins

Two domains can be recognized in the ULT1 and ULT2 protein sequences that have been found in transcription factors. The Prosite program(pit.georgetown.edu)revealed a significant structural homology between the N-terminal region of the ULT proteins (Fig. 2B) and a conserved SAND domain found in animal proteins. The SAND domain is an evolutionarily conserved ∼80-100 amino acid DNA-binding motif that takes its name from the Sp100, AIRE-1, NucP41/75 and DEAF-1/suppressin proteins found in humans and Drosophila melanogaster(Gibson et al., 1998). The ULT1 and ULT2 proteins, as well as the other ULT-like plant sequences, share∼75% identity within the SAND domain(Fig. 2C).

The three-dimensional structures of several SAND domains have been determined by NMR and x-ray crystallography(Bottomley et al., 2001 Surdo et al., 2003). The SAND domain is a compact, strongly twisted α/β fold consisting of five antiparallel β-sheets alternating with four α-helices(Fig. 2C). However, the primary sequence of the SAND domain is poorly conserved between family members. The highest degree of amino acid conservation is found between two otherwise unrelated proteins from C. elegans, CeC25G4.4 and CeC44F1.2, which share 57% identity within the SAND domain. Most of the animal proteins share less than 30% identity within the SAND domain, and the pair-wise comparison score can be as low as 7% identity, as shown for the human AIRE-1 and GMEB1/2 proteins. Thus, the similarity between animal SAND domains instead resides at the secondary and consequent tertiary structure level. Similarly, the major conservation of the ULT SAND domains is at the level of the secondary structure: The PsiPred program (McGuffin et al., 2000) predicts the β1, β2, β3 and β5 strands, as well as the α2 and α4 helices in the ULT proteins(Fig. 2C). The program did not detect the α1 and α3 helices or the β4 sheet, probably because of their extremely small size. Only two conserved cores are highlighted by multiple alignment of the SAND domains, the TPxxFE and the KDWK motifs (Fig. 2C). The TPxxFE motif is perfectly conserved among all the putative ULT-like proteins in plants (Fig. 2B,C). The KDWK core is not conserved in ULT1 and ULT2 nor in the mouse and human AIRE-1 proteins at the primary sequence level, but the secondary structure is conserved. The ult1-2 mutation, which causes a null mutant phenotype(see below), lies within the α2 helix of the SAND domain(Fig. 2B,C).

The ULT1 and ULT2 proteins are highly cysteine rich, with cysteine residues accounting for 9.7% of the total amino acid content of each protein(Fig. 2B). One particular arrangement of cysteine residues near the C terminus of the ULT1 and ULT2 proteins is highly similar to that of a B-box motif found in many eukaryotes(Fig. 2D). In these organisms,the B-box domain has been proposed to function in protein-protein and in protein-RNA interactions (Borden,1998 Torok and Etkin,2000). B-box domains are associated with cysteine-rich zinc-binding motifs in otherwise unrelated proteins, many of them transcription factors, that participate in a wide range of cellular processes(Borden, 1998 Torok and Etkin, 2000). The putative B-box region is more highly conserved between ULT1 and the homologous sequences than the rest of the protein(Fig. 2B).

Subcellular localization of the ULT proteins

In animals, SAND domain-containing proteins are found in the nucleus, in the cytoplasm, or in both compartments(Gross and McGinnis, 1996 Jimenez-Lara et al., 2000 Peterson et al., 2004). Similarly, eukaryotic proteins containing B-box domains have been localized to either the nucleus or the cytosol (Borden,1998 Torok and Etkin,2000). The computer programs Prosite(Hulo et al., 2004 Sigrist et al., 2002), PSORT(Nakai and Kanehisa, 1992),SignalP (Nielsen et al.,1997), and NLSdb (Nair et al.,2003) each predict the ULT1 and ULT2 proteins to be localized to the cytosol, based on the absence of any sorting or signal peptide. However,both ULT proteins are small enough to diffuse passively into the nucleus through the nuclear pores (Raikhel,1992). Subcellular localization experiments using enhanced green fluorescent protein (EGFP) as a marker showed that ULT1-EGFP and ULT2-EGFP fusion constructs transiently transformed into onion epidermal cells are localized in both the nucleus and the cytosolic compartments(Fig. 3A).

Subcellular localization of the ULT proteins. (A) Dark-field exposure of an onion epidermal cell transiently expressing an ULT1-EGFP fusion protein. The GFP signal is detected in both the nucleus (arrowhead) and the cytosol. In the cytosol, the fusion protein appears to be distributed in cytoplasmic streams(arrows). (B) Confocal image of a root from a 35S::ULT1-EGFP T2 transgenic plant. (C,D) Confocal image of a petal from a 35S::ULT1-EGFP T2 transgenic plant. (D) DAPI staining of the nuclei and cell walls in the petal shown in C.(E) An immunoblot of protein extracts from inflorescence meristem tissue,using anti-GFP serum. WT, extract from a wild type Ler plant EGFP,extract from a 35S:EGFP transgenic plant ULT-EGFP, extract from a 35S::ULT1-EGFP T2 transgenic plant. (F,G) Dark-field exposures of onion epidermal cells transiently expressing the ULT1-GUS-EGFP fusion protein. The GFP signal is detected in the cytosol and the perinuclear region (F) or both in the cytosol and throughout the nucleus (G). N, nucleus CS, cytoplasmic streams.

Subcellular localization of the ULT proteins. (A) Dark-field exposure of an onion epidermal cell transiently expressing an ULT1-EGFP fusion protein. The GFP signal is detected in both the nucleus (arrowhead) and the cytosol. In the cytosol, the fusion protein appears to be distributed in cytoplasmic streams(arrows). (B) Confocal image of a root from a 35S::ULT1-EGFP T2 transgenic plant. (C,D) Confocal image of a petal from a 35S::ULT1-EGFP T2 transgenic plant. (D) DAPI staining of the nuclei and cell walls in the petal shown in C.(E) An immunoblot of protein extracts from inflorescence meristem tissue,using anti-GFP serum. WT, extract from a wild type Ler plant EGFP,extract from a 35S:EGFP transgenic plant ULT-EGFP, extract from a 35S::ULT1-EGFP T2 transgenic plant. (F,G) Dark-field exposures of onion epidermal cells transiently expressing the ULT1-GUS-EGFP fusion protein. The GFP signal is detected in the cytosol and the perinuclear region (F) or both in the cytosol and throughout the nucleus (G). N, nucleus CS, cytoplasmic streams.

