Arabidopsis thaliana (Col-0) wild-type, transgenic and mutant plants were cultivated in pots with soil substrate (50% peat moss, 30% perlite and 20% sand) under a temperature of 22°C, humidity of 40% - 60% and 14 h light provided by white-colored LED panels at an intensity of 150 µmol m^-2 s⁻¹/10 h dark photoperiod. Seeds were surface sterilized with 70% ethanol for 1 min and then with 1% sodium hypochlorite for 20 min. After washing three times with distilled water, seeds were sown in Petri dishes on standard cultivation medium (SCM, 1/2 MS basal salts, 0.4 mg L^–1 thiamine HCl, 100 mg L^–1 myo-inositol, 10 g L^–1 sucrose, 7 g L^–1 agar). Plates were cold stratified at 4°C for 3 days, then moved to the cultivation chamber and kept under conditions mentioned above.

Genomic DNA was isolated from young rosette leaves using the innuPREP plant DNA kit (Analytik Jena, Jena, Germany) or with the CTAB method (Rogers and Bendich, 1989). For preparing the pSYT4-GUS transcriptional fusion construct, a 2,053-bp-long DNA stretch located upstream of the start codon was amplified by PCR from genomic DNA and inserted into the pPCV812 vector (Koncz et al., 1994) between the XhoI and BamHI restriction sites. For protein localization studies, we prepared pSYT4::SYT4::Dendra2::pASYT4 (hereafter abbreviated as SYT4-Dendra2) and pSYT4::SYT4::EGFP::pASYT4 (SYT4-GFP) transcriptional-translational fusion constructs. Genomic SYT4, with a 2,087 bp long DNA sequence located upstream of the start codon and a 485 bp long DNA stretch located downstream of the stop codon, was amplified by PCR from genomic DNA of Arabidopsis (Col-0). Dendra2 was amplified from p35S::Dendra2 (Jásik et al., 2013) and EGFP from pCS2 + EGFP-NB plasmids (Boggetti et al., 2012). Fragments were sequentially inserted into the pAMPAT-MSC vector (GenBank: AY436765.1) between the AscI and NotI sites. DNA stretches for Dendra2 and EGFP were attached to the 3′ end of the SYT4 genomic DNA sequence from which the stop codon was removed. All fragments were amplified using Q5^® Hot Start High-Fidelity DNA Polymerase and primers available in Supplementary Table 1 . All enzymes were purchased from New England BioLabs (Frankfurt, Germany). The correctness of sequences and read frames were verified by sequencing. Plasmids were transferred into the GV3101 (pMP90RK) Agrobacterium tumefaciens strain (Koncz and Schell, 1986), and this was used to transform Arabidopsis thaliana (Col-0) using the floral-dip method (Clough and Bent, 1998). The transformation was performed twice, and eight pots containing 5–8 Arabidopsis plants were used for each construct. Selection of primary transformants was carried out on the standard cultivation medium with 7.5 mg L^-1 phosphinothricin for the SYT4-GFP and SYT4-Dendra2 constructs and with 15 mg L^-1 hygromycin for the pSYT4-GUS construct. Both chemicals were purchased from Duchefa (Haarlem, The Netherlands). Based on the resistance, we selected 31 plants for pSYT4-GUS, 87 plants for SYT4-GFP, and 54 plants for the SYT4-Dendra2 construct. The expression of the fusion proteins was verified in offspring seedlings under an epifluorescence microscope, and the activity of the SYT4 promoter was monitored using a GUS histochemical assay. We then subjected the lines with signals to segregation analysis based on resistance (see above) to eliminate lines with multiple T-DNA inserts. We finally selected homozygous lines with a single insertion and obtained 11 homozygous lines with one insertion for pSYT4-GUS, 10 for SYT4-GFP, and 7 for SYT4-Dendra2. The presence of correct fusions in transgenic lines was verified by PCR. The SYT1-Dendra2 line was described previously (Lešková et al., 2019, Lešková et al., 2020).

