Skip to main content
NCBI home page
Wiley Open Access Collection logoLink to Wiley Open Access Collection
. 2022 Sep 13;112(2):451–459. doi: 10.1111/tpj.15958

DEFECTIVELY ORGANIZED TRIBUTARIES 5 is not required for leaf venation patterning in Arabidopsis thaliana

Daniela Vlad 1,✉, Jane A Langdale 1,✉
PMCID: PMC9826136  PMID: 36042697

SUMMARY

The search for genetic regulators of leaf venation patterning started over 30 years ago, primarily focused on mutant screens in the eudicotyledon Arabidopsis thaliana. Developmental perturbations in either cotyledons or true leaves led to the identification of transcription factors required to elaborate the characteristic reticulated vein network. An ortholog of one of these, the C2H2 zinc finger protein DEFECTIVELY ORGANIZED TRIBUTARIES 5 (AtDOT5), was recently identified through transcriptomics as a candidate regulator of parallel venation in maize (Zea mays) leaves. To elucidate how AtDOT5 regulates vein patterning, we generated three independent loss‐of‐function mutations by gene editing in Arabidopsis. Surprisingly, none of them exhibited any obvious phenotypic perturbations. To reconcile our findings with earlier reports, we re‐evaluated the original Atdot5‐1 and Atdot5‐2 alleles. By genome sequencing, we show that reported mutations at the Atdot5‐1 locus are actually polymorphisms between Landsberg erecta and Columbia ecotypes, and that other mutations present in the background most likely cause the pleiotropic mutant phenotype observed. We further show that a T‐DNA insertion in the Atdot5‐2 locus has no impact on leaf venation patterns when segregated from other T‐DNA insertions present in the original line. We thus conclude that AtDOT5 plays no role in leaf venation patterning in Arabidopsis.

Keywords: leaf development, gene editing, venation patterning, Arabidopsis thaliana

Significance Statement

An understanding of gene function is often derived on the basis of loss‐of‐function mutant phenotypes, and thus correct identification of mutated loci is crucial. Through gene editing we reveal previous mis‐identification of a causative mutation that led to inappropriate functional assignation for the DEFECTIVELY ORGANIZED TRIBUTARIES 5 gene in Arabidopsis thaliana.

INTRODUCTION

Vascular plants rely on a complex network of veins to transport water, nutrients and sugar throughout the plant and to serve as structural support. How the vascular network is established and maintained has been the focus of research for decades, spanning disciplines from physiology and development to hydraulics and mathematics (reviewed in Nelson & Dengler, 1997, Sack & Scoffoni, 2013, De Rybel et al., 2016, Perico et al., 2022). Early physiological studies revealed that auxin induces vascular formation after wounding, leading to the hypothesis that canalization of auxin flow through developing veins facilitates self‐organization of the vascular network (Sachs, 1969; Sachs, 1981). Attempts to validate this hypothesis at the molecular level are still ongoing, with new discoveries alternately supporting or refuting the possibility (reviewed in Rolland‐Lagan & Prusinkiewicz, 2005, Bennett et al., 2014, Ravichandran et al., 2020, Perico et al., 2022). Central to these endeavors are screens for mutants with perturbed vascular patterning that enable causative mutations to be identified and thus genetic regulators of the patterning process to be revealed. Mutants with perturbed vascular patterning in roots and/or shoots have been reported in both eudicotyledons and monocotyledons, with most studies to date focused on the eudicot species Arabidopsis thaliana (e.g., Candela et al., 1999; Carland et al., 1999; Hardtke & Berleth, 1998; Petricka et al., 2008; Scarpella & Meijer, 2004; Smillie et al., 2012).

The extent to which patterning mechanisms are conserved in eudicot leaves that elaborate reticulate vein networks and monocot leaves that develop parallel veins is unknown; however, transcriptomic studies identified an ortholog of the Arabidopsis DEFECTIVELY ORGANIZED TRIBUTARIES 5 (AtDOT5) gene as a candidate regulator of leaf venation patterning in maize (Zea mays) (Wang et al., 2013). This observation suggests that aspects of the patterning process may be shared in eudicots and monocots. The Arabidopsis defectively organized tributaries (Atdot) mutants were isolated from an extensive forward genetics screen for altered vein patterning in young leaves of approximately 30 000 individuals (Petricka et al., 2008). The plants were derived from three different mutagenized populations, two generated in the Columbia‐0 (Col‐0) background using either diepoxybutane (Clay & Nelson, 2005) or activation tagging mutagenesis (Weigel et al., 2000) and one obtained by Dissociation (Ds) transposon mutagenesis in the Landsberg erecta (Ler) background (Bancroft et al., 1992). The pleiotropic Atdot5‐1 mutant was isolated from the Ds mutagenized Ler background and was classically mapped to the At1g13290 (AtDOT5) locus (Petricka et al., 2008), which encodes a WIP family C2H2 zinc finger protein with predicted transcription factor activity (Appelhagen et al., 2010). Atdot5‐1 mutants exhibited narrower leaves with misaligned veins, delayed leaf initiation, reduced apical dominance, short roots and enhanced auxin sensitivity. The Atdot5‐1 allele differed from wild type at four amino acid positions, all outside of the C2H2 and WIP domains. A second allele, Atdot5‐2, contained a T‐DNA insertion upstream of the C2H2 domain that conditioned an embryo lethal phenotype (Alonso et al., 2003; Petricka et al., 2008). A second T‐DNA insertion in the line containing the Atdot5‐2 allele was subsequently reported in the 5′ untranslated region (UTR) of At2g26740 (https://abrc.osu.edu/stocks/631530), which encodes one of the Arabidopsis SOLUBLE EPOXIDE HYDROLASES (AtSEH) proteins (Kiyosue et al., 1994; Pineau et al., 2017). Constitutive expression of the AtDOT5 genomic sequence from wild‐type Ler was reported to only partially complement the leaf initiation defects of the Atdot5‐1 mutant (Petricka et al., 2008). To date, the mechanism by which AtDOT5 regulates vein patterning in Arabidopsis has not been elucidated.