To determine the relevance of this localization pattern in vivo, we generated transgenic ult1-1 plants stably expressing either the ULT1-EGFP or ULT2-EGFP fusion protein under the control of the 35S promoter. Transgenic plants that expressed the ULT1 or ULT2 protein with the EGFP moiety attached to either the N terminus or the C terminus had a wild-type appearance, indicating that the fusion proteins are functional in either orientation and can rescue the ult1-1 mutant phenotypes. Visualizing the ULT1-EGFP or ULT2-EGFP fusion proteins in the roots or petals of the transgenic plants, we observed signal in both the nucleus and the cytosol(Fig. 3B-D). Immunoblotting of extracts from the transgenic plant using an anti-GFP antibody showed that the observed localization pattern is not an artifact due to the cleavage of the fusion protein (Fig. 3E). The same ULT-EGFP fusion proteins in combination with a nuclear localization signal (NLS) or a nuclear export signal (NES) also complement the mutant phenotype when expressed in an ult1-1 background (data not shown).

However, as the ULT-EGFP fusion proteins are still smaller than the nuclear pore exclusion size they may enter or exit the nucleus passively, especially when expressed at high levels under the 35S promoter. To prevent passive entry into or exit from the nucleus, we fused each ULT protein to a combined GUS(β-glucuronidase)-EGFP protein(Grebenok et al., 1997). When bombarded into onion epidermal cells, the constructs gave a GFP and a GUS signal primarily in the cytosol for some cells and equivalently in the cytosol and the nucleus for others (Fig. 3F,G). Thus, the ULT1 and ULT2 proteins have a dual localization in the nucleus and in the cytosol, and may function in both compartments.

ULT1 and ULT2 expression analysis

We used RT-PCR to determine the distribution of ULT1 and ULT2 mRNA transcripts in wild-type tissues. As shown in Fig. 4, ULT1transcripts could be amplified from all tissues tested: roots, 8-day-old seedlings, mature leaves, stems, inflorescences, pollen and siliques. ULT2 expression was specific to the reproductive developmental stage,being detected only in inflorescences, pollen and siliques. For both genes,the highest level of expression was observed in inflorescence tissues.

Expression profiles of the ULT1 and ULT2 genes. RT-PCR analysis was performed on RNA extracts from various wild type Lertissues: roots (R), 8-day-old seedlings (S), mature rosette leaves (L), stems(St), inflorescence apices (In), pollen (P) and siliques (Si). ULT1transcripts were amplified from all tissues examined, whereas ULT2transcripts were detected only during the reproductive phase in inflorescences, pollen and siliques. EF1α was amplified as a control. In addition, control amplification reactions were run with each set of primers using genomic DNA (gDNA) as a template.

Expression profiles of the ULT1 and ULT2 genes. RT-PCR analysis was performed on RNA extracts from various wild type Lertissues: roots (R), 8-day-old seedlings (S), mature rosette leaves (L), stems(St), inflorescence apices (In), pollen (P) and siliques (Si). ULT1transcripts were amplified from all tissues examined, whereas ULT2transcripts were detected only during the reproductive phase in inflorescences, pollen and siliques. EF1α was amplified as a control. In addition, control amplification reactions were run with each set of primers using genomic DNA (gDNA) as a template.

We then performed in situ hybridization experiments to localize the ULT1 and ULT2 mRNAs more precisely in the tissues where they could be detected by RT-PCR. ULT1 and ULT2 transcripts can be detected throughout the inflorescence meristem, and a weak signal can also be detected in the inflorescence vascular tissues(Fig. 5A,B). Neither ULT1 nor ULT2 transcripts are detectable in stage 1 flower meristems budding from the flanks of the inflorescence meristem, but they reappear much stronger in late stage 2 primordia. As soon as the sepal primordia initiate (stage 3), ULT1 and ULT2 expression is excluded from these organ primordia and becomes restricted to the floral meristem (Fig. 5B). As flower development continues, ULT1 and ULT2 transcripts become further restricted to stamen and carpel primordia(Fig. 5D-G). Expression in carpels was detected only in the adaxial domain, corresponding to the region of ovule formation. Hybridization to cross-sections of mature flowers reveals specific signal throughout the ovules and in the tapetum tissue of the anthers(Fig. 5H-I). Thus, the ULT1 and ULT2 mRNA expression patterns are coincident in inflorescence meristems, floral meristems and developing flowers.

ULT1 and ULT2 mRNA expression patterns in inflorescence and flower tissues. RNA localization by in situ hybridization with ULT1 (A,D,H) and ULT2 (B,F,I) antisense probes hybridized to wild-type Ler tissues. (A-C) Longitudinal sections through the inflorescence meristem (ifm) and adjacent floral meristems (fm). ULT1mRNA is localized throughout the inflorescence meristem. No signal was detected in stage 1 floral meristems (white arrowheads). ULT1transcripts reappeared in late stage 2 floral primordia (black arrowheads). As soon as the sepals initiate (stage 3 flower), ULT1 expression becomes restricted to the center of the floral meristem. (C) Control hybridization with an ULT2 sense probe. (D-G) Longitudinal sections through stage 7-8 flowers. ULT1 (D) and ULT2 (F) mRNA was detected in stamen (St) and carpel (Ca) primordia. In both cases the signal appears stronger on the adaxial side of the carpels (arrows). (E,G) Control hybridizations with ULT1 and ULT2 sense probes. (H-J)Transverse sections through mature flowers. ULT1 (H) and ULT2 (I) mRNA was detected in ovules (white arrowheads) and tapetum tissue in the anthers (black arrowheads). (J) Control hybridization with an ULT1 sense probe. Scale bars: 50 μm.