Promoter activity in different organs was estimated using pSYT4::GUS transgenic plants in combination with histochemistry and fluorometry described previously (Jásik et al., 2011). For whole-mount histochemistry, seedlings growing in vitro were incubated in X-GlcA reagent mixture (1 mM 5-Bromo-4-chloro-3-indolyl-β-D-glucuronic acid [Duchefa, Haarlem, Netherlands] dissolved in 50 mM phosphate buffer (pH 7.3) supplemented with 10 mM EDTA, 2 mM ferrocyanide, 2 mM ferricyanide, 0.1% Triton X-100 [v/v] and 10% methanol). Samples were kept in the dark at 37°C. Blue-stained samples were treated with 70% ethanol to remove the chlorophyll and mounted to 80% glycerol for overall examination. Alternatively, after the GUS procedure, seedling parts were fixed with 4% glutaraldehyde, embedded in Stedman wax (Vitha et al., 2000), and sectioned. After dewaxing in ethanol, sections were stained with 0.1% basic fuchsine. GUS activity in complex organs of plants grown in the pots was analyzed by histochemistry, also using cryosections as described by Krausko et al. (2021). Shortly, cryosections collected in microcentrifuge tubes in cryostat were washed with MTSB, then treated with X-GlcA reagent mixture, and stained with 0.1% basic fuchsine. Sections were then washed and embedded in glycerol on microscopic slides. For the GUS fluorometry assay, samples of different organs of plants cultivated in the pots in growth stage 6.50 (50% of flowers to be produced have opened, Boyes et al., 2001) were homogenized in protein extraction buffer (50 mM phosphate buffer, 10 mM EDTA, 0.1% Triton X-100, 0.1% SDS) in 1.5 mL Eppendorf microcentrifuge tubes with LLG metal micro pestle fitted in a hand drill. Extracts were centrifuged at 16,000 x g and 4°C, and protein concentration in the supernatant was determined with a DC protein assay kit (Bio-Rad, Hercules, California, USA). Afterward, samples were diluted with extraction buffer to equal protein concentration. The reaction was carried out in 50 µL extraction buffer containing 10 µg total proteins and 2 mM 4-Methylumbelliferyl-ß-D-glucuronide trihydrate (4-MUG, Duchefa, Haarlem, Netherlands) in a black 96 well-plate (BRAND, Wertheim, Germany). The plates were incubated in the dark at 37°C, and after 2 h, the reaction was stopped with 200 μL 0.2 M Na₂CO₃. Fluorescence signal intensity was determined using a microplate reader Fluoroskan Ascent^® FL (Thermo Fisher Scientific, Waltham, USA), employing 355 nm excitation and 485 nm emission filters. The promoter activity in seedlings influenced by various factors was determined using the GUS fluorometric technique. Seedlings were grown on SCM in the growth chamber at 22°C under continuous light with 100 µmol m^-2 s^-1 light intensity. Petri dishes were kept in a vertical position. Ten-day-old seedlings were transferred to new SCM supplemented with different substances or kept under different conditions. After 24 h and 48 h, the roots and aboveground parts were separated, and intact samples were placed in 96-well microplates containing a 150 µl fluorometry extraction buffer with 2 mM MUG substrate. The microplates with samples were incubated in the dark for two hours at 37°C. The reaction was then stopped by adding 150 µl of 0.2 M Na₂CO₃, and fluorescence signal intensity was recorded as mentioned above.

For analysis of overall transcript abundance, total RNA was obtained from different parts of plants in growth stage 6.50, as characterized previously (Boyes et al., 2001). For transcript estimation in mutant alleles, RNA was isolated from roots or aboveground parts of seedlings originating from homozygous mutant plants (obtained as described below). RNA was isolated using the RNeasy plant mini kit (Qiagen, Hilden, Germany). DNA contamination was removed with a TURBO DNA-free™ kit (Thermo Fisher Scientific, Waltham, MA, USA). First-strand cDNA for transcript abundance analysis in different organs was synthesized from 500 ng of total RNA using a FIREScript® RT cDNA synthesis KIT (Solis BioDyne, Estonia) and a mix of oligo-dT and random primers. The first-strand cDNA for transcript analysis in mutants was synthesized using the same kit employing oligo-dT or oligo-dT and a random primer mix. Alternatively, cDNA was prepared using SOLIScript® RT cDNA synthesis KIT (Solis BioDyne, Estonia) and the gene-specific primer for SYT4 ( Supplementary Table 1 ).

