In plants the dorsoventral boundary of leaves defines an axis of symmetry through the centre of the organ separating the top (dorsal) and bottom (ventral) tissues. Although the positioning of this boundary is critical for leaf morphogenesis, how the boundary is established and how it influences development remains unclear. Using live-imaging and perturbation experiments we show that leaf orientation, morphology and position are pre-patterned by HD-ZIPIII and KAN gene expression in the shoot, leading to a model in which dorsoventral genes coordinate to regulate plant development by localizing auxin response between their expression domains. However we also find that auxin levels feedback on dorsoventral patterning by spatially organizing HD-ZIPIII and KAN expression in the shoot periphery. By demonstrating that the regulation of these genes by auxin also governs their response to wounds, our results also provide a parsimonious explanation for the influence of wounds on leaf dorsoventrality.https://doi.org/10.7554/eLife.27421.001
Lateral organ development in plants and animals typically involves several processes occurring in a coordinated manner. These include organ positioning, the specification of different cell types and organ morphogenesis. Spatial cues specifying these processes are usually provided by a molecular pre-pattern present in precursor tissues, or from inductive signals emanating from neighboring regions. Unlike animals however, plant organs such as leaves arise continuously in regular patterns around the shoot apical meristem (SAM). Nevertheless, certain features of leaves are relatively constant including the restriction of their formation to the meristem periphery and their flattened, dorsoventral (top-bottom) orientation with respect to the shoot apex. How are these fundamental features specified?
Since the 1950s wounding experiments involving the isolation of leaf primordia from the meristem have suggested the presence of an inductive signal from the meristem that promotes dorsal identity within leaf primordia at the time of organ initiation (Reinhardt et al., 2005; Sussex, 1951). A variant on this theme is the proposal that transient auxin depletion in the adaxial (adjacent to the shoot axis) tissues of leaf primordia promotes dorsal identity (Qi et al., 2014). In contrast, other studies suggest that dorsoventral patterning is pre-established, being directly derived from central-peripheral patterning of the shoot (Hagemann and Gleissberg, 1996; Husbands et al., 2009; Kerstetter et al., 2001; Koch and Meinhardt, 1994). Supporting this proposal is the observation that transcription factors involved in both dorsal and ventral cell fate in the leaf are expressed in the SAM in a central and peripheral manner respectively (Emery et al., 2003; Kerstetter et al., 2001; McConnell et al., 2001; Yadav et al., 2013). Thus, the manner in which leaf dorsoventrality is first established in leaves remains unresolved (Kuhlemeier and Timmermans, 2016).
To explain how leaf dorsoventrality regulates morphogenesis, Waites and Hudson, 1995 took a cue from wing development in Drosophila (Diaz-Benjumea and Cohen, 1993) and proposed that tissues located along the boundary between dorsal and ventral leaf tissues, i.e. along the leaf margin, act as organizers by producing mobile signals that pattern lamina growth (Waites and Hudson, 1995). So far, several genes have been identified that are expressed along this boundary including KLUH (Anastasiou et al., 2007) and the WOX family transcription WOX1, WOX2 and PRS (Haecker et al., 2004; Matsumoto and Okada, 2001; Nakata et al., 2012). The WOX genes in particular are required for proper lamina growth but are expressed only at the boundary region suggesting they may promote long-range patterning (Nakata et al., 2012). However although their ectopic expression can cause filamentous outgrowths at the leaf base, overall lamina growth is fairly normal (Nakata et al., 2012) suggesting that additional factors must direct patterning. The WOX genes are also known to regulate the expression of both dorsal and ventrally expressed transcription factors and miRNAs (Nakata et al., 2012; Zhang et al., 2014; Zhang et al., 2017) indicating that the WOX genes promote lamina growth at least in part by maintaining the integrity of dorsal and ventral expression domains. Overall, this leaves the question of how dorsoventral boundaries actually regulate morphogenesis still unanswered.
Besides influencing leaf differentiation and shape, genes involved in leaf dorsoventrality influence leaf position. For instance, Arabidopsis plants mutant for the KANADI (KAN) genes develop leaves ectopically from the hypocotyl and leaf tissues (Izhaki and Bowman, 2007) while plants mutant for the Class III HD-ZIP (HD-ZIPIII) genes develop leaves from the center of the shoot (Emery et al., 2003). These observations indicate that the developmental mechanisms that specify leaf dorsoventrality may also be involved in organ positioning although how these processes relate is unclear.