Given the potential for conserved function of AtDOT5 orthologs in both eudicot and monocot leaf vein patterning, we sought to determine the mechanism of gene function. To this end we used CRISPR/Cas9 to generate three independent loss‐of‐function alleles in Arabidopsis. Here we report the characterization of those mutants and a re‐analysis of the original mutant alleles.

RESULTS & DISCUSSION

Gene edited loss‐of‐function Atdot5 mutants show no vein patterning or morphological defects

To generate null alleles of AtDOT5, CRISPR/Cas9 was used in conjunction with a single guide RNA (sgRNA) that was designed to target the first exon of the At1g13290 locus. The guide was predicted to bind 100 bp downstream from the start codon and to induce mutations that would disrupt both the WIP and the C2H2 zinc finger domains (Figure 1a). T1 plants were screened for potential mutations by genomic PCR and three different loss‐of‐function alleles were identified in which deletions or insertions led to a premature stop codon (Figure 1a). Two alleles, containing a 5‐bp deletion (C11.1_4) and a 1‐bp insertion (C11.4_A3), arose in the same background (line C11). In order to segregate away any potential interactors from the shared C11 background that might influence the mutant phenotype, the two lines were independently backcrossed as the male parent to wild‐type Col‐0 and then selfed to F2. The third allele, containing a 1‐bp deletion, was isolated as a heterozygous, transgene free T2 line and was fixed as homozygous (F12.12_5) in the T3 generation. For all three mutant alleles, segregating wild‐type siblings were fixed in the same generation for use as controls in phenotyping experiments.

Figure 1.

Figure 1

Open in a new tab

CRISPR‐generated Atdot5 loss‐of‐function mutants resemble wild‐type plants.

(a) Schematic of the AtDOT5 locus targeted for mutagenesis and representation of three edited alleles. Black boxes depict exons, with the overlying red box depicting the WIP domain and the blue boxes depicting the zinc finger domains. The sgRNA complementary sequence is underlined and the PAM site is shown in bold italic. White triangles mark deletions and the black inverted triangle marks an insertion. (b–m) Phenotypic characterization of segregating wild‐type (F12.12_3) (b, d, f, h, j and l) and homozygous Atdot5 loss‐of‐function mutants (F12.12_5) (c, e, g, i, k and m). Plants were imaged prior to bolting (b, c) and after flowering (d, e). Flower morphology appeared normal (f, g) and vascular patterning in both stems (h, i) and leaves (j–m) was similar in wild‐type and loss‐of‐function lines. (l) and (m) are insets of (j) and (k), respectively, as indicated. Scale bars: 1 cm (b, c); 5 cm (d, e); 0.5 mm (f, g); 200 μm (h, i); and 5 mm (j, k).

To characterize the mutant phenotype, homozygous mutants from all three independent lines were grown to maturity alongside wild‐type controls. Unexpectedly, mutant plants were developmentally indistinguishable from segregating wild types at all stages of development. Leaf initiation was unperturbed (Figure 1b,c), there was no loss of apical dominance or delayed flowering (Figure 1d,e), floral organs developed normally and the plants were fertile (Figure 1f,g). Furthermore, cross‐sections revealed no differences in patterning of the stem vasculature (Figure 1h,i) and paradermal views revealed normal leaf venation networks (Figure 1j–m). As such, contrary to previous reports, these results indicated that AtDOT5 function is not necessary for leaf venation patterning or for the regulation of overall plant morphology.

The Atdot5‐2 allele conditions no obvious developmental defects

To try and reconcile our finding that AtDOT5 is dispensable for normal plant development with earlier reports, we re‐evaluated the genotype and phenotype of the original Atdot5‐2 mutant line. The Atdot5‐2 allele was acquired as a homozygous line in the Col‐0 background (SALK_148869c). In this line a T‐DNA is inserted in the first exon of AtDOT5 (At1g13290), upstream of the WIP and C2H2 zinc finger domains (Figure 2a). The genotype was confirmed as homozygous by genomic PCR using primers flanking the T‐DNA insertion point together with a primer in the T‐DNA left border (Figure S1). Importantly, the SALK_148869c line differs from the original SALK_148869 line in that the second insertion at the At2g26740 locus has been segregated away (Figure S1) and the mutant phenotype is not embryo lethal. The phenotype of the SALK_148869c mutant should thus reflect loss of function of AtDOT5. In experiments analogous to those performed with the gene edited Atdot5 loss‐of‐function alleles (Figure 1), the Atdot5‐2 mutant phenotype was compared to that of Col‐0 wild‐type plants. Atdot5‐2 mutants exhibited normal leaf, rosette and flower development, no loss of apical dominance and full fertility (Figure 2b–g). Crucially, no changes were observed in either stem (Figure 2h,i) or leaf vasculature patterning (Figure 2j–m). These data are consistent with our hypothesis that loss of AtDOT5 function does not lead to perturbed vein patterning and suggest that the reported embryo lethal phenotype in the original SALK_148869 line (Petricka et al., 2008) was caused by loss of function of the soluble epoxide hydrolase encoded by the At2g26740 locus. It is notable that this finding would have been revealed in the original study if allelism tests had been carried out between the SALK_148869 line and the Atdot5‐1 mutant.

Figure 2.