Transcript abundance estimation in six different organs and mutant alleles (characterized below) was performed on Bio-Rad CFX Connect real-time PCR detection system with SsoAdvanced™ universal SYBR® green supermix. We carried out four independent biological replicates and two technical replicates. PROTEIN PHOSPHATASE 2A SUBUNIT A3 (PP2A, AT1G13320), MONENSIN SENSITIVITY1 (MON1, AT2G28390) and POLYUBIQUITIN 10 (UBQ10, AT4G05320) shown previously to have expression stability throughout development and under a range of environmental conditions (Czechowski et al., 2005) were used for normalization of gene expression. Sequences of primers are available in Supplementary Table 1 . Results were analyzed using the Bio-Rad CFX Manager 3.1 software, and expressions were quantified through the ΔCt method. Differences in relative expression between the organs were estimated through one-way ANOVA and Student's t-test. Differences in the abundance of transcript in roots of mutant alleles were evaluated also by qPCR as mentioned above. Additionally, we tested the presence of transcripts in roots and aboveground parts of seedlings of the mutant alleles using different combinations of primers and c-DNAs by semi-quantitative RT-PCR. Sequences of primers used in testing are listed in Supplementary Table 1 .

We purchased five alleles from The European Arabidopsis Stock Centre (NASC: http://arabidopsis.info/). SALK_201787C (named here syt4–1) was from the Salk collection (Alonso et al., 2003), two alleles SAILseq_652_E05.1 (syt4–2) and SAIL_359_H05 (syt4–3) from the Syngenta population (Sessions et al., 2002), and two alleles GK-215E11 (syt4–4) and GK668A12 (syt4–5) from Gabi-Kat collection (Rosso et al., 2003). Syngenta seeds for both alleles were sown in pots, and next-generation seedlings were subjected to segregation analysis using 7,5 mg/L phosphinothricin. Two lines for each allele showing resistance of all seeds were genotyped by PCR. We used combinations of gene-specific primers located upstream and downstream of expected insertion sites and SAIL-LB2 T-DNA insert-specific primers. Fragments were sequenced with the SAIL-LBJJ primer. Original seeds of the Salk syt4–1 allele have been sown in pots, and individual plants were genotyped with the SALK-LBJJ insert-specific primer and gene-specific primers. PCR fragments were sequenced with the SALK-LBb1.3 primer. Seeds of the syt4–4 and syt4–5 alleles were subjected to segregation analysis on SCM with 7.5 mg L^-l sulfadiazine. Two lines for each allele showing resistance of all seedlings were genotyped using combinations of gene-specific and GABI-LBJJ and GABI-RBJJ insert-specific primers. The PCR products obtained with GABI-LBJJ and gene-specific primers were sequenced with GABI-LBo8409 primer, the product of GABI-RBJJ primer and insert-downstream primer obtained in syt4–4 lines were sequenced with the GABI-RBo3144/35st primer.

Different plant organs and seedlings were processed according to Vitha et al. (2000). The samples were fixed with 4% formaldehyde in MTSB for 3 h, washed thrice with the same buffer, and dehydrated with ethanol and then embedded in Steedman’s wax. Six µm thick sections were prepared using a rotary microtome. Section ribs were mounted on glass covered with glycerol albumin and, after drying, dewaxed with ethanol and processed for immunolocalization. Mouse monoclonal antibody against Dendra2 (clone OTI1G6, Origene Tech. Inc., Rockville, MD, dilution 1:200) was combined with in rat grown antibodies against JIM13 (Knox et al., 1991) or LM5 (Jones et al., 1997) (1:100). Donkey anti-Mouse IgG (H+L) antibody, DyLight™ 550 (1:150, ab96876, Abcam, Cambridge, MA, USA) used for Dendra2 and Goat Anti-Rat IgG-FITC (1:150, OB3050–02, SouthernBiotech, Birmingham, AL, USA) for JIM13 and LM5 visualization. Alternatively, we combined goat Anti-Mouse IgG (whole molecule)–FITC antibody (F9006, Sigma-Aldrich) and Goat anti-Rat IgG (H+L), Alexa Fluor® 555 (1:150, A-21434, Invitrogen, Carlsbad, CA, USA).