In this study we investigate the origin of dorsoventral patterning in detail. We show that new organs are centered on a pre-patterned boundary region located between the expression domains of HD-ZIPIII and KAN genes and that both KAN and HD-ZIPIII genes act to repress organogenesis where they are expressed. This leads to a model in which dorsoventral genes control leaf morphogenesis and positioning by localizing auxin transcriptional response, which then polarizes cells non-cell autonomously (Bhatia et al., 2016). However, we also show that dynamic auxin levels play a central role in determining boundary position by spatially organizing HD-ZIPIII and KAN1 expression in the meristem periphery. By demonstrating that the regulation of these genes by auxin also governs their response to wounds, our results also provide a parsimonious explanation for the influence of wounds on leaf dorsoventrality.
To better understand how leaf dorsoventrality is first established we used confocal microscopy to monitor the expression of genes involved in leaf dorsoventrality in the shoot apical meristem (SAM) in combination with the auxin efflux carrier PIN-FORMED1 (PIN1) using functional fluorescent protein reporters. In general, we found proteins involved in leaf dorsoventrality to be localized in non-overlapping concentric patterns with the dorsal Class III HD-ZIP protein REVOLUTA (REV) (Otsuga et al., 2001) being detected centrally, as marked by the expression of REVp::REV-2 × YPet (REV-2 × YPet), and the ventral protein KANADI1 (KAN1) (Kerstetter et al., 2001) expressed peripherally, as marked by the expression of KAN1p::KAN1−2 × GFP (KAN1−2 × GFP) (Figure 1, A to E). These concentric domains not only encircled the meristem but also extended contiguously around initiating leaves and floral bracts. Imaging PIN1p::PIN1-CFP (PIN1-CFP) (Gordon et al., 2007a) together with these reporters revealed regions of high PIN1-CFP expression, marking positions of organ inception (Heisler et al., 2005; Reinhardt et al., 2003), to be centered on a narrow region or ‘gap’ located between the dorsal and ventral domains (Figure 1D–K). As found previously, REV expression extended out into the PIN1-marked primordia over time (Heisler et al., 2005), reducing the ‘gap’ between the REV and KAN domains (Figure 1—figure supplement 1A–F,M–P). Nevertheless, at later stages a more substantial ‘gap’ reappeared between the REV and KAN1 domains (Figure 1C and D insets; Figure 1—figure supplement 1G–L and Q–R) indicating that the distance between the REV and KAN1 domains can change dynamically. Time-lapse imaging revealed that few cells alter their KAN1 expression during these early stages (Figure 2A–F) suggesting that the boundary of KAN1 expression in the SAM marks cells destined to form the future dorsoventral boundary of the organ.
Ventrally expressed MIR165/166 (Kidner and Martienssen, 2004; Merelo et al., 2016; Nogueira et al., 2007; Yao et al., 2009) also appeared active in the SAM periphery as marked by a MIR166Ap::GFPER reporter and MIR165/166 biosensor (Figure 3A–I), consistent with previous studies (Miyashima et al., 2013). Both KAN1−2 × GFP and MIR166Ap::GFPER re-established their expression in the SAM periphery after organ outgrowth (Figure 1E arrowhead; Figure 1—figure supplement 1I,K and L and Figure 3C arrowhead). As members of the WOX family of transcription factors including WOX1 and PRS are expressed at dorsoventral boundaries (Nakata et al., 2012) we imaged functional translational fusions to both PRS (PRSp::PRS- 2 × GFP) and WOX1 (WOX1p:: 2 × GFP-WOX1) to examine their expression in the shoot relative to leaves. We found that although 2 × GFP-WOX1 expression was limited to the margins and middle domain of leaves (Figure 4A and B) and absent in the inflorescence meristem (Figure 4C), in the vegetative shoot PRS-2 ×GFP expression extended from the leaf middle domain to surround the SAM in a region between the KAN1 and REV expression domains (Figure 4D–H). In the inflorescence meristem however, PRS-2 × GFP expression was restricted to early cryptic bract primordia (Figure 4I and J). In contrast to all genes described so far, a FILp::dsRED-N7 marker was expressed in both the abaxial (away from the shoot axis) and adaxial regions of the primordium at inception, consistent with a previous study (Tameshige et al., 2013) (Figure 5). Overall these data reveal that in many respects although not all, dorsoventral patterning within young leaf primordia, including the middle region (Nakata et al., 2012), corresponds to and is contiguous with, the patterning of dorsoventral gene expression in the SAM.