Figure 2

Open in a new tab

Homozygous Atdot5‐2 mutants (SALK_148869c) exhibit normal leaf venation patterns. (a) Schematic of the AtDOT5 locus showing the position of the T‐DNA insertion in the SALK_148869c line. Black boxes depict exons, with the overlying red box depicting the WIP domain and the blue boxes depicting the zinc finger domains. (b–m) Phenotypic characterization of Col‐0 (b, d, f, h, j and l) and Atdot5‐2 mutants (c, e, g, i, k and m). Plants were imaged prior to bolting (b, c) and after flowering (d, e). Flower morphology appeared normal (f, g) and vascular patterning in both stems (h, i) and leaves (j–m) was similar in Col‐0 and the T‐DNA line SALK_148869c. (l) and (m) are insets of (j) and (k), respectively, as indicated. Scale bars: 1 cm (b, c); 5 cm (d, e); 0.5 mm (f, g); 200 μm (h, i); and 5 mm (j, k).

Developmental defects exhibited by the Atdot5‐1 mutant cannot be explained by mutations in the AtDOT5 gene

Given that the new gene edited alleles and the Atdot5‐2 allele, all of which are in the Col‐0 background, do not lead to perturbed leaf venation patterns or to any general morphological defects, we considered whether the Atdot5‐1 mutant phenotype was specific to the Ler background and thus to any genetic interactions present therein. To this end, we re‐evaluated the phenotype and genotype of the original Atdot5‐1 mutant line.

Throughout development, Atdot5‐1 mutants exhibited a pleiotropic phenotype characterized by variable seedling morphology, altered phyllotaxy, delayed leaf initiation (Figure 3a,b), delayed flowering and loss of apical dominance (Figure 3c,d) (Petricka et al., 2008). The mutant also showed various defects in flower morphology that impacted on fertility and seed set (Figure 3e–g). Stamens either matured too quickly (Figure 3f) or did not elongate sufficiently (Figure 3g), failing in both cases to efficiently deliver viable pollen to the stigma. Additional vascular bundles were evident in the stem vasculature (Figure 3h,i) and the nearly glabrous and irregularly shaped leaves (Figure 3b) displayed conspicuous vein patterning defects (Figure 3j–m). Specifically, the leaf venation pattern was less complex in Atdot5‐1 leaves (Figure 3k,m) compared to wild type (Figure 3j,l) with fewer tertiary veins evident, most of the quaternary veins absent and higher order veins completely absent. These observations are consistent with the report that suggested AtDOT5 influences multiple aspects of plant development, including leaf venation patterning (Petricka et al., 2008).

Figure 3.

Figure 3

Open in a new tab

Pleiotropic phenotype of the Atdot5‐1 mutant. (a, b) Contrasting rosette phenotypes of Ler (a) and the Atdot5‐1 mutant (b) prior to flowering. (c–g) Flowering phenotype of Ler (c, e) and Atdot5‐1 mutant (d, f, g) plants. Mutant plants flower late and display reduced apical dominance (d). Floral morphology is perturbed with stamens either extending above the carpel (f, white arrow) or failing to fully emerge (g, white arrow). (h–m) Transverse sections through stems (h, i) and paradermal views of cleared juvenile leaves (j–m) from Ler (h, j, l) and Atdot5‐1 mutant (i, k, m) plants. Additional vascular bundles are present in mutant stems (i, black arrows). (j) and (k) are insets of (l) and (m), respectively, as indicated. Scale bars: 1 cm (a, b); 5 cm (c, d); 0.5 mm (e–g); 200 μm (h, i); and 2.5 mm (l, m).

In an attempt to explain the conflicting phenotypic differences between Atdot5‐1 versus the gene edited and Atdot5‐2 loss‐of‐function mutants, we compared the AtDOT5 locus in Ler and Col‐0 Arabidopsis accessions. To this end, the coding sequence (CDS) of AtDOT5 (At1g13750) from the recently published Ler genome (Zapata et al., 2016) was aligned to the Col‐0 AtDOT5 (At1g13290) reference sequence (TAIR). The two sequences differ by 10 single nucleotide polymorphisms (SNPs), six of which result in amino acid changes at positions 36 (T to A), 64 (S to T), 244 (S to G), 267 (V to E), 269 (E to K) and 294 (Y to C). In the original report by Petricka et al. (2008), nine point mutations were reported to alter the sequence of the AtDOT5 gene in the Atdot5‐1 mutant background, with four of them resulting in amino acid changes at positions 46 (T to A), 64 (S to T), 244 (S to G) and 294 (Y to C). The Ler genome sequence reveals that three of the four amino acid changes reported as mutagenesis‐induced in the Atdot5‐1 background correspond to natural variation present between Ler and Col‐0. We could not validate the fourth change (46 – T to A) in the genome of Ler or in the genome of the Atdot5‐1 mutant (see below). As such, mutations in AtDOT5 cannot explain the phenotypic changes observed in the Atdot5‐1 mutant.

Genome sequencing of the Atdot5‐1 mutant reveals multiple polymorphisms and a Ds transposon insertion

To identify potential causative mutations underlying the Atdot5‐1 phenotype, the genome of mutant plants was sequenced at 30‐fold coverage. When the sequence of the AtDOT5 locus in the mutant background was aligned with the published Ler genome, no significant changes were identified in AtDOT5 or in any of the 10 adjacent genes either upstream or downstream (Figure S2). As such, we cannot explain how partial complementation of the Atdot5‐1 mutant phenotype was achieved when sequences from this genomic region were used in transgenic complementation experiments (Petricka et al., 2008).