All examinations were performed with Olympus FV1000 confocal laser scanning microscope. Seedlings were grown as described previously (Jásik et al., 2013; Lešková et al., 2019). The signal of aniline blue, sirofluor and aesculin were excited with a 405 nm laser and collected with 480–495 nm barrier filter, EGFP, the green population of Dendra2, and FITC were excited with an Argon/2 488 nm laser and captured with a 505 to 525 nm barrier filter, red form of Dendra2, basic fuchsine, DyLight™ 550 and Alexa Fluor^® 555 were excited with 543 nm laser and signal collected with a 560–620 nm barrier filter. ImageJ software was used to quantify the signal intensity.

Germination assay was performed with syt4–2 and syt4–3 mutants. Seeds of mutants and the SAIL wild type were sown in Petri dishes on SCM with 1% sucrose or without sucrose. After stratification as described above, Petri dishes were moved to permanent light (100 µmol m^-2 s^-1, 22°C) and the proportion of seeds with visible radicles was calculated after 24 and 48 h. In addition, the proportion of seedlings with hypocotyl and cotyledons fully emerged from the seed coat was evaluated after 48 h. To study the inhibitory effect of different factors on the elongation of primary roots, seeds of all five insertion mutant alleles and wild types from the same collection as mutant alleles were surface-sterilized and stratified, as described above. Seeds germinated vertically for another three days in permanent light to produce seedlings for experiments. Then, seedlings of the same size with roots approximately 5 mm long were transferred on the media with different factors in square Petri dishes. We tested the effects of IAA (0.1 µM), NAA (0.1 µM), ABA (20 µM), SA (20 µM), BAP (0.5 µM), mannitol (150 mM), NaCl (75 mM) and sucrose (0 and 6%). SCM with 1% sucrose was used as a control. Seedlings of individual mutant alleles were grown in the same Petri dish with corresponding wild-type seedlings. Twenty seedlings were used for each genotype and treatment, and the experiments were repeated minimally three times. Petri dishes were kept in a vertical position under the conditions mentioned above. After 3 days, the seedlings were photographed, and the root length increments were analyzed. Root lengths of mutants and wild-type plants affected by factors were related to root lengths of seedlings growing on SCM.

The figure images were processed by ImageJ, Adobe Photoshop CS2 (Adobe Systems, Mountain View, USA), and Microsoft Publisher software (Microsoft, Redmond, USA). The length of the roots was measured with ImageJ software using a neuroanatomy-SNT plugin. In charts, bars correspond to the standard errors of means. Significances between samples were analyzed by ANOVA and Student's t-test. Values in charts are given as the mean ± SE.

Since the expression of the SYT4 gene was not previously characterized, we analyzed it in different organs. We first studied SYT4 promoter activity using the uidA reporter gene that encodes beta-glucuronidase (GUS). Three homozygotic lines with a single copy of the T-DNA insert were used to analyze the activity of the SYT4 promoter in different organs by GUS fluorometry assay. Plants grew in pots and were in the middle flowering stage (stage 6.50, Boyes et al., 2001). The promoter activity patterns among organs were similar for all three lines; however, the fluorescence values among lines varied considerably. The highest promoter activity was detected in roots, less in inflorescence and stem, followed by siliques, and the lowest promoter activity was identified in both leaf types ( Figure 1A ).