ULT1 and ULT2 mRNA expression patterns in inflorescence and flower tissues. RNA localization by in situ hybridization with ULT1 (A,D,H) and ULT2 (B,F,I) antisense probes hybridized to wild-type Ler tissues. (A-C) Longitudinal sections through the inflorescence meristem (ifm) and adjacent floral meristems (fm). ULT1mRNA is localized throughout the inflorescence meristem. No signal was detected in stage 1 floral meristems (white arrowheads). ULT1transcripts reappeared in late stage 2 floral primordia (black arrowheads). As soon as the sepals initiate (stage 3 flower), ULT1 expression becomes restricted to the center of the floral meristem. (C) Control hybridization with an ULT2 sense probe. (D-G) Longitudinal sections through stage 7-8 flowers. ULT1 (D) and ULT2 (F) mRNA was detected in stamen (St) and carpel (Ca) primordia. In both cases the signal appears stronger on the adaxial side of the carpels (arrows). (E,G) Control hybridizations with ULT1 and ULT2 sense probes. (H-J)Transverse sections through mature flowers. ULT1 (H) and ULT2 (I) mRNA was detected in ovules (white arrowheads) and tapetum tissue in the anthers (black arrowheads). (J) Control hybridization with an ULT1 sense probe. Scale bars: 50 μm.

Next, we determined the expression patterns of both genes in seedlings and embryos. ULT1 is expressed throughout the vegetative SAM and in young leaf primordia (Fig. 6A). A stronger ULT1 signal is detected on the adaxial side of the leaf primordia, as observed for the carpel primordia. The antisense ULT2mRNA probe did not hybridize to seedling tissues(Fig. 6B), confirming that ULT1 but not ULT2 is expressed during the vegetative stage. In mature embryos, both ULT1 and ULT2 transcripts are detected in the SAM, and ULT2 expression is also observed in the RAM(Fig. 6D-G). The expression is detected in very restricted domains corresponding to meristematic cells localized at the apices. Interestingly, in all earlier analyzed stages - from the eight-cell stage to the bending cotyledon stage - ULT1 and ULT2 transcripts are localized throughout the embryo, occasionally displaying a stronger signal between the developing cotyledons and in the vasculature (Fig. 6J-P). This suggests that ULT expression becomes tissue-restricted only at the time when the embryo enters the maturation phase of development. No signal was detected in the suspensor or in the endosperm at any stage(Fig. 6J-P), showing that ULT1 and ULT2 gene expression is embryo specific.

ULT1 and ULT2 mRNA expression patterns in seedlings and embryos. RNA localization by in situ hybridization with ULT1(A,D,J,K,N,P) and ULT2 (B,E,G,L,M,O) antisense probes hybridized to wild-type Ler seedlings (A-C) and embryos (D-Q). (A-C) Longitudinal sections through 7-day-old seedlings. (A) ULT1 transcripts are localized throughout the vegetative SAM and in young leaf primordia. (B) ULT2 transcripts are not detected in seedlings. (C) Control hybridization with an ULT1 sense probe. (D-H) Longitudinal sections through mature embryos. (D) ULT1 expression is restricted to the embryonic SAM (arrowhead). (E) ULT2 transcripts can be detected in the embryonic SAM and RAM (arrowheads). (G) Higher magnification of ULT2 expression in the RAM. (F,H) Control hybridization with ULT1 and ULT2 sense probes, respectively. (I-Q) Longitudinal sections through embryos at different stages of embryogenesis. (I) Early globular stage. (J) Eight-cell-stage embryo. (K) Late triangle stage. (L)Early heart stage. (M) Late heart stage. (N) Early torpedo stage. (O) Torpedo stage. (P,Q) Bending cotyledon stage. (I,Q) Control hybridizations with an ULT1 sense probe. Scale bars: 50 μm in A-F 25 μm in G-Q.

ULT1 and ULT2 mRNA expression patterns in seedlings and embryos. RNA localization by in situ hybridization with ULT1(A,D,J,K,N,P) and ULT2 (B,E,G,L,M,O) antisense probes hybridized to wild-type Ler seedlings (A-C) and embryos (D-Q). (A-C) Longitudinal sections through 7-day-old seedlings. (A) ULT1 transcripts are localized throughout the vegetative SAM and in young leaf primordia. (B) ULT2 transcripts are not detected in seedlings. (C) Control hybridization with an ULT1 sense probe. (D-H) Longitudinal sections through mature embryos. (D) ULT1 expression is restricted to the embryonic SAM (arrowhead). (E) ULT2 transcripts can be detected in the embryonic SAM and RAM (arrowheads). (G) Higher magnification of ULT2 expression in the RAM. (F,H) Control hybridization with ULT1 and ULT2 sense probes, respectively. (I-Q) Longitudinal sections through embryos at different stages of embryogenesis. (I) Early globular stage. (J) Eight-cell-stage embryo. (K) Late triangle stage. (L)Early heart stage. (M) Late heart stage. (N) Early torpedo stage. (O) Torpedo stage. (P,Q) Bending cotyledon stage. (I,Q) Control hybridizations with an ULT1 sense probe. Scale bars: 50 μm in A-F 25 μm in G-Q.

ULT2 overexpression can rescue the ult1-1 mutant phenotypes

Because the ULT1 and ULT2 gene expression patterns overlap in inflorescence and floral meristems, we asked if ULT2 could mimic ULT1 function in these tissues. We transformed a d35S::ULT2 sense construct into ult1-1 plants, in order to increase the level of ULT2expression in this mutant background. We analyzed the capacity of the d35S::ULT2 transgene to rescue the ult1-1 phenotypes, and compared its effects with those of a d35S::ULT1 transgene, by scoring floral organ number and flowering time.