Given that the Atdot5‐1 line was identified in a collection of transposon‐tagged mutants, we next looked for evidence of transgenes and/or transposons in the Atdot5‐1 mutant genome. A single transgene insertion was identified, corresponding to the transposon‐containing construct used in the mutagenesis process (Bancroft et al., 1992). Transgene reassembly from sequence traces showed that the Ds element was inserted in the CDS of the kanamycin resistance gene (NPTII) in the Atdot5‐1 mutant line, instead of in the streptomycin phosphotransferase gene where it was positioned in the original transformation construct (Figure 4a,b). This observation suggests that the Ds was transactivated by an autonomous Activator element at some point since the original lines were generated. Transposition of Ds is further evidenced by duplication of an 8‐bp sequence ‘GCAGCTGT’ at the insertion point in NPTII. Although not well supported, a single pair of reads, one read corresponding to the Ds element and its paired read mapping to positions 11 454 793–11 454 942 on chromosome 4, suggests that a Ds element might also be inserted upstream of the final exon of AtLer‐4G47010, the Ler ortholog of AT4G20370 in Col‐0 that encodes TWIN SISTER OF FT (TSF). However, because loss‐of‐function mutations in TSF do not condition phenotypes similar to Atdot5‐1 (Yamaguchi et al., 2005) and there was just a single read, we discounted the possibility that the insertion was significant. No other Ds insertion events were detected in the genome.

Figure 4.

Figure 4

Open in a new tab

Transgene insertion on chromosome 1 in the Atdot5‐1 background. (a) Diagram showing the insertion point of the HmR Ds construct in the Atdot5‐1 mutant genome, illustrating the position of the Ds in the NPTII gene. (b) Diagram showing the original transformation construct as depicted by Bancroft et al. (1992) with the Ds inserted in the SPT gene. (c) DRL1 transcript levels in wild‐type Ler and Atdot5‐1 seedlings, as determined by qRT‐PCR. Transcript levels were normalized against Act2 transcript levels and an unpaired t‐test demonstrated that there is no statistically significant difference between the two samples (P = 0.2395). Black circles represent individual datapoints and the red diamonds represent the mean in each case.

The sequence flanking the Ds‐containing transgene in the Atdot5‐1 genome indicates that the transgene is inserted between the DEFORMED ROOTS AND LEAVES (DRL1) locus (At1G13870) and a gene of unknown function (At1G13880) (Figure 4a). DRL1 was first identified by Ds tagging with the reported loss‐of‐function drl1‐1 mutant caused by a transposed Ds (tDs) in the DRL1 CDS (Bancroft et al., 1993). Genomic PCR confirmed that the Ds‐containing transgene is present at the same genomic location in both Atdot5‐1 and drl1‐1 mutants, suggesting that it may have been the source of the tDs that inserted into the DRL1 coding region (Figure S3). Although some aspects of the Atdot5‐1 mutant phenotype resemble those found in drl1 mutants, some are noticeably different. For example, drl1‐1 mutants do not form inflorescences (Bancroft et al., 1993) and drl1‐2 mutants show no venation patterning defects even though leaves are narrower than wild type (Nelissen et al., 2003). As such, we hypothesized that the transgene insertion upstream of the DRL1 locus in Atdot5‐1 has no functional significance. This hypothesis is supported by the fact that any phenotypic consequences of the insertion would have been segregating in the progenitor lines of the drl1‐1 mutant, and none were reported (Bancroft et al., 1993). As a final verification, we assessed whether DRL1 gene expression is perturbed in Atdot5‐1 mutants. DRL1 encodes a putative elongator associated protein that is expressed in all organs during wild‐type development (Nelissen et al., 2003). Notably, quantitative reverse transcriptase‐PCR (qRT‐PCR) using RNA extracted from pooled 7‐day‐old seedlings demonstrated that transcript levels are not significantly different between wild‐type Ler and Atdot5‐1 mutant lines (Figure 4c), and the genome sequence reveals no mutations in the DRL1 CDS. We thus conclude that the pleiotropic Atdot5‐1 mutant phenotype is not caused by a transposon or transgene insertion in the genome, nor by any loss or gain of DRL1 function.

To identify potential causative mutations in the Atdot5‐1 genome, sequence variations that have the potential to disrupt gene function were identified. A total of eight exonic frameshift deletions and 10 exonic frameshift insertions were identified (Table 1). Previous reports of phenotypes associated with loss of function at two of the identified loci – AT3G49360 (Xiong et al., 2009) and At5G54600 (Liu et al., 2013) – suggest that the frameshifts observed at these loci are not responsible for the Atdot5‐1 phenotype. The other sequence variants cannot be eliminated as potential causative mutations without further investigation.

Table 1.

Frameshift mutations identified in the Atdot5‐1 genome

Annotation Type of mutation Chr TAIR 10 Description
AtLer‐1G22960.1 Deletion before ATG Chr1 AT1G12330 Cyclin‐dependent kinase‐like protein
AtLer‐1G22970.1 Deletion exon 3 Chr1 Not annotated
AtLer‐1G22970.1 Deletion exon 4 Chr1
AtLer‐1G38900.1 Deletion 11‐bp deletion, heterozygous Chr1 AT1G27580 F‐box protein (DUF295)
AtLer‐3G50050.1 Deletion exon 7 Chr3 Not annotated
AtLer‐4G42200.1 Deletion exon Chr4 AT4G16230 GDSL‐like lipase/acylhydrolase superfamily protein
AtLer‐4G62980.1 Deletion exon 4, premature stop codon Chr4 AT4g34200 PGDH1; phosphoglycerate dehydrogenase 1; EDA9
AtLer‐5G70960.1 Deletion Chr5 AT5G56200 C2H2 type zinc finger transcription factor family
AtLer‐1G22980.1 Insertion exon 2 Chr1 Not annotated
AtLer‐1G47950.1 Insertion Chr1 Not annotated
AtLer‐1G78630.1 Insertion before ATG Chr1 AT1G65295
AtLer‐2G35290.1 Insertion no stop codon Chr2 AT2G18160 ATBZIP2; basic leucine‐zipper 2; FLORAL TRANSITION AT THE MERISTEM3; FTM3; GBF5; G‐BOX BINDING FACTOR 5; bZIP2
AtLer‐3G18110.1 Insertion before ATG Chr3 AT3G07970 Pectin lyase‐like superfamily protein
AtLer‐3G27460.1 Insertion before ATG Chr3 AT3G16340 Taurine‐transporting ATPase
AtLer‐3G63470.1 Insertion exon 4 Chr3 Not annotated
AtLer‐3G68040.1 Insertion premature stop codon Chr3 AT3G49360 PGL2. Acts redundantly with PGL1 and PGL5 (Xiong et al., 2009)
AtLer‐5G23930.1 Insertion before ATG Chr5 AT5G14200 Isopropylmalate dehydrogenase 1
AtLer‐5G74960.1 Insertion Chr5 AT5G54600 PLASTID RIBOSOMAL PROTEIN L24, RPL24, SUPPRESSOR OF VARIEGATION 8, SVR8 (Liu et al., 2013)
Open in a new tab