Then, we investigated the activity of the promoter in different organs during development by GUS histochemical detection. Using the whole-mount GUS assay, we observed blue staining in vascular bundles of all plant organs ( Figures 1B–L ), and in addition in pollen grains of open flowers ( Figure 1L ), in vasculature at chalazal ends of ovules ( Figure 1M ) and seeds ( Figure 1N ), and floral organ abscission zones ( Figure 1L ). No promoter activity was detectable during embryo development and maturation ( Figure 1O ). However, when we analyzed imbibing seeds, we saw promoter activity in embryo roots and hypocotyls 24 h after sowing ( Figure 1P ). After 72 h, the staining was observable in vascular bundles of whole seedlings ( Figure 1Q ), although coloring was interrupted, especially in the cotyledons. In the seedling root tips, a clear signal was apparent already in isodiametric cells in protophloem (PP) files ( Figures 1B–D ) and more proximally also in incipient metaphloem cells (MP), companion cells (CC), phloem pole pericycle cells and cells of the phloem pole of procambium (PC; Figures 1B, C, E ). In several transgenic lines (three from eleven with one insertion of T-DNA), we observed intense staining in the root cap (RC; Supplementary Figure 2A ), but other lines show only a feeble reaction even after extended treatment (24 h). Several GUS-positive cell files were observed in the phloem of the root differentiation zone, and staining intensity varied among files ( Figures 1F, G ). No GUS reaction was detected in the lateral root primordium ( Supplementary Figure 2C ); however, intense staining was seen in emerged lateral roots in their central basal parts and developing PP ( Figure 1H ). In the leaf, the GUS reaction was observed mainly in the primary vein ( Figure 1I ). In the young, just emerging inflorescence stem (developmental stage 5, Boyes et al., 2001), GUS activity was limited to the phloem PC ( Figure 1J ). Secondary thickening stems had positive cells in the phloem area, but cells showed different staining intensities ( Figure 1K ).

At first, we analyze the pSYT4 DNA sequence used in our constructs employing the PlantCARE database of plant cis-acting regulatory elements (Lescot et al., 2002). We identified several cis-regulatory elements ( Supplementary Table 2 ). Besides five MYB recognition and binding sites and three MYC cis-elements that are responsive to different stresses, we also found one DRE1 drought responsiveness element, four ABRE abscisic acid-responsive elements, two ARE regulatory element essential for the anaerobic induction, one TGA auxin-responsive element, LTR cis-acting element involved in low-temperature responsiveness and finally the light-responsive regulatory elements such as G-box, AE-box, and TCT-motif.

To understand how the expression of SYT4 is regulated at the promoter level, we examined the effects of different factors on the promotor activity using GUS quantitative fluorometry. For experiments, we used 10-day-old seedlings that were large enough to produce a sufficient quantity of signal to perform analysis with single intact roots and aboveground parts. Seedlings were exposed to 21 treatments for 24 h and 48 h, and then the roots and aboveground parts were separated and immersed in the MUG reaction solution. Results are summarized in Figure 1R . Application of growth regulators, namely NAA, 2,4-D, kinetin, zeatin, BAP, and ABA, for 24 h and 48 h significantly reduced the promoter activity in roots; however, the activity of the promoter in the aboveground parts was unaffected ( Figure 1R ). The exposure of seedlings to osmotics, namely, PEG, sucrose, and mannitol, severely reduced SYT4 promoter activity in roots and excluded PEG also in aboveground parts ( Figure 1R ). Finally, also NaCl, cold and peroxide treatment dramatically decreased promoter activity in roots and aboveground parts ( Figure 1R ).

Furthermore, we wanted to know whether the overall promoter activity was reduced proportionally in the phloem throughout the root or only in its particular parts. Qualitative histochemical analysis showed that the decrease in promoter activity detected by the fluorometry was mainly due to the absence of promoter activity in the phloem of the matured root region. In contrast, the promoter was still active in the phloem in the root tip. This was evident in the case of 2,4-D ( Supplementary Figure 2F ), mannitol ( Supplementary Figure 2G ), and NaCl ( Supplementary Figure 2H ) treatment. Under NaCl and mannitol stress, we observed remarkable variability of SYT4 promoter activity among cells in the phloem files. In roots growing on SCM, the SYT4 promoter activity was not interrupted in matured roots, and the promoter strength did not change so drastically from cell to cell ( Supplementary Figure 2E , see also Figure 1B ).

In the next step, we estimated SYT4 transcript abundance in different organs of Arabidopsis, as in the case of the promoter, in the middle stage of flowering, as characterized previously (Boyes et al., 2001). We employed qPCR and primers designed to bridge exon 6/7 and exon 7/8 junctions to amplify specifically SYT4 cDNAs. Values normalized to three commonly used reference genes PP2A, MON1 and UBQ10 (Czechowski et al., 2005) are shown in Figure 1S . The SYT4 transcript was most abundant in the roots, followed by the stems. Rosette and cauline leaves showed the lowest transcript abundance.