Transgenic d35S::ULT plants display a gradient of phenotypes that correlates with the level of ULT gene overexpression. Those plants expressing the highest levels of ULT1 or ULT2 show dramatic vegetative phenotypes as soon as a few days after germination (C.C.C. and J.C.F.,unpublished). Consequently, we performed the complementation analysis on d35S::ULT2 ult1-1 lines that had a wild-type appearance at the vegetative stage. As expected, RT-PCR experiments showed that these lines display a more moderate increase in ULT2 gene expression than the dramatically affected overexpression lines (data not shown). By analyzing these moderate overexpression lines, we found that the d35S::ULT2 transgene complements the ult1-1 mutant phenotypes to the same extent as the d35S::ULT1 transgene (Fig. 7). Indeed, ult1-1 plants containing either of these constructs display floral organ number and bolting time phenotypes close to those of the wild type. Thus, although the endogenous level of ULT2 is not sufficient to overcome the effect of the ult1-1 mutation, an increase in the amount of wild-type ULT2 protein in the ult1-1 background allows complete rescue of the ult1-1 mutant phenotypes. These data indicate that, when expressed at higher levels, wild-type ULT2 protein can functionally compensate for mutant ULT1-1 protein.

Rescue of the ult1-1 mutant phenotype by an ULT2transgene. (A) Floral organ number in wild-type Ler plants, ult1-1 plants and ult1-1 plants containing the d35S::ULT1 or d35S::ULT2 construct. Graph shows the mean number of organs in the first ten flowers of 10 plants (n=100 flowers), and the standard error is indicated. For the transgenic lines in the ult1-1 mutant background,the mean organ number was calculated from the first ten bolting T1 plants that did not show an overexpression phenotype. (B) Days to bolting after germination for Ler plants, ult1-1 plants and ult1-1 plants containing the d35S::ULT1 or d35S::ULT2 construct. The mean number of days to bolting was calculated from the same populations of plants that were used for the floral organ counts in (A) (n=10 plants), and the standard error is indicated.

Rescue of the ult1-1 mutant phenotype by an ULT2transgene. (A) Floral organ number in wild-type Ler plants, ult1-1 plants and ult1-1 plants containing the d35S::ULT1 or d35S::ULT2 construct. Graph shows the mean number of organs in the first ten flowers of 10 plants (n=100 flowers), and the standard error is indicated. For the transgenic lines in the ult1-1 mutant background,the mean organ number was calculated from the first ten bolting T1 plants that did not show an overexpression phenotype. (B) Days to bolting after germination for Ler plants, ult1-1 plants and ult1-1 plants containing the d35S::ULT1 or d35S::ULT2 construct. The mean number of days to bolting was calculated from the same populations of plants that were used for the floral organ counts in (A) (n=10 plants), and the standard error is indicated.

Identification of ult1 and ult2 T-DNA alleles

In order to examine the full spectrum of biological functions for ULT1 and ULT2, we have obtained an insertion allele of each gene. The ult1-3 allele contains a T-DNA insertion in the first exon of ULT1, 155 bp after the start codon(Fig. 2B). RT-PCR experiments show that ult1-3 plants do not accumulate ULT1 transcripts(Fig. 8A) and place the ult1-3 allele as a null allele. The inflorescence and flower phenotypes of plants carrying the ult1-3 mutation are indistinguishable from those of ult1-2 plants(Fig. 1C,D, Fig. 8B), indicating that the ult1-2 EMS line is a phenotypic null allele probably because of the lack of functional ULT1 protein. WUS molecular marker analysis confirms that the ult1-3 allele phenocopies the ult1-2allele (Fig. 1K,L). In ult1-3 inflorescences, the mean organizing center size is 7.62±0.48 cells in width, 3.12±0.33 cells in height, and 18±1.22 cells in total. These values are not significantly different from ult1-2 (Fig. 1P). Surprisingly, the ult1-1 EMS mutation has a more severe effect on inflorescence meristem size, WUS domain expansion and floral organ number than either the ult1-2 or the ult1-3 null mutations(Fig. 1B-D,J-L,P Fig. 8B). In addition, ult1-2 and ult1-3 mutant plants flower only two days later than the wild type (Fig. 8C),whereas ult1-1 plants are more dramatically affected, flowering up to 2 weeks later than wild-type plants.

ULT1 and ULT2 T-DNA insertion alleles. (A) RT-PCR on wild type Ler, ult1-3 and ult2-1 T-DNA insertion mutant inflorescences. ULT1 transcripts could be amplified from Ler(wild-type) plants but not from ult1-3 plants, while ULT2transcripts were detected in wild-type Col-0 and ult2-1/+heterozygous plants but not in ult2-1 homozygous plants after 40 cycles of PCR. However, after 45 cycles a faint signal corresponding to correctly spliced ULT2 transcript was detected in the ult2-1homozygous lane. EF1α was amplified as a control. Additional control amplification reactions were run with each set of primers using genomic DNA (gDNA) as a template. (B) Floral organ number in Ler, ult1-1,ult1-2 and ult1-3 mutant plants. Graph shows the mean number of organs in the first ten flowers of 10 plants (n=100 flowers), and the standard error is indicated. (C) Mean days to bolting after germination for Ler, ult1-1, ult1-2 and ult1-3 plants (n=10 plants). The standard error is indicated. (D) Floral organ number in ult1-3 homozygous plants, ult1-1/+ heterozygous plants and ult1-1/ult1-3 plants. Graph shows the mean number of organs in the first ten flowers of four or six plants (n=40 flowers for ult1-3 and n=60 flowers for the other genotypes), and the standard error is indicated. (E) Mean days to bolting after germination for ult1-3 homozygous plants, ult1-1/+ heterozygous plants and ult1-1/ult1-3 plants (n=4 plants for ult1-3 and n=6 flowers for the other genotypes). The standard error is indicated.