The finding that sites of organ inception marked by PIN1 are located between KAN1 and REV expression domains in the SAM suggest that both KAN1 and REV may act to repress organ inception where they are expressed. Supporting this proposal, leaves develop ectopically in kan1 kan2 kan4 and rev phb phv mutants (Izhaki and Bowman, 2007). However it is possible that the HD-ZIPIII genes influence organ initiation only indirectly by promoting SAM formation during embryogenesis (Emery et al., 2003; Izhaki and Bowman, 2007). To distinguish between these possibilities we induced the expression of MIR165/166-resistant REVr-2 ×VENUS throughout the epidermis using a pOp6/LhGR two-component (Samalova et al., 2005) system with the ATML1 promoter (Sessions et al., 1999) driving LhGR and found that it caused an arrest of organ formation and repression of KAN1−2 × GFP in both the vegetative and inflorescence meristems (Figure 6A–E). Despite the lack of organs, stem growth for the inflorescence meristem continued without any obvious change to meristem size, indicating that the phenotype initially influenced organ initiation (inset in Figure 6E). Similar results were observed after induction of a short tandem target mimicry construct designed to repress MIR165/166 activity (Yan et al., 2012) (Figure 6F and G) or after epidermal induction of MIR165/166-resistant PHAVOLUTA (Figure 6H–J). Similarly, plants expressing KAN1 ectopically in the epidermis also stopped making new organs (Figure 6K and L) and the inflorescence meristem took on a dome shape before eventually arresting (Figure 6M). We conclude both the KAN and Class III HD-ZIP genes regulate organ positioning at least in part by repressing organ initiation where they are expressed.
To test whether boundaries between KAN1 and Class III HD-ZIP expression in the SAM can play an instructive role in positioning new organs and determining their subsequent dorsoventrality, we induced KAN1−2 × GFP expression ectopically at the center of the SAM using a pOp6/LhGR two-component (Samalova et al., 2005) system and the CLV3 promoter driving LhGR. After KAN1−2 × GFP induction, most seedlings initiated several new leaves before their growth stopped. Confocal imaging of seedlings five days after stratification (DAS) on dexamethasone (DEX) induction medium revealed that new organs, marked by high levels of PIN1-CFP expression, formed ectopically at the perimeter of an enlarged and irregular central domain of induced KAN1−2 × GFP expression, in which REV-2 × YPet expression had been repressed (Figure 7A and B). Although endogenous KAN1 was not monitored, ectopic KAN1−2 × GFP was only detected within or bordering organs during their initiation while REV-2 × YPet expression was often restricted during later developmental stages (Figure 7C–H). This indicates that patterns of KAN1 gene expression within the SAM can influence subsequent organ development. In particular, we noted that the distal margins of developing leaf primordia always correlated with boundaries of REV expression within the epidermis, even when REV expression was abnormally restricted (arrow heads in Figure 7C–H; Video 1). Several classes of phenotype, including leaves with an inverted orientation, could be distinguished at maturity (Figure 7I–T) that correlated with the patterns of REV-2 ×YPet expression observed during early development. These distinct morphological classes can be explained according to the configuration of the HD-ZIPIII-KAN boundaries at organ inception. Specifically, we infer that the number and orientation of boundaries within organ founder cells (as specified by high auxin levels) determines the configuration of later leaf marginal tissue (Figure 7P–S).
The influence of dorsoventral gene expression on organogenesis suggests that dorsoventral boundaries may function generally to localize auxin response. In support of this proposal it has been reported that in the inflorescence meristem the auxin transcriptional marker DR5 is only responsive to auxin in the shoot periphery (de Reuille et al., 2006). We investigated DR5 expression (Liao et al., 2015) in the vegetative meristem by examining its expression both in the wild type and after 1-N-Naphthylphthalamic acid (NPA) treatment at 3 DAS. In mock-treated seedlings we found DR5 to be expressed at the locations of incipient primordia and at the distal tip of existing leaf primordia (Figure 8A and B). In contrast, NPA-treated seedlings expressed DR5 in a ring encircling the SAM prior to organ emergence and in the middle domain or dorsoventral boundary region of developing leaves (Figure 8C and D). Similar experiments with seedlings expressing the ratiometric R2D2 intracellular auxin reporter (Liao et al., 2015) revealed a generally broader distribution of auxin compared to the DR5 transcriptional response, especially after NPA treatment, where signal was not restricted to the meristem periphery or leaf tip (Figure 8E–H). These results indicate that like the inflorescence meristem, auxin transcriptional response outside the periphery of the vegetative meristem appears repressed. Since we also observed PRS expression localized to the meristem periphery (Figure 4D), we tested whether PRS as well as WOX1 are auxin inducible and found that both genes respond to auxin treatment within 12 and 15 hr respectively although their response was restricted to the boundary region (Figure 8I–P). Measuring transcript levels using qPCR indicated that the auxin response for both genes occurs at the transcriptional level (Figure 8Q).