To examine the SYT4 protein expression pattern at the cellular and intracellular level, we created homozygous transgenic plants expressing SYT4-EGFP and SYT4-Dendra2 fusions under the native SYT4 promoter and containing the endogenous 3′UTR sequence. In ovules and seeds, the fluorescence signal was detected only in the phloem at the chalazal end and funicules ( Figures 2A, B ). In developing embryos, fusion proteins first appeared in the shoot and root apical meristem cells at the late heart stage ( Figures 2C–E ) and in the PC tissue at the torpedo stage ( Figure 2F ). No signal was detected in fully developed dried seeds. A slight signal appeared in germinating embryos 24 hours after plating in the PC of hypocotyls and roots. After 48–72 h, the fusion protein was visible in immature PP elements of whole seedlings ( Figures 2G, H ) and detectable in the cells of shoot ( Figure 2I ) and root stem cell niche ( Figure 2J and Supplementary Figure 2B). Here, the protein was most abundant in the columella initials but was also present in the quiescent center cells (QC), endodermis/cortex and lateral RC/epidermis initials, and the outermost layer of the RC but not in initials in the stele. Then, we examined the SYT4 patterning during phloem development using cross-sections. In the distal part of the root tip, only PP elements showed a fluorescent signal ( Figure 2K ). More proximally to the shoot, the protein also appeared in phloem PC and MP and on their sides localized CC ( Figure 2L ). In young developing sieves, SYT4 became concentrated as patches at the cell peripheries ( Figure 2M, N ). When we analyzed a 1 cm long inflorescence stem (the principal growth stage 5, inflorescence emergence; Boyes et al., 2001), the stem was in the stage of primary growth, and we detected a strong signal in the phloem pole of PC ( Figure 2O ).

Precise cellular localization of SYT4 and its dynamics we studied in PP cells. Since the PP in the Arabidopsis root tip consists of only two columns of cells, these are readily observable, and the signal intensities of EGFP and Dendra2 are easily quantifiable with a confocal laser scanning microscope. We detected a slight signal already in the first early dividing PP cell approximately 50 µm from QC ( Figure 3A , see also Figures 2J, K ). However, incipient MP and PC file cells were negative within this distance from the QC. The signal was visible in PP meristematic cells as a network in the cytoplasm and a ring around the nuclei. It was also significantly enriched at the site of cell contacts of neighboring cells in the PP file ( Figure 3A ). The signal intensity increased dramatically in the more proximally located PP cells. However, as cells started to elongate and deposit callose in their transverse cell walls, the amount of SYT4 gradually decreased in the cells ( Figures 3B, C ). Using the SYT4-Dendra2 transgenic line, we demonstrated that the high abundance of SYT4 was due to its extensive synthesis in meristematic PP cells, as shown by the increasing intensity of the green signal emitted by the SYT4-Dendra2 population synthesized after the photoconversion. However, protein synthesis dropped in 11 to 14 cells in the PP column (Figures 3D, E). The abundance of the old red population of fusion protein synthesized before photoconversion was steadily reduced in differentiating SE.

We could not detect the signal in early lateral root primordia ( Supplementary Figure 2D ). However, the intense fluorescence appeared firstly in emerging lateral roots from parent roots in a group of cells in the central basal domain, and the weaker signal was seen in the central apical domain ( Figure 4A ). Here, SYT4 became visible during new root stem cell niche establishment ( Figure 4B ). Later, the signal was strong in the PP files, connected with the group of cells mentioned above ( Figure 4C ), and finally, in the cells connecting the SE of lateral and primary roots ( Figure 4D ). As described above, SYT4 was abundant in SE precursors and less profuse in maturing elements. We wanted to verify this finding by studying more developed organs. On the cross-section of differentiated root parts, cells exhibiting green fluorescence were scattered in phloem poles but were no longer present in the PP/MP areas ( Figure 4E ). We also observed the signal in several parallel cell files in the phloem; however, rarely in functional sieve tubes labeled by aesculin ( Figure 4F ). Then, we analyzed shoots and roots undergoing secondary growth. On the free-hand sections and cryosections, we saw SYT4 only in centripetally located cells of the phloem. Here, the green signal partially overlapped with the blue signal emitted by aniline-stained callose deposited in maturing sieves ( Figure 4G ). Centrifugally located cells showed only a blue signal. SYT4 distribution contrasted with the appearance of SYT1 protein, which was present in different tissues ( Supplementary Figure 2I ). Also, in the phloem, SYT1 was visible in all cell types ( Supplementary Figure 2J ), whereas SYT4 was accumulated at the periphery of particular phloem cells ( Supplementary Figure 2K ).