ULT1 and ULT2 T-DNA insertion alleles. (A) RT-PCR on wild type Ler, ult1-3 and ult2-1 T-DNA insertion mutant inflorescences. ULT1 transcripts could be amplified from Ler(wild-type) plants but not from ult1-3 plants, while ULT2transcripts were detected in wild-type Col-0 and ult2-1/+heterozygous plants but not in ult2-1 homozygous plants after 40 cycles of PCR. However, after 45 cycles a faint signal corresponding to correctly spliced ULT2 transcript was detected in the ult2-1homozygous lane. EF1α was amplified as a control. Additional control amplification reactions were run with each set of primers using genomic DNA (gDNA) as a template. (B) Floral organ number in Ler, ult1-1,ult1-2 and ult1-3 mutant plants. Graph shows the mean number of organs in the first ten flowers of 10 plants (n=100 flowers), and the standard error is indicated. (C) Mean days to bolting after germination for Ler, ult1-1, ult1-2 and ult1-3 plants (n=10 plants). The standard error is indicated. (D) Floral organ number in ult1-3 homozygous plants, ult1-1/+ heterozygous plants and ult1-1/ult1-3 plants. Graph shows the mean number of organs in the first ten flowers of four or six plants (n=40 flowers for ult1-3 and n=60 flowers for the other genotypes), and the standard error is indicated. (E) Mean days to bolting after germination for ult1-3 homozygous plants, ult1-1/+ heterozygous plants and ult1-1/ult1-3 plants (n=4 plants for ult1-3 and n=6 flowers for the other genotypes). The standard error is indicated.

The ult2-1 allele contains a T-DNA insertion in the intron of ULT2, 408 bp after the start codon. Because a weak band corresponding to correctly-spliced ULT2 cDNA could be amplified from inflorescence tissues from some homozygous mutant individuals after 45 cycles of RT-PCR, we cannot conclude that ult2-1 is a null allele(Fig. 8A). ult2-1mutant plants do not display any inflorescence or flower phenotypes, and are indistinguishable from wild-type plants (data not shown). Determining whether the presence of ULT2 protein is required for proper reproductive meristem activity will rely on the identification and analysis of a true null allele for the ULT2 locus.

The ULT1-1 mutant protein has semi-dominant effects

Because the ult1-1 mutant phenotype is more dramatic than that of the ult1-3 null mutant, we analyzed the effect of the ult1-1mutation in the heterozygote state. ult1-1 behaves as a slight semi-dominant allele when heterozygous: of 18 ult1-1/+ plants scored,five had five sepals and/or petals in the first one or two flowers and one had six petals in the first flower. All ult1-1/+ plants are wild type with respect to stamen and carpel number and floral determinacy, indicating that the ult1-1 mutation is recessive with respect to these traits.

To test whether the semi-dominant effect of the ULT1-1 mutant protein is altered in the absence of wild-type ULT1 protein, we compared the floral organ number and flowering time phenotypes of ult1-1/+ plants with those of ult1-1/ult1-3 plants. We found that ult1-1/ult1-3 plants are more severely affected than either ult1-3 homozygous plants or ult1-1/+ heterozygous plants with respect to sepal/petal number and also flowering time (Fig. 8D,E). Thus, eliminating wild-type ULT1 protein enhances the effects of the ult1-1 mutation on flowering time and on floral organ number in the outer two whorls.

Down-regulation of both ULT genes leads to early arrest of the vegetative SAM

Antisense plants carrying a d35S::ULT1 AS construct generated in the Ler wild-type background show a dramatic reduction in the level of both ULT1 and ULT2 transcripts(Fig. 9A). Some plants from the antisense lines fail to germinate (data not shown), while the rest display a range of shoot and floral meristem defects(Fig. 9B-I).

Phenotypes of ULT antisense (AS) lines. (A) RT-PCR on inflorescences of the least affected ULT AS plants (those showing the flower phenotypes illustrated in G-I). The expression of both ULT1 and ULT2 is downregulated. EF1α was amplified as a control. (B) Vegetative phenotypes of the most severely affected ULT AS lines. The plants were grouped into three classes based on their SAM termination phenotypes (class 1 plants terminate the earliest). Shoot apex bright-field images (first row), SEM images (second row), and longitudinal sections (third row) are shown for 7-day-old wild-type Ler seedlings and 14-day-old ULT AS seedlings.(C-E) Reproductive phenotypes of the ULT AS lines. (C) Six-week-old Ler plants. (D,E) Six-week-old ULT AS plants showing reduced flower number and premature arrest of the axillary inflorescence meristems. (F-I)Flower phenotypes of the least affected ULT AS lines. (F) Ler flower.(G) ULT AS flower with extra sepals and petals. (H) Siliques from a Ler flower and from an ULT AS flower with extra carpels. (I) Silique from an ULT AS flower dissected open to reveal the presence of a fifth whorl carpeloid structure developing inside the fourth whorl (arrow). Scale bars: 20μm.

Phenotypes of ULT antisense (AS) lines. (A) RT-PCR on inflorescences of the least affected ULT AS plants (those showing the flower phenotypes illustrated in G-I). The expression of both ULT1 and ULT2 is downregulated. EF1α was amplified as a control. (B) Vegetative phenotypes of the most severely affected ULT AS lines. The plants were grouped into three classes based on their SAM termination phenotypes (class 1 plants terminate the earliest). Shoot apex bright-field images (first row), SEM images (second row), and longitudinal sections (third row) are shown for 7-day-old wild-type Ler seedlings and 14-day-old ULT AS seedlings.(C-E) Reproductive phenotypes of the ULT AS lines. (C) Six-week-old Ler plants. (D,E) Six-week-old ULT AS plants showing reduced flower number and premature arrest of the axillary inflorescence meristems. (F-I)Flower phenotypes of the least affected ULT AS lines. (F) Ler flower.(G) ULT AS flower with extra sepals and petals. (H) Siliques from a Ler flower and from an ULT AS flower with extra carpels. (I) Silique from an ULT AS flower dissected open to reveal the presence of a fifth whorl carpeloid structure developing inside the fourth whorl (arrow). Scale bars: 20μm.