CONCLUSION

Gene editing technologies allow for unambiguous assignment of mutant phenotypes to loss‐of‐function alleles. This advance has enabled more robust hypotheses of gene function to be proposed than was previously possible. Genetic screens that utilized highly mutagenic chemicals or non‐specific insertion tags such as T‐DNA or transposons inevitably led to ‘noisy’ genomes that could mask the actual mutation of interest, and RNAi suppression lines similarly led to imprecise interpretations of gene function. The discovery by gene editing that the long‐standing AUXIN BINDING PROTEIN played no role in auxin homeostasis is probably one of the most high‐profile cases of mistaken identity in plant biology (Gao et al., 2015) but others have been reported (Bergelson et al., 2016; Yoshida et al., 2018) and there will undoubtedly be more over the coming years. In this context, we have shown here that the AtDOT5 gene does not regulate venation patterning in the Arabidopsis leaf. This finding raises the question of whether the gene name should be changed. If so, a process needs to be established to enable changes to be made in a systematic way that can also be applied to any future findings of mistaken gene identity.

EXPERIMENTAL PROCEDURES

Plant material

Atdot5‐1 seeds were obtained from David Diaz Ramirez and Nayelli Marsch Martinez (Biotechnology and Biochemistry Department, Center for Research and Advanced Studies [CINVESTAV‐IPN] Irapuato Unit, Mexico). The homozygous T‐DNA line SALK_148869c was obtained from NASC (stock number N648869). Landsberg (Ler) and Columbia‐0 (Col‐0) seeds were originally obtained from Lehle Seeds but have been propagated in the lab for many years. Seeds were sown directly on soil (Levington Seed modular compost), stratified at 4°C for 2 days to break dormancy and then transferred to a controlled environment chamber (CER) with a set temperature of 21°C and a 16‐h photoperiod. Col‐0 plants used to generate CRISPR lines were grown in the greenhouse under the same temperature and photoperiod conditions as the CER.

Construct design and plant transformation

A short sgRNA was designed against the first exon of the At1g13290 gene reference sequence using the CRISPOR online tool (Concordet & Haeussler, 2018). Constructs were generated using parts of a modular cloning system based on Golden Gate technology (Weber et al., 2011). The guide sequence was integrated by PCR into an RNA scaffold derived from EC15768 (Hughes & Langdale, 2022), modified by the addition of 34 bp (5′‐CTAGACCCAGCTTTCTTGTACAAAGTTGGCATTA‐3′) at the 3′ end as found on pICH86966 (Nekrasov et al., 2013). The scaffold was assembled into a Golden Gate level 1 module (position 3, reverse) downstream of the AtU6‐26 promoter. The promoter was amplified by PCR from Col‐0 genomic DNA using the following primers: AtU6‐26pF: 5′‐cactctgtggtctcaGGAGAAGCTTCGTTGAACAACGGA‐3′ and AtU6‐26pR 5′‐cactctgtggtctcaCAATCACTACTTCGACTCTAGCTG‐3′, containing 4‐bp sequences and BpiI restriction sites compatible with the PU Level 0 vector, EC41295.

The Cas9p gene (Ma et al., 2015) obtained by synthesis as an SC module (CDS) was cloned under the control of the AtYAO (At4g05410) promoter (Yan et al., 2015) in a Golden Gate level 1 module corresponding to position 2, forward. The AtYAO promoter was obtained by PCR from genomic DNA with primers pAtYAO‐F (5′‐cactctgtggtctcaGGAGACCCAAATCAACAGCTGCAA‐3′) and pAtYAO‐R (5′‐cactctgtggtctcaCATTTCTTCTCTCTCTCACTCCCTCT‐3′) and cloned as described above into a PU Level 0 vector, EC41295. To terminate transcription the t‐NOS terminator module, EC41421, was used. To allow for the selection of transgenic plants, a module containing the bar gene fused to the Agrobacterium tumefaciens nopaline synthase (NOS) promoter and terminator was cloned adjacent to the T‐DNA left border of the pICSL4723 Golden Gate level 2 backbone, in position 1, reverse. Arabidopsis Col‐0 plants were transformed by floral dipping (Clough & Bent, 1998).

Genotyping

To genotype CRISPR lines, 96 T1 basta resistant seedlings were analyzed using a CAPS marker designed to cut the wild‐type sequence at the predicted editing site (3 bp into the guide sequence from the PAM) (Figure S4). The undigested fragment was sequenced to identify the nature of the mutations. The T2 progeny were screened to select mutant lines free of the transgene. When it was not possible to segregate the mutation from the transgene, the lines were backcrossed onto Col‐0.