Since cells in the phloem of Arabidopsis organs are tiny and difficult to specify without fixation and proper sectioning, we employed histoimmunology techniques to identify cells expressing SYT4. We co-labeled sections of roots with the Dendra2-specific antibody and JIM13-specific antibody recognizing SE (Šamaj et al., 1998) and LM5-specific antibody recognizing CC (Torode et al., 2018), respectively. The signal for Dendra2 was localized within cells, with the cell walls labeled with JIM13 ( Figure 4H ). When we combined antibodies against LM5 and Dendra2, labeling was detected in distinct but adjacent cells in phloem islets in the inner part of the phloem ( Figure 4I ). However, in outside phloem layers, labeling of Dendra2 was not detected ( Figure 4J ).

Based on available data from the TAIR database, we selected five mutant alleles of SYT4 for deep analysis. By standard procedures, we isolated two homozygous lines for each selected allele. By PCR-based approach, we showed the presence of inserts and the absence of the entire SYT4 gene in all lines. The positions of primers are in Figure 5A , and the results of PCR genotyping are in Supplementary Figure 1 . Then, we re-evaluated the positions of T-DNA insertions by sequencing PCR fragments. We have established that four alleles, syt4–1, syt4–2, syt4–3 and syt4–5 contained complex T-DNA inserts. The inserts occurred as inverted T-DNA repeats around the right border, so they are bound to the SYT4 gene loci by left borders. The syt4–4 contains a single T-DNA molecule or direct T-DNA repeats with both LB and RB at junctions. Next, we verified the T-DNA insertion sites in the genomes and completed the missing T-DNA/plant DNA junction sequences for all mutant alleles. Flanking insertion sequences from the TAIR database indicated that the T-DNA insertion disrupted exon 10 in the Salk syt4–1; however, we identified that this allele had inverted T-DNA repeats in the 10th intron at 2144 bp downstream of the start codon. Also, the Syngenta syt4–2 allele with expected T-DNA insertion in the exon 10 had this insert actually in the 10th intron at 2193 bp downstream of the start codon and 23 bp were deleted during the integration process. As expected, the syt4–3 allele had integrated T-DNA insert in exon 11, precisely at 2637 bp downstream of the start codon and 10 bp were deleted. According to the original data, the syt4–4 GABI-Kat mutant should have integrated T-DNA in the second exon; however, we have found that this position was actually in the third intron at 448 bp downstream of ATG. Finally, in the syt4–5 GABI-Kat allele, the T-DNA was linked to the genome by one of the left borders in the intron 6 as previously determined, precisely at 1346 bp downstream of the start codon; however, the second LB of inverted repeat was attached to the genomic sequence at the beginning of exon 7 and 15 nucleotides were lost from the genomic sequence. The junction sequences, structures of T-DNA insertions and consequences caused by insertions in the mutant alleles are given in Supplementary Table 3 .

We then tested the presence and patterns of SYT4 transcripts in all syt4 alleles by RT-PCR. By qPCR we quantify the abundance of SYT4 transcript combining primers designed to bridge exon 6/7 and exon 7/8 junctions ( Figure 5A ) to anneal specifically to SYT4 cDNA. For the syt4–5 allele, we used a forward primer located slightly more upstream since the original forward primer partially overlapped the deleted DNA stretch. This procedure with primers that anneal in the middle of cDNA and PP2A as reference gene revealed more than 50 times increased abundance of SYT4 transcript in syt4–4 roots and almost 30 times decreased abundance of the transcript in syt4–3 roots, while in syt4–1, syt4–2 and syt4–5 transcript abundance was affected only slightly ( Figure 5C ). The pattern obtained in semi quantitative RT-PCRs with the same primers and 30 cycles shown in Figure 5B confirmed the synthesis of fragments of the correct size for SYT4 cDNA and the absence of genomic contamination.