The most strongly affected plants have severely disorganized SAMs that resemble those of fas1 or fas2 plants(Leyser and Furner, 1992 Kaya et al., 2001), with highly aberrant lateral organ initiation(Fig. 9B). Although Ler wild-type seedlings develop four true leaves after 7 days of development, ULT AS seedlings have formed only two cotyledons (class 1), two barely developed filamentous leaves (class 2) or two to three stunted leaves(class 3) after 14 days of development(Fig. 9B, first row). The wild-type SAM is a dome-shaped structure that produces lateral organs in a regular phyllotaxy, but no meristematic structure can be detected between the two cotyledons of class 1 ULT AS plants (arrowhead). Class 2 plants initiate leaf primordia at a greatly reduced rate and their SAMs are very small and flat (asterisk), while class 3 plants produce small leaf primordia(arrowheads) around a reduced SAM composed of few enlarged cells(Fig. 9B, second row). Comparison of sections through 7-day-old Ler and 14-day-old ULT AS seedlings (Fig. 9B, third row)shows that class 1 and class 2 AS seedlings lack more than a few meristematic cells (arrowheads). Sections through class 3 ULT AS shoot apices reveal a small group of enlarged cells that are not organized into layers as in the wild type. After the termination of the primary SAM some ULT AS plants initiate axillary meristems, which generate one or more inflorescences(Fig. 9D) much later than wild-type plants (Fig. 9C). These axillary inflorescence meristems can also arrest precociously, after the production of a couple of flowers (Fig. 9E).

The least severely affected ULT AS plants produce flowers that resemble those of ult1-2 mutants (Fig. 9F-I). These plants form flowers with supernumerous floral organs(Fig. 9G) when compared with wild-type plants (Fig. 9F). Five sepals and five petals are observed in some flowers(Fig. 9G), and others form up to four carpels (Fig. 9H). Flowers from the ULT AS lines also display a partial loss of determinacy, in that supernumerous carpels can develop as fifth whorl structures within the fourth whorl gynoecium (Fig. 9I, arrow).


Stem Growth

Growth in plants occurs as the stems and roots lengthen. Some plants, especially those that are woody, also increase in thickness during their life span. The increase in length of the shoot and the root is referred to as primary growth, and is the result of cell division in the shoot apical meristem. Secondary growth is characterized by an increase in thickness or girth of the plant, and is caused by cell division in the lateral meristem. Figure 4 shows the areas of primary and secondary growth in a plant. Herbaceous plants mostly undergo primary growth, with hardly any secondary growth or increase in thickness. Secondary growth or “wood” is noticeable in woody plants it occurs in some dicots, but occurs very rarely in monocots.

Figure 4. In woody plants, primary growth is followed by secondary growth, which allows the plant stem to increase in thickness or girth. Secondary vascular tissue is added as the plant grows, as well as a cork layer. The bark of a tree extends from the vascular cambium to the epidermis.

Some plant parts, such as stems and roots, continue to grow throughout a plant’s life: a phenomenon called indeterminate growth. Other plant parts, such as leaves and flowers, exhibit determinate growth, which ceases when a plant part reaches a particular size.

Primary Growth

Most primary growth occurs at the apices, or tips, of stems and roots. Primary growth is a result of rapidly dividing cells in the apical meristems at the shoot tip and root tip. Subsequent cell elongation also contributes to primary growth. The growth of shoots and roots during primary growth enables plants to continuously seek water (roots) or sunlight (shoots).

The influence of the apical bud on overall plant growth is known as apical dominance, which diminishes the growth of axillary buds that form along the sides of branches and stems. Most coniferous trees exhibit strong apical dominance, thus producing the typical conical Christmas tree shape. If the apical bud is removed, then the axillary buds will start forming lateral branches. Gardeners make use of this fact when they prune plants by cutting off the tops of branches, thus encouraging the axillary buds to grow out, giving the plant a bushy shape.

Secondary Growth

The increase in stem thickness that results from secondary growth is due to the activity of the lateral meristems, which are lacking in herbaceous plants. Lateral meristems include the vascular cambium and, in woody plants, the cork cambium (see Figure 4).

Figure 5. Lenticels on the bark of this cherry tree enable the woody stem to exchange gases with the surrounding atmosphere. (credit: Roger Griffith)

The vascular cambium is located just outside the primary xylem and to the interior of the primary phloem. The cells of the vascular cambium divide and form secondary xylem (tracheids and vessel elements) to the inside, and secondary phloem (sieve elements and companion cells) to the outside. The thickening of the stem that occurs in secondary growth is due to the formation of secondary phloem and secondary xylem by the vascular cambium, plus the action of cork cambium, which forms the tough outermost layer of the stem. The cells of the secondary xylem contain lignin, which provides hardiness and strength.

In woody plants, cork cambium is the outermost lateral meristem. It produces cork cells (bark) containing a waxy substance known as suberin that can repel water. The bark protects the plant against physical damage and helps reduce water loss. The cork cambium also produces a layer of cells known as phelloderm, which grows inward from the cambium. The cork cambium, cork cells, and phelloderm are collectively termed the periderm. The periderm substitutes for the epidermis in mature plants. In some plants, the periderm has many openings, known as lenticels, which allow the interior cells to exchange gases with the outside atmosphere (Figure 5). This supplies oxygen to the living and metabolically active cells of the cortex, xylem and phloem.

Annual Rings

Figure 6. The rate of wood growth increases in summer and decreases in winter, producing a characteristic ring for each year of growth. Seasonal changes in weather patterns can also affect the growth rate—note how the rings vary in thickness. (credit: Adrian Pingstone)

The activity of the vascular cambium gives rise to annual growth rings. During the spring growing season, cells of the secondary xylem have a large internal diameter and their primary cell walls are not extensively thickened. This is known as early wood, or spring wood. During the fall season, the secondary xylem develops thickened cell walls, forming late wood, or autumn wood, which is denser than early wood. This alternation of early and late wood is due largely to a seasonal decrease in the number of vessel elements and a seasonal increase in the number of tracheids. It results in the formation of an annual ring, which can be seen as a circular ring in the cross section of the stem (Figure 6). An examination of the number of annual rings and their nature (such as their size and cell wall thickness) can reveal the age of the tree and the prevailing climatic conditions during each season.