These results not only reveal that general auxin response is maximized at the dorsoventral boundary but also, that genes already known to be expressed at the boundary are auxin responsive. Considering our results altogether and the finding that KAN1 represses PRS and WOX1 in ventral tissues (Nakata et al., 2012), we suggest a general scenario in which KAN and HD-ZIPIII genes repress auxin response where they are expressed, leaving a narrow domain of auxin responsive cells in between their expression domains.
We tested this proposal in silico by implementing a previous model for phyllotaxis (Jönsson et al., 2006; Smith et al., 2006), now supported experimentally (Bayer et al., 2009; Bhatia et al., 2016; Heisler et al., 2010), that incorporates the polarization of PIN1 towards cells with high intracellular auxin concentrations. By assuming that both KAN1 and REV repress auxin-induced transcription and by including a narrow region of cells located between the KAN1 and REV expression domains the model was able to self-organize the periodic formation of auxin maxima along the boundary as predicted, compared to a broader distribution of maxima when such boundaries are not included (Figure 9A–D; compare 9B to 9D).
So far, our results indicate that both HD-ZIPIII and KAN1 suppress auxin-induced gene expression where they are expressed. However this does not exclude the possibility that auxin may play a role in patterning HD-ZIPIII and KAN expression. Indeed, auxin promotes the expression of several HD-ZIPIII genes in rice (Itoh et al., 2008) and in Arabidopsis, REV expression typically extends towards PIN1 polarity convergence patterns in the meristem periphery and disappears in axil regions where auxin is depleted (Heisler et al., 2005). To further investigate any potential regulation of REV and KAN expression by auxin we treated inflorescence meristems with synthetic auxin NAA(1-Naphthylacetic acid)as well as auxin transport inhibitor NPA (N-1-Naphthylthalamic acid) to prevent rapid auxin redistribution and examined the response of both REV and KAN1. We found that the region in between the REV and KAN1 domains narrowed due to a significant expansion of REV expression within 15 hr of treatment (Figure 10A and B). REV also became expressed in cells between the primordia and meristem where REV had presumably reduced its expression previously (Figure 10C and D). This was confirmed by following cells over time where we found that although KAN expression appeared static, more cells expressed REV (Figure 10C–H). A more subtle response was observed as early as 6 hr upon combined NAA and NPA application (Figure 10—figure supplement 1). Local application of IAA to pin1-4 meristems also resulted in a local response by REV within 16 hr of auxin application (Figure 10—figure supplement 2). In contrast, exogenous auxin had little effect on a pCLV3:GFP-ER marker for central zone identity (Figure 10—figure supplement 3). Apart from changes in REV we noticed that over the course of 48 hr, KAN1 expression did not appear as expected in between the meristem and floral bracts of stages i1 and i2 (but not P1) at the time of treatment (Figure 10I and J), compared to mock treated plants (Figure 10K and H), where KAN expression appeared adjacent to the primordia within 24 hr. In vegetative meristems this was more obvious where contiguous REV expression often appeared between the meristem and developing leaves where KAN1 expression would normally be expressed (Figure 11A–H; Video 2). These results indicate exogenous auxin promotes HD-ZIPIII expression locally and that exogenous auxin can prevent the establishment of KAN1 expression if applied at stages i1 and i2 of organ development.
To test whether REV or KAN1 expression depend on endogenous auxin for their patterns of expression we treated triple marker plants with the TIR1 antagonist auxinole (Hayashi et al., 2012) and found that after 18 hr of treatment KAN expression had expanded slightly toward the meristem center (Figure 12A and B; Figure 12—figure supplement 1A–D). As auxinole competes with endogenous auxin for the TIR1 binding pocket, we attempted to reduce endogenous auxin signaling further by simultaneously treating the meristems with yucasin (Nishimura et al., 2014) and kyn (He et al., 2011) to block auxin synthesis. This combination of treatments led to an arrest of organogenesis and ‘pin’ phenotype as well as a much more significant expansion and restriction of KAN1 and REV expression centrally, respectively (Figure 12C and D; Figure 12—figure supplement 1E–H Figure 12—figure supplement 2). KAN expression not only expanded into those cells originally located between the REV and KAN1 domains but also cells that had previously been expressing REV at high levels. However KAN1 expression remained excluded from the central most cells of the shoot and established floral bracts and leaf primordia, where REV expression still remained (Figure 12D; Figure 12—figure supplement 2. Extending the drug treatment time did not lead to a further change in expression. Monitoring the ratiometric auxin sensor R2D2 in response to a similar combined inhibitor treatment confirmed a strong decrease in overall intracellular auxin levels, including in the central meristem region and maturing organs (Figure 12E and F), indicating that auxin independent mechanisms regulate the expression patterns of these genes in the central zone and in organs after their initiation. In contrast to REV and KAN1, auxin depletion did not influence size of the central zone, as marked by the expression of pCLV3:GFP-ER (Figure 12—figure supplement 3).