Further, we tested all mutant alleles with different combinations of primers and an extended number of cycles (40–45) to see how SYT4 transcript look in different syt4 alleles. Positions of primers are accessible in Figure 5A . In alleles syt4–1 and syt4–2, which have insertions in the 10th intron, all combinations of primers from upstream of insertion sites produced fragments of the correct size for SYT4 cDNA. An example is presented in Figure 5D . If we employed the reverse primer, which bridges the exon 10/11 junction in combination with the forward primer positioned in the exon 10, upstream insertion site, we obtained the same fragment as in the wild type, and the fragment size corresponds to cDNA suggesting splicing in this site was not affected by T-DNA inserts in the intron 10. Genomic DNA does not produce the amplicon. When we analyzed the syt4–3 allele, which has an insertion in the exon 11, we obtained fragments with all primer pair combinations from upstream of the T-DNA insert. The patterns were not identical to those of wild-type plants because multiple amplicons were produced in the syt4–3 allele. An example is in Figure 5D . When we combined primers from both sides of the T-DNA insertion, we failed to amplify the fragment in mutant plants ( Figure 5D ). In the syt4–4 allele, which contained T-DNA in the third intron, RT-PCR fragments were obtained using primers annealing upstream or downstream of the insertion site; however, primer combinations from both sides of the T-DNA insert have not produced fragments ( Figure 5D ). In the syt4–5 allele, which has the T-DNA insertion at the intron 6/exon 7 border, all combinations of primers produced fragments of the correct size for SYT4 cDNA, similar to the wild type ( Figure 5D ).

Firstly, we looked at the appearance of syt4 mutant plants growing in in vitro conditions on SCM supplemented with 1% sucrose under a 14 h light at an intensity of 150 µmol m^-2 s^-1/10 h dark photoperiod. We did not observe changes in growth in any of the five alleles, but they resembled the wild type ( Figure 6A ). Then, we estimated plants growing in pots. We did not observe differences in phenotype between mutants and corresponding controls (syt4–3 mutant and control plants in the stage of fluorescence emergence are shown in Figure 6B ). Then, we compared more advanced plants (growth stage 6.50 as characterized by Boyes et al., 2001) and failed to find growth defects in insertional alleles ( Figure 6C ). We checked siliques and observed normal seed development in all mutant alleles.

Then, we estimated if a mutation in the SYT4 gene affects seed germination. When we analyzed the germination rate after 48 h, we failed to find differences between the wild type and two syt4 alleles ( Figure 6D ). However, observation after 24 h showed that the syt4–3 allele, but not the syt4–2 allele which show no altered transcript, is delayed in germination compared to the wild type ( Figure 6D ). This was confirmed by observing the proportion of fully emerged seedlings from seed coat after 48 h ( Figure 6D ), which was significantly lower in syt4–3 alleles than in the wild type. The presence or absence of sucrose did not play a role in these processes. Finally, we performed the primary root growth test. We took advantage of the knowledge from studying the SYT4 promoter and estimated the influence of factors affecting its activity to find a possible effect on the mutants’ phenotype. The growth increment of roots of syt4–1, syt4–2, and syt4–5 alleles was not different from wild types under any of the investigated factors ( Figure 6E ). When we analyzed the syt4–4 mutant, we observed a slight relative decrease in root elongation under conditions such as mannitol and NaCl; however, a statistically significant difference between the wild type and syt4–4 was detected only under the absence of sucrose ( Figure 6E ). The root growth in the syt4–3 mutant was significantly more reduced than in the wild type under the influence of all factors except BAP and ABA, and the difference was highly significant when seedlings were grown without exogenous sucrose ( Figure 6E ). The influence of sodium chloride is shown in Figure 6F . The importance of exogenous sucrose for the growth of syt4–3 and syt4–4 mutants is also confirmed by extended cultivation of seedlings in vitro. An increased number of stunted seedlings was apparent in both cases, and in the syt4–3 allele, the number of such seedlings was higher than in the syt4–4 allele ( Figure 6G ).