Apical Meristem Structure

The apical meristem is located just below the root cap in the roots, as seen in the image below. The actual apical meristem is a cluster of densely packed and undifferentiated cells. From these cells will come all of the various cell structure the plant uses. An undifferentiated apical meristem cell will divide again and again, slowly becoming a specialized cell.


Where are cytokinins produced in plants?

Cytokinins are more abundant in developing tissues and organs, such as root tip, shoot apex, cambium, and immature organs, and initially it was thought that cytokinins are synthesized in these limited tissues and organs. However, recent studies provide us with insight into cytokinin-producing and -degrading sites. There are seven IPT genes (IPT1 and IPT3IPT8), two CYP735A genes (CYP735A1 and A2), seven functional LOG genes (LOG1LOG5, LOG7 and LOG8) and seven CKX genes (AtCKX1AtCKX7) in Arabidopsis [3]. These gene families show various expression patterns [27, 29, 31–33]. Notably, IPT3 is expressed in the phloem in both roots and aerial organs, suggesting that the precursor of iP-type cytokinins could be synthesized in a wide range of plant parts [27, 29] (Fig. 4). On the other hand, CYP735A is predominantly expressed in root vasculature [31]. Such spatial distribution of IPT gene and CYP735A expression causes the preferential synthesis of tZ in roots and of iP in shoots. Expression of the LOG gene family covers almost all organs. Thus, the activation step of cytokinin synthesis can occur wherever the LOG genes are expressed [32]. These expression patterns suggest that cytokinins play roles in both long-distance and local signaling.

Spatial expression patterns of IPT3 and CYP735A2 in Arabidopsis. Spatial expression patterns of IPT3 and CYP735A2 are indicated in red and blue, respectively. IPT3 is predominantly expressed in phloem, and CYP735A2 in root vasculature. tZR is the major form of xylem cytokinins, and iPR and cZR are found in phloem [40], suggesting that iP-type and tZ-type cytokinins are directionally translocated between organs. The Arabidopsis picture is modified from Sowerby et al. [53]


SHake

Our laboratory uses genetics to study plant development. We are interested in identifiying the genes that control plant architecture and determining their mechanism of action. Plant architecture results from activities of meristems, thus our lab has a strong focus on the regulation of meristem activity. We characterize mutants that are affected in vegetative and/or inflorescence shoot meristems. The genes are identified through positional cloning and studied using a battery of molecular techniques. Maize is our model organism, but we often study a gene or trait in other grasses such as Brachypodium and Setaria . The laboratory research falls into three categories: 1) understanding the role and regulation of knotted1-like (knox) homeodomain transcription factors, 2) identifying genes that regulate inflorescence architecture in maize and the impacts of drought on inflorescence meristems, 3) determining the cellular signaling mechanisms that pattern a maize leaf.

Educational Background:

The role of knox genes in maize and other species

Knotted1-like homeobox (knox) genes are expressed in meristems and are specifically down-regulated as leaf primordia initiate. Dominant mutants exist in maize that misexpress knox genes in leaves, leading to striking proximal-distal defects. Recessive knox mutants in maize, rice and Arabidopsis fail to elaborate a shoot, thus revealing a function for knox genes in the meristem. We have identified the targets of KNOTTED1 in maize and are carrying out similar experiments in rice and sorghum. Most of the hormone pathways are regulated by KN1. The present goal is to determine which targets are most critical in the meristem and which targets are involved in the leaf patterning defect. We are also seeking to understand the chromatin context of KN1 regulation.


A dominant Knotted1 mutant that results from a transposon insertion in the intron. The normally smooth leaves have bumps (knots) that result from blade cells adopting sheath fate.

Inflorescence architecture in maize and other grasses

Nearly all grasses are characterized by the spikelet, a short branch that contains floral meristems. The diversity in the grasses reflects variation in the organization of spikelets that ultimately originates from activities of meristems. With NSF support, we identified many of the genes that regulate inflorescence architecture in maize. These genes are defined by their mutant phenotypes. Our goal now is to ask how drought affects expression of these master regulatory genes and to identify genes that are responsible for the natural diversity in maize inflorescence architecture.

A scanning electron micrograph of an ear primordia showing rows of spikelet meristems, each of which will develop into a kernel.

Positional signaling in maize leaf development

The maize leaf offers a unique opportunity to study how cells respond to their position and differentiate accordingly. The proximal part of the maize leaf is the sheath, a tissue that wraps around the culm. The distal part of the leaf is the blade, which lays flat to optimize photosynthesis. At the junction of the sheath and blade is the ligule region. In addition to this major proximal-distal axis, the abaxial-adaxial and medial-lateral axes are also defined by distinct cell types and tissue organization. How these axes are established and coordinated is not known. We are using maize mutants to identify the genes that regulate this coordinate system.


The front (abaxial surface) and back (adaxial surface) of a maize leaf.

Tsuda, K. Kurata, N. Ohyanagi, H. and Hake, S. (2014) Genome-Wide Study of KNOX Regulatory Network Reveals Brassinosteroid Catabolic Genes Important for Shoot Meristem Function in Rice. Plant Cell 26:3488-500.

Lewis MW, Bolduc N, Hake K, Htike Y, Hay A, Candela H, Hake S. (2014) Gene regulatory interactions at lateral organ boundaries in maize. Gene regulatory interactions at lateral organ boundaries in maize. Development 141:4590-7.

Thompson BE, Basham C, Hammond R, Ding Q, Kakrana A, Lee TF, Simon SA, Meeley, R, Meyers, BC and Hake, S. (2014) The dicerlike-1 homologue, fuzzy tassel, is required for the regulation of meristem determinacy in the inflorescence and vegetative growth in maize. Plant Cell Dec 2.

Eveland, A.L., Goldshmidt, A., Pautler, M., Morohashi, K., Liseron-Monfils, C., MW, L., Kumari S, Hiraga S, Yang F, Unger-Wallace E, Olson A, Hake S, Vollbrecht E, Grotewold E, Ware D, and Jackson, D. (2014). Regulatory modules controlling maize inflorescence architecture. Genome Res.24:431-43.