Genome sequence

DNA was isolated from pooled 3‐week‐old Atdot5‐1 mutant seedlings and sequenced. Plant whole‐genome sequencing was performed by Novogene Cambridge using a standard Illumina pair‐end sequencing protocol with 30× coverage. 5G data were obtained and mapped to the Ler genome available from the 1001 Genomes Browser (https://1001genomes.org/data/MPIPZ/MPIPZJiao2020/releases/current/strains/Ler/). Standard data analysis provided by Novogene included extracting polymorphisms and predicting their impact on gene function. SNP, insertion/deletion, structural variation and copy number variation analyses were included in the results. Limitations on the bioinformatics pipeline did not allow mapping of the transgene and transposon insertions which were assembled and mapped by manually searching sequence traces.

qRT‐PCR

Total RNA was extracted from pooled 7‐day‐old seedlings using the Qiagen RNeasy Plant Mini Kit and complementary DNA was synthesized using the Maxima cDNA Synthesis kit (Thermo Fisher Scientific, Vilnius, Lithuania). qRT‐PCR analysis was performed on a StepOnePlus Real‐Time PCR System (Applied Biosystems, Life Technologies, Marsiling, Singapore) using SYBR™ Green PCR Master Mix (Ref. 4309155, Applied Biosystems by Thermo Fisher Scientific, Life Technologies, Warrington, UK) with the following cycle conditions: 95°C for 10 min, then 40 cycles of 95°C for 15 sec, 60°C for 10 sec and 72°C for 15 sec.

To amplify the DRL1 gene, primers were designed against the CDS using Primer3Plus (DRL1qRT‐F: 5′‐GTTGGACAGAGCGACACAAG‐3′, DRL1qRT‐F: 5′‐GTGGACCGCTTAGACTCGAT‐3′) and expression levels were normalized to the expression of Actin2 amplified using previously published primers (Liu et al., 2009).

Three biological replicates for Ler, four biological replicates for the Atdot5‐1 mutant and three technical replicates for each sample were run. The Cq values and primer efficiencies were calculated for each sample using the R package qpcR (Ritz & Spiess, 2008) and relative expression levels were calculated using the EasyqpcR package based on previously published algorithms (Hellemans et al., 2007). The box plot was generated using RStudio.

Histology

To analyze leaf venation patterns, leaf 5 from approximately 3‐week‐old (CRISPR mutants, Salk lines, wild‐type Ler and Col‐0) or approximately 4‐week‐old (Atdot5‐1) plants was fixed in 3:1 ethanol:acetic acid and chlorophyll was cleared by successive washes with 70% ethanol followed by overnight incubation in histoclear. Leaf vasculature was imaged using a Leica S9i stereomicroscope against a dark background.

Young stem segments were cut 1 cm from the base of the inflorescence soon after bolting. Where possible the plants were selected to have stems of equivalent heights. In the case of the Atdot5‐1 mutant and the Ler wild‐type control, lateral shoots were used instead of the main inflorescence because the mutant lacked apical dominance. One‐centimeter segments were fixed overnight using 3:1 ethanol:acetic acid and then infiltrated with paraffin wax in a Tissue‐Tek VIP machine (Sakura, www.sakura.eu) following a protocol described previously (Hughes et al., 2019). Ten‐micrometer sections were stained using a 1% Safranin solution in 50% ethanol, counterstained with a 0.04% fast‐green solution in 95% ethanol and mounted in DPX mounting medium. Brightfield images were obtained using Leica LASX software and a DFCT000T camera fitted on a Leica DMRB microscope.

AUTHOR CONTRIBUTIONS

DV and JAL conceived and designed the experiments. DV carried out the experiments and analyzed the data. DV and JAL wrote the manuscript.

CONFLICT OF INTEREST

The authors have no competing interests to declare.

Supporting information

Figure S1. Validation of T‐DNA insertion in AtDOT5 in the SALK_148869c line.

Figure S2. Snapshots of sequencing reads for genes flanking AtDOT5 in the Atdot5‐1 genome.

Figure S3. Confirmation of shared transgene position in Atdot5‐1 and Atdrl1‐1 mutants.

Figure S4. PCR assay to genotype gene edited alleles of AtDOT5.

Click here for additional data file. (2.1MB, docx)

ACKNOWLEDGMENTS

We are grateful to Julie Bull, Roxaana Clayton and Lizzie Jamieson for technical support; John Baker for photography; Steve Kelly for help with bioinformatic analyses; and Tom Hughes, Sophie Johnson, Julia Lambret‐Frotte, Chiara Perico, Sovanna Tan and Maricris Zaidem for discussion throughout the experimental work and during manuscript preparation. This work was funded by the Bill and Melinda Gates Foundation C4 Rice grant awarded to the University of Oxford (2015‐2019, OPP1129902; 2019‐2024, INV‐002970).

Contributor Information

Daniela Vlad, Email: daniela.vlad@biology.ox.ac.uk.

Jane A. Langdale, Email: jane.langdale@biology.ox.ac.uk.

DATA AVAILABILITY STATEMENT

All data generated or analyzed during this study are included in this published article and its supplementary information files, except for the sequence reads of the Atdot5‐1 mutant genome which are available at ArrayExpress https://www.ebi.ac.uk/arrayexpress under accession number E‐MTAB‐12010.