Overall these data indicate that dynamic changes in auxin levels specifically within the meristem periphery play a central role in regulating dorsal and ventral identities. While high levels of auxin act to both trigger and maintain HD-ZIPIII expression, this only occurs in cells not already expressing KAN1. Such high levels of auxin also prevent the acropetal expansion of KAN1 expression, not only in cells within primordia but also in regions between the primordia and meristem. As our auxin treatments only prevented KAN1 expression from establishing around primordia at stages, i1 and i2 and not around P1, which corresponds to a stage when auxin depletion has already occurred (Heisler et al., 2005), our data also suggest that once cells are exposed to low auxin levels, their fate becomes determined, i.e subsequent auxin exposure cannot reverse their trajectory towards ventral or peripheral cell identity. Outside the meristem periphery, both in the meristem central zone and within more established organs, REV and KAN expression appears comparatively auxin insensitive.
Since auxin is required to maintain REV at the expense of KAN expression in the meristem periphery and wounding causes repolarization of PIN1 away from wound sites (Heisler et al., 2010) resulting in auxin depletion (Landrein et al., 2015), we investigated whether wounding results in ectopic KAN1 and reductions in REV expression. Firstly, using a pulsed IR laser to ablate cells adjacent to young organ primordia we confirmed that auxin levels decrease in the vicinity of wounds by monitoring the expression of the R2D2 ratiometric marker (Figure 13A and B). Next, we monitored REV, KAN and PIN1 expression in response to such wounds. We found that KAN1 became expressed in cells adjacent to the wound on either side, regardless of wound orientation with respect to the SAM (Figure 13C and D; Figure 13—figure supplement 1A–H). Such a response argues against the possibility that ectopic KAN is the result of interruption of a signal emanating from the meristem that promotes dorsal and represses ventral identity (Sussex, 1955). Instead, it supports the proposal that KAN1 expression is promoted in the vicinity of wounds in general, possibly due to low auxin levels. To test this hypothesis, we repeated these experiments while treating the wounded meristems with combinations of NAA and NPA over a 48 hr period and found that the induction of KAN1 expression around wounds could be completely eliminated if NAA and NPA were combined (Figure 13E–G; Figure 13—figure supplement 1I–L). Although a similar response to wounding was found to occur in vegetative meristems, the wound response typically involved a more substantial reorganization of meristem structure, possibly due to the small size of the vegetative meristem relative to the wounds (Figure 13—figure supplement 1M–O). When new leaves subsequently formed, they were properly oriented with respect to the new meristem organization.