Bolduc, N. Tyers, R. Freeling, M. Hake, S. (2014) Unequal redundancies in KNOX genes. Plant Physiology 164: 229-38

O'Connor, D. L., Runions, A., Sluis, A., Bragg, J., Vogel, J. Prusinkiewicz, P. Hake, S. A Division in PIN-Mediated Auxin Patterning During Organ Initiation in Grasses (2014) Plos Computational Biology Jan 30, 2014.

Moon, J. Candela, H. and Hake, S. 2012. The Liguleless narrow mutation affects proximal-distal signaling and leaf growth. Development 140: 405-412.

Bolduc, N., Yilmaz, A., Mejia-Guerra, M.K., Morohashi, K., O'Connor, D., Grotewold, E., and Hake, S. 2012. Unraveling the KNOTTED1 regulatory network in maize meristems Genes & Development 26: 1685-1690.

Chuck, G., Tobias, C., Sun, L., Kraemer, F., Li, C., Dibble, D., Arora, R., Bragg, J. N., Vogel, J. P., Singh, S., Simmons, B., Pauly, M., Hake, S. (2011) Overexpression of the maize Corngrass1 microRNA prevents flowering, improves digestibility and increases starch content of switchgrass PNAS 108:17550-17555.

Johnston, R., Candela, H., Hake, S. Foster, T. (2010) The maize milkweed pod1 mutant reveals a mechanism to modify organ morphology. Genesis 48:416-423.

Chuck, G. Whipple, C., Jackson, D. and Hake, S. (2010) The maize SBP-box transcription factor encoded by tasselsheath4 regulates bract development and establishment of meristem boundaries. Development 137:1243-1250

Ramirez, J. Bolduc, N., Lisch, D., and Hake, S. (2009) Distal expression of knotted1 in maize leaves leads to re-establishment of proximal/distal patterning and leaf dissection. Plant Physiology 151:1878-88.

Thompson, B., Bartling, L., Whipple, C., Hall, D. Schmidt, R, and Hake, S. (2009) bearded-ear encodes a MADS-box transcription factor that controls floral meristem identity and determinacy in maize. Plant Cell 21:2578-90.

Bolduc, N. and Hake, S. (2009) The maize transcription factor KNOTTED1 directly regulates the gibberellin catabolism gene ga2ox1. Plant Cell 21:1647-58.

Chuck G, Meeley R, Hake S. (2008) Floral meristem initiation and meristem cell fate are regulated by the maize AP2 genes ids1 and sid1. Development 135:3013-9.

Candela, H. Johnston, R., Gerhold, A., Foster, T. and Hake, S. (2008) The milkweed pod1 gene encodes a KANADI protein that is required for abaxial-adaxial patterning in maize leaves. Plant Cell 20: 2073-2087.

Magnani, E. and Hake, S. (2008) KNOX lost the OX: The Arabidopsis KNATM gene defines a novel class of KNOX transcriptional regulators missing the homeodomain. Plant Cell 20: 875-887.

Chuck, G. Meeley, R. Irish, E., Sakai, H. Hake, S. (2007) The tasselseed4 microRNA of maize controls meristem cell fate and sex determination by targeting the indeterminate spikelet1/Tasselseed6 gene. Nature Genetics 12:1517-1521 PDF 448k

McSteen, P., Malcomber, S. Skirpan, A., Lunde, C., Wu, X., Kellogg, E. and Hake, S. (2007) barren inflorescence2 encodes a co-ortholog of the PINOID serine/threonine kinase and is required for organogenesis during inflorescence and vegetative development in maize. Plant Phys. 144:100-11.

Chuck, G, Cigan, M., Saeteurn, K. Hake, S. (2007) The heterochronic maize mutant Corngrass1 results from overexpression of a tandem microRNA. Nature Genetics, 39:544-549. PDF 448k

Bortiri, E., Chuck, G., Vollbrecht, E., Rocheford, T., Martienssen, R., Hake, S. ramosa2 encodes a LATERAL ORGAN BOUNDARY domain protein that determines the fate of stem cells in branch meristems of maize. Plant Cell. 2006 Mar18(3):574-85. PDF 448k

Bommert, P., Lunde, C., Nardmann, J., Vollbrecht, E., Running, M., Jackson, D., Hake, S. and Werr, W. (2005) thick tassel dwarf1 encodes a putative maize orthologue of the Arabidopsis CLAVATA1 leucine-rich repeat receptor-like kinase. Development 132:1235-1245.

Hake, S. Smith, H. M. S., Magnani, E., Holtan, H., Mele, G., and Ramirez, J. (2004) The role of knox genes in plant development. Annual Review of Cell and Developmental Biology. 20:125-151

Magnani E, Sjolander K, Hake S. (2004) From endonucleases to transcription factors: evolution of the AP2 DNA binding domain in plants. Plant Cell 16:2265-77.

Member - ARS Hall of Science - 2013
ARS Senior Scientist of the Year - 2011
Fellow - American Association for the Advancement of Science - 2009
Member - National Academy of Sciences - 2009
Stephen Hales Prize - American Society of Plant Biology - 2008
Jeanette Siron Pelton Award - Botanical Society of America - 1996


Additional file 1: Figure S1.

Distinction between zones at the shoot apex, based on curvature. From the curvature map, the boundary (B) could be recognized by its negative Gaussian curvature (light green to blue). Organs (O) are located outside of the boundaries and exhibit highly positive Gaussian curvature (orange to red). Old organs were excluded from the analysis. The meristem was subdivided into central zone (CZ) and peripheral zone (P), assuming that the thickness of peripheral zone ring is roughly similar to the diameter of the central zone. (PNG 167 kb)

Additional file 2: Table S1.

Sample size and statistical analysis. (ODS 5 kb)

Additional file 3: Table S2.

Raw data (relative signal intensities) for individual apices. (ODS 5 kb)

Additional file 4: Figure S2.

Variable auxin pattern in NPA-treated plants expressing DII-Venus. Shoot apical meristems from seedlings grown on NPA-containing medium from germination. At t = 0 h, plants were taken off the drug. Scale bar, 20 μm. (JPG 1268 kb)