REFERENCES

  1. Alonso, J.M. , Stepanova, A.N. , Leisse, T.J. , Kim, C.J. , Chen, H.M. , Shinn, P. et al. (2003) Genome‐wide Insertional mutagenesis of Arabidopsis thaliana . Science, 301, 653–657. [DOI] [PubMed] [Google Scholar]
  2. Appelhagen, I. , Huep, G. , Lu, G.‐H. , Strompen, G. , Weisshaar, B. & Sagasser, M. (2010) Weird fingers: functional analysis of WIP domain proteins. FEBS Letters, 584, 3116–3122. [DOI] [PubMed] [Google Scholar]
  3. Bancroft, I. , Bhatt, A.M. , Sjodin, C. , Scofield, S. , Jones, J.D.G. & Dean, C. (1992) Development of an efficient two‐element transposon tagging system in Arabidopsis thaliana . Molecular & General Genetics, 233, 449–461. [DOI] [PubMed] [Google Scholar]
  4. Bancroft, I. , Jones, J.D.G. & Dean, C. (1993) Heterologous transposon tagging of the DRL1 locus in Arabidopsis. The Plant Cell, 5, 631–638. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Bennett, T. , Hines, G. & Leyser, O. (2014) Canalization: what the flux? Trends in Genetics, 30, 41–48. [DOI] [PubMed] [Google Scholar]
  6. Bergelson, J. , Buckler, E.S. , Ecker, J.R. , Nordborg, M. & Weigel, D. (2016) A proposal regarding best practices for validating the identity of genetic stocks and the effects of genetic variants. The Plant Cell, 28, 606–609. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Candela, H. , Martinez‐Laborda, A. & Micol, J.L. (1999) Venation pattern formation in Arabidopsis thaliana vegetative leaves. Developmental Biology, 205, 205–216. [DOI] [PubMed] [Google Scholar]
  8. Carland, F. , Berg, B.L. , Fitzgerald, J.N. , Jinamornphongs, S. , Nelson, T. & Keith, B. (1999) Genetic regulation of vascular tissue patterning in Arabidopsis. The Plant Cell, 11, 2123–2138. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Clay, N.K. & Nelson, T. (2005) The recessive epigenetic swellmap mutation affects the expression of two step II splicing factors required for the transcription of the cell proliferation gene STRUWWELPETER and for the timing of cell cycle arrest in the Arabidopsis leaf. The Plant Cell, 17, 1994–2008. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Clough, S.J. & Bent, A.F. (1998) Floral dip: a simplified method for Agrobacterium‐mediated transformation of Arabidopsis thaliana . The Plant Journal, 16, 735–743. [DOI] [PubMed] [Google Scholar]
  11. Concordet, J.‐P. & Haeussler, M. (2018) CRISPOR: intuitive guide selection for CRISPR/Cas9 genome editing experiments and screens. Nucleic Acids Research, 46, W242–W245. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. De Rybel, B. , Mahonen, A.P. , Helariutta, Y. & Weijers, D. (2016) Plant vascular development: from early specification to differentiation. Nature Reviews. Molecular Cell Biology, 17, 30–40. [DOI] [PubMed] [Google Scholar]
  13. Gao, Y. , Zhang, Y. , Zhang, D. , Dai, X. , Estelle, M. & Zhao, Y. (2015) Auxin binding protein 1 (ABP1) is not required for either auxin signaling or Arabidopsis development. Proceedings of the National Academy of Sciences of the United States of America, 112, 2275–2280. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Hardtke, C.S. & Berleth, T. (1998) The Arabidopsis gene MONOPTEROS encodes a transcription factor mediating embryo axis formation and vascular development. EMBO Journal, 17, 1405–1411. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Hellemans, J. , Mortier, G. , De Paepe, A. , Speleman, F. & Vandesompele, J. (2007) qBase relative quantification framework and software for management and automated analysis of real‐time quantitative PCR data. Genome Biology, 8, R19. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Hughes, T.E. & Langdale, J.A. (2022) SCARECROW is deployed in distinct contexts during rice and maize leaf development. Development, 149, e200410. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Hughes, T.E. , Sedelnikova, O.V. , Wu, H. , Becraft, P.W. & Langdale, J.A. (2019) Redundant SCARECROW genes pattern distinct cell layers in roots and leaves of maize. Development, 146, e177543. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Kiyosue, T. , Beetham, J.K. , Pinot, F. , Hammock, B.D. , Yamaguchi‐Shinozaki, K. & Shinozaki, K. (1994) Characterization of an Arabidopsis cDNA for a soluble epoxide hydrolase gene that is inducible by auxin and water stress. The Plant Journal, 6, 259–269. [DOI] [PubMed] [Google Scholar]
  19. Liu, J. , Elmore, J.M. , Fuglsang, A.T. , Palmgren, M.G. , Staskawicz, B.J. & Coaker, G. (2009) RIN4 functions with plasma membrane H+‐ATPases to regulate stomatal apertures during pathogen attack. PLoS Biology, 7, e1000139. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Liu, X. , Zheng, M. , Wang, R. , Wang, R. , An, L. , Rodermel, S.R. et al. (2013) Genetic interactions reveal that specific defects of chloroplast translation are associated with the suppression of var2‐mediated leaf variegation. Journal of Integrative Plant Biology, 55, 979–993. [DOI] [PubMed] [Google Scholar]
  21. Ma, X. , Zhang, Q. , Zhu, Q. , Liu, W. , Chen, Y. , Qiu, R. et al. (2015) A robust CRISPR/Cas9 system for convenient, high‐efficiency multiplex genome editing in monocot and dicot plants. Molecular Plant, 8, 1274–1284. [DOI] [PubMed] [Google Scholar]
  22. Nekrasov, V. , Staskawicz, B. , Weigel, D. , Jones, J.D.G. & Kamoun, S. (2013) Targeted mutagenesis in the model plant Nicotiana benthamiana using Cas9 RNA‐guided endonuclease. Nature Biotechnology, 31, 691–693. [DOI] [PubMed] [Google Scholar]