In this study we shed new light on a long-standing question regarding leaf dorsoventrality in plants – when and how is it established? The early work of Sussex, based on histological analysis and wounding experiments, suggested that initiating leaf primordia require an inductive signal from meristem tissues to specify dorsal cell fate (Sussex, 1955). This proposal has been further supported by a more recent study in tomato (Reinhardt et al., 2005). In contrast, other workers in the field have claimed that dorsoventrality arises directly from radial patterning of the shoot (Husbands et al., 2009; Kerstetter et al., 2001), with some authors also linking the organogenic capacity of the SAM peripheral zone to the meristematic nature of leaf margins (Hagemann and Gleissberg, 1996) and dorsoventral boundaries (Koch and Meinhardt, 1994). Our results support elements of these latter proposals by revealing that organs are pre-patterned by domains of KAN and HD-ZIPIII expression not only in terms of their differentiation but also their position and morphology. The effect of ectopic KAN1 expression at the centre of the shoot on subsequent leaf positioning and growth is particularly striking and indicates that the spatial arrangement of HD-ZIPIII and KAN expression present within organ founder cells is incorporated into organs as they initiate and directs patterns of morphogenesis (Figure 7P–S; Figure 14). However, the exact configuration of this pre-pattern, including its propagation into developing organs, is dynamic and depends on changing auxin levels, although this regulation is unlikely to be direct. While high auxin levels promote HD-ZIPIII and represses KAN1 this is only true for cells not already expressing KAN1. Hence, at sites of incipient organ formation, where auxin is concentrated in between the two expression domains, HD-ZIPIII expression is promoted and can expand peripherally, but only in cells adaxial to where KAN is expressed. At the same time, this high level of auxin prevents KAN expression from expanding acropetally into HD-ZIPIII expressing cells, causing the boundary to remain relatively static with respect to the underlying cells (Figure 14A). However if auxin levels drop, for instance when PIN1 reverses polarity away from cells between the meristem and primordium (Heisler et al., 2005; Qi et al., 2014), HD-ZIPIII expression decreases and KAN expression takes its place (Figure 14B and C). Hence high levels of auxin ‘lock-in’ the spatial arrangement of HD-ZIPIII and KAN expression within organ founder cells at organ inception until a later stage when the pattern becomes auxin independent. In between primordia, where auxin levels are lower, cells continuously transition from expressing REV in the central zone (where HD-ZIPIII expression appears auxin insensitive) to expressing KAN1 as they are displaced peripherally, as also indicated by the expression patterns of these genes in pin1 meristems (Figure 10—figure supplement 2A,E and I).
Overall these results imply that the dorsoventral patterning of organs results from a four-step process: (i) signals during embryonic development establish concentric patterns of Class III HD-ZIP and KAN gene expression; (ii) the boundary between these domains helps to define sites for auxin-dependent organogenesis i.e. the meristem peripheral zone; (iii) As organs form, dynamic auxin levels modulate the patterns of HD-ZIPIII and KAN expression, thereby helping to position new organs and establish their dorsoventrality; (iv) patterns of HD-ZIPIII and KAN gene expression within organs are stabilized and dictate future patterns of morphogenesis.
The regulation of HD-ZIPIII and KAN expression by auxin is not only relevant to understanding wild type organ development but also for understanding the reorganization of tissue types in response to wounds. Wounds in the meristem outer cell layer specifically alter cell polarities such that PIN1 becomes polarized away from wounds in adjacent cells (Heisler et al., 2010), leading to auxin depletion (Landrein et al., 2015). Consistent with this, we observe KAN1 to be upregulated and REV to be repressed around wounds. Furthermore, such changes can be completely blocked by auxin application. Altogether then, our results strongly indicate that wounds re-pattern peripheral cell identities due to the depletion of auxin in their vicinity. Specifically, we propose that auxin depletion triggers a local commitment to peripheral cell fate (which includes expressing KAN1), thereby providing a parsimonious explanation for the wound-associated disruptions to leaf polarity observed by Sussex and others (Sussex, 1955). However further experiments using the vegetative meristems of plants with larger meristems will be needed to confirm this proposal since we were not successful in altering the dorsoventrality of Arabidopsis leaves by wounding and therefore could not test whether auxin treatment could prevent such alterations.
Our conclusion that auxin promotes dorsal cell fate while inhibiting ventral cell fate contrasts with those of another study reporting that transient high levels of auxin inhibit establishment or maintenance of dorsal fate and/or promote establishment or maintenance of ventral cell fate (Qi et al., 2014). These findings were based on auxin application experiments in tomato that resulted in the ventralization of leaves as well as the observation that Arabidopsis pin1 and pid rev double mutants produce trumpet or rod-shaped shaped leaves. Although our results cannot easily explain the tomato data, since we find that high auxin levels are required to maintain REV expression during organ establishment, we would expect that in pin and pid mutants, lower auxin levels may well result in lower expression levels of REV and possibly other Class III HD-ZIPs, leading to leaf ventralization as reported in these mutant backgrounds (Qi et al., 2014). Further auxin application experiments in tomato that include an analysis of gene expression and auxin level changes may clarify the relationship between auxin and dorsoventral patterning in tomato compared to Arabidopsis.