  23. Nelissen, H. , Clarke, J.H. , De Block, M. , De Block, S. , Vanderhaeghen, R. , Zielinski, R.E. et al. (2003) DRL1, a homolog of the yeast TOT4/KTI12 protein, has a function in meristem activity and organ growth in plants. The Plant Cell, 15, 639–654. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Nelson, T. & Dengler, N. (1997) Leaf vascular pattern formation. The Plant Cell, 9, 1121–1135. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Perico, C. , Tan, S. & Langdale, J.A. (2022) Developmental regulation of leaf venation patterns: monocot versus eudicots and the role of auxin. New Phytologist, 234, 783–803. [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. Petricka, J.J. , Clay, N.K. & Nelson, T.M. (2008) Vein patterning screens and the defectively organized tributaries mutants in Arabidopsis thaliana . The Plant Journal, 56, 251–263. [DOI] [PubMed] [Google Scholar]
  27. Pineau, E. , Xu, L. , Renault, H. , Trolet, A. , Navrot, N. , Ullmann, P. et al. (2017) Arabidopsis thaliana EPOXIDE HYDROLASE1 (AtEH1) is a cytosolic epoxide hydrolase involved in the synthesis of poly‐hydroxylated cutin monomers. New Phytologist, 215, 173–186. [DOI] [PubMed] [Google Scholar]
  28. Ravichandran, S.J. , Linh, N.M. & Scarpella, E. (2020) The canalization hypothesis ‐ challenges and alternatives. New Phytologist, 227, 1051–1059. [DOI] [PubMed] [Google Scholar]
  29. Ritz, C. & Spiess, A.‐N. (2008) qpcR: an R package for sigmoidal model selection in quantitative real‐time polymerase chain reaction analysis. Bioinformatics, 24, 1549–1551. [DOI] [PubMed] [Google Scholar]
  30. Rolland‐Lagan, A.‐G. & Prusinkiewicz, P. (2005) Reviewing models of auxin canalization in the context of leaf vein pattern formation in Arabidopsis. The Plant Journal, 44, 854–865. [DOI] [PubMed] [Google Scholar]
  31. Sachs, T. (1969) Polarity and the induction of organized vascular tissues. Annals of Botany, 33, 263–275. [Google Scholar]
  32. Sachs, T. (1981) The control of the patterned differentiation of vascular tissues. Advances in Botanical Research, 9, 151–262. [Google Scholar]
  33. Sack, L. & Scoffoni, C. (2013) Leaf venation: structure, function, development, evolution, ecology and applications in the past, present and future. New Phytologist, 198, 983–1000. [DOI] [PubMed] [Google Scholar]
  34. Scarpella, E. & Meijer, A.H. (2004) Pattern formation in the vascular system of monocot and dicot plant species. New Phytologist, 164, 209–242. [DOI] [PubMed] [Google Scholar]
  35. Smillie, I.R. , Pyke, K.A. & Murchie, E.H. (2012) Variation in vein density and mesophyll cell architecture in a rice deletion mutant population. Journal of Experimental Botany, 63, 4563–4570. [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Wang, P. , Kelly, S. , Fouracre, J.P. & Langdale, J.A. (2013) Genome‐wide transcript analysis of early maize leaf development reveals gene cohorts associated with the differentiation of C4 Kranz anatomy. The Plant Journal, 75, 656–670. [DOI] [PubMed] [Google Scholar]
  37. Weber, E. , Engler, C. , Gruetzner, R. , Werner, S. & Marillonnet, S. (2011) A modular cloning system for standardized assembly of multigene constructs. PLoS One, 6, e16765. [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Weigel, D. , Ahn, J.H. , Blazquez, M.A. , Borevitz, J.O. , Christensen, S.K. , Fankhauser, C. et al. (2000) Activation tagging in Arabidopsis. Plant Physiology, 122, 1003–1013. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Xiong, Y. , DeFraia, C. , Williams, D. , Zhang, X. & Mou, Z. (2009) Characterization of Arabidopsis 6‐Phosphogluconolactonase T‐DNA insertion mutants reveals an essential role for the oxidative section of the plastidic pentose phosphate pathway in plant growth and development. Plant and Cell Physiology, 50, 1277–1291. [DOI] [PubMed] [Google Scholar]
  40. Yamaguchi, A. , Kobayashi, Y. , Goto, K. , Abe, M. & Araki, T. (2005) TWIN SISTER OF FT (TSF) acts as a floral pathway integrator redundantly with FT. Plant and Cell Physiology, 46, 1175–1189. [DOI] [PubMed] [Google Scholar]
  41. Yan, L. , Wei, S. , Wu, Y. , Hu, R. , Li, H. , Yang, W. et al. (2015) High‐efficiency genome editing in Arabidopsis using YAO promoter‐driven CRISPR/Cas9 system. Molecular Plant, 8, 1820–1823. [DOI] [PubMed] [Google Scholar]
  42. Yoshida, Y. , Sarmiento‐Mañús, R. , Yamori, W. , Ponce, M.R. , Micol, J.L. & Tsukaya, H. (2018) The Arabidopsis phyB‐9 mutant has a second‐site mutation in the VENOSA4 gene that alters chloroplast size, photosynthetic traits, and leaf growth. Plant Physiology, 178, 3–6. [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Zapata, L. , Ding, J. , Willing, E.‐M. , Hartwig, B. , Bezdan, D. , Jiao, W.‐B. et al. (2016) Chromosome‐level assembly of Arabidopsis thaliana Ler reveals the extent of translocation and inversion polymorphisms. Proceedings of the National Academy of Sciences of the United States of America, 113, E4052–E4060. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

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

Supplementary Materials

Figure S1. Validation of T‐DNA insertion in AtDOT5 in the SALK_148869c line.

Figure S2. Snapshots of sequencing reads for genes flanking AtDOT5 in the Atdot5‐1 genome.

Figure S3. Confirmation of shared transgene position in Atdot5‐1 and Atdrl1‐1 mutants.

Figure S4. PCR assay to genotype gene edited alleles of AtDOT5.

Click here for additional data file. (2.1MB, docx)

Data Availability Statement

All data generated or analyzed during this study are included in this published article and its supplementary information files, except for the sequence reads of the Atdot5‐1 mutant genome which are available at ArrayExpress https://www.ebi.ac.uk/arrayexpress under accession number E‐MTAB‐12010.

Articles from The Plant Journal are provided here courtesy of Wiley

RESOURCES