How do DV boundaries control organ position and shape? Both our data as well as data from previous studies suggest this is through the regulation of auxin perception. For instance, organ initiation requires high auxin levels but auxin can only trigger organogenesis in the peripheral zone (Reinhardt et al., 2000) where the HD-ZIPIII/KAN boundary occurs. A similar relationship holds for leaves where auxin application results in growth but only from the leaf margins, again corresponding to a DV boundary (Koenig et al., 2009). We further show that the localization of auxin response to DV boundaries applies generally, i.e. DR5 expression appears higher at such boundaries compared to the broader predicted auxin distribution, as marked by R2D2. How is this restriction achieved? Imaging data reveals that the locations of PIN1 polarity convergences correspond to a region of cells in between the expression domains of HD-ZIPIII and KAN where their expression is low or absent. Given our results as well as genetic and molecular data indicating that both the HD-ZIPIII and KAN genes repress auxin activity (Huang et al., 2014; Merelo et al., 2013; Müller et al., 2016; Zhang et al., 2017), we propose that this localized absence of expression results in a local de-repression of auxin-induced transcription. Such localized auxin activity has recently been shown to orient cell polarity, including microtubule orientations, in a non-cell autonomous manner (Bhatia et al., 2016). Hence, we suggest that the proposed ability of dorsoventral boundaries to act as organizers analogous to those of the Drosophila wing (Diaz-Benjumea and Cohen, 1993; Waites and Hudson, 1995) rests in-part on their ability to orient cell polarity non-cell autonomously by localizing auxin activity (Figure 14D). The resulting growth may occur either in a periodic fashion along the boundary typical of phyllotaxis and complex leaves, or in a more continuous manner typical of simple leaves, depending on the strength of auxin transport or other modifications to the cell polarity feedback system (Bilsborough et al., 2011; Koenig et al., 2009). Investigating the dynamic consequences of directly juxtaposing REV and KAN expression should enable the testing of these hypotheses as well as lead to a better understanding of how auxin-responsive boundary domains are first established.
Plants were grown on soil at 22°C in continuous light-conditions and cultivated either on soil or on GM medium (1% sucrose, 1 × Murashige and Skoog basal salt mixture, 0.05% MES 2-(MN-morpholino)-ethane sulfonic acid, 0.8% Bacto Agar, 1 % MS vitamins, pH 5.7 with 1 M potassium hydroxide solution).
Multiply transgenic lines were generated by Agrobacterium-mediated transformation into stable transgenic lines or by genetic crossing. The FILp::dsREDN7 and PIN1p::PIN1-GFP transgenes have been described elsewhere (Heisler et al., 2005). pREV::REV-2×VENUS in the T-DNA vector pMLBART (Gleave, 1992) is a modification of pREV::REV-VENUS (Heisler et al., 2005) that contains a translational fusion to two tandem copies of the fluorescent protein VENUS (Nagai et al., 2002). REVp::REV-2×Ypet containing a C-terminal fusion to the 2 × Ypet (Nguyen and Daugherty, 2005) in pMOA36 T-DNA (Barrell and Conner, 2006) was transformed into a PINp::PIN1-CFP line (Gordon et al., 2007b). The KAN1p::KAN1−2 × GFP transgene in pMOA34 T-DNA was created by amplifying 8.7 kb of KAN1 (At5g16560) genomic sequences with primers KAN1g F and KAN1g R (Supplementary file 1A) as a translational fusion to a 9 Ala linker and 2 × GFP followed by OCS terminator sequences. When transformed into kan1-2 kan2-1 segregating plants, this construct complements the mutant phenotype. The triple marker line was generated by transforming KAN1p::KAN1−2 × GFP into a REVp::REV-2 × Ypet; PIN1p::PIN1-CFP transgenic line. KAN1p::KAN1−2 × CFP or KAN1p::KAN1−2 × Ypet containing a fusion to 2 copies of CFP or Ypet, respectively, were constructed similarly. KAN1p::KAN1−2 × Ypet and PIN1p::PIN1-GFP were combined in T-DNA vector BGW (Karimi et al., 2002) by Gateway technology (Invitrogen) for generation of a double marker transgenic line.
The KAN1 cDNA was amplified by PCR with primers K1 cDNA F and K1 cDNA R to generate C-terminal translational fusion to a 9 Ala linker followed by single GFP or 2 × GFP followed by pea rbcS E9 terminator sequence (Zuo et al., 2001) and cloned into the pOp6/LhGR two-component system (Craft et al., 2005) for dexamethasone-inducible misexpression.
An ATML1p::LhGR driver containing 3.4 kb of the L1-specific ATML1 gene (At4g21750) fused to the chimeric LhGR transcription factor and a 6Op::KAN1-GFP expression construct in a pSULT sulfadiazine-resistant T-DNA vector (ATML1 >>KAN1 GFP) was generated. The pSULT T-DNA vector was derived from pMLBART by replacing the NOSp::BAR gene with 1’−2’p::SUL (Rosso et al., 2003), a plant selectable marker that confers resistance to sulfadiazine herbicide to create pSULT. A CLV3p::LhGR driver containing 1.49 kb of upstream regulatory sequences was PCR amplified with primers CLV3p F and CLV3p R along with 1.35 kb of downstream regulatory sequences with primers CLV3utr F and CLV3utr R was combined with 6Op::KAN1−2 × GFP in pSULT T-DNA vector (CLV3 >>KAN1-2×GFP).
ATML1 >>REVr-2×VENUS is a sulfadiazine-resistant T-DNA vector to misexpress microRNA resistant REV-2 ×VENUS fusion, where 6Op::REVr-2×VENUS was constructed by cloning a 1148 bp BamHI-XcmI microRNA resistant REV cDNA (a gift from J. Bowman) harbouring two previously characterized silent mutations that disrupt the binding of MIRNA 165/166 to the coding sequence of REV as previously described (Emery et al., 2003) downstream of the 6Op and in frame with the wild type REV-2 ×VENUS coding sequences. miR166Ap::GFPER T-DNA construct was kindly provided by K. Nakajima (Miyashima et al., 2011). The MIR165/166 biosensor was created based on the design presented by Smith Z. R. et al. (Smith and Long, 2010) in the AlcR/AlcA expression system (Roslan et al., 2001) for ethanol-inducible expression. The sequences conferring MIR165/166 sensitivity from the REV coding sequence (REV) and the sequences conferring MIR165/166 insensitivity (REVr) were fused to mCherry (Shaner et al., 2004) with endoplasmic reticulum localization sequences mCherryER, which was synthesized de novo (Genscript). The MIR165/166-mCherryER biosensors (both biosensor and control) were cloned as HindIII-BamHI fragments downstream of the AlcA regulatory sequences in the UBQ10p:AlcR_BJ36 plasmid vector. UBQ10p:AlcR was constructed by cloning the UBQ10 promoter 2 kb fragment upstream of AlcR and the OCS terminator. Both UBQ10p::AlcR and AlcA::REV-mCherryER or AlcA::REVr-mCherryER components were combined in the T-DNA vector pMOA34.
The WOX1p::2 × GFP-WOX1 construct in pMLBART T-DNA vector was generated as follows: 2.2 kb of WOX1 (At3g18010) upstream promoter sequence was amplified with primers WOX1p F and WOX1p R and cloned using restriction enzymes KpnI and BamHI. 3.6 kb of WOX1 coding sequence plus 1.65 kb 3’-regulatory sequences was amplified from wild-type Col-O genomic DNA with the primers WOX1g F and WOX1g R and cloned using restriction enzymes BglII and SpeI. 2 copies of GFP were inserted in frame at the start of the WOX1 coding sequence at the BamHI and BglII sites. A double marker was generated by transforming the WOX1p::2 × GFP-WOX1 into a PIN1p::PIN1-CFP transgenic line.
The PRSp::PRS-2×GFP construct in pMOA34 T-DNA vector was made by amplification of 3.9 kb PRS (At2g28610) genomic sequence (similar to [Shimizu et al., 2009]) with primers PRSg F and PRSg R to create a C-terminal fusion to 2 × GFP followed by OCS 3’regulatory sequences. Marker combinations were generated by transforming the PRSp::PRS-2×GFP into either a PIN1p::PIN1-CFP transgenic line or into REVp::REV-2×YPET PIN1p::PIN1-CFP line. (PRSp::PRS-2×GFP) and (KAN1p::KAN1−2 × CFP) were combined in T-DNA vector BGW (Karimi et al., 2002) by Gateway technology (Invitrogen) for generation of a double marker transgenic lines.
A short tandem target mimic (STTM) construct to target MIR165/166 (Yan et al., 2012) was generated in the pOp6/LhGR two-component system for dexamethasone-inducible expression with a UBQ10p::GRLh driver. STTM MIR165/166-88 sequence (Yan et al., 2012) was synthesized de novo (Genscript) and cloned downstream of 6 × Op to create 6 × Op::STTM 165/166. Both components were combined in a sulfadiazine T-DNA pSULT (UBQ10 >>STTM 165/166).
ATML1 >>PHVr is a sulfadiazine-resistant T-DNA vector containing a mutated version of PHV cDNA (a gift from J. Bowman) with a Gly to Asp amino acid change that disrupts the miRNA165/166 binding site in the PHV gene (McConnell et al., 2001). 6 × Op::PHVr was constructed by cloning a 2.6 kb XhoI-BamHI PHVr cDNA downstream of 6 × Op and upstream of pea3A terminator sequences. ATML1 >>PHVr was transformed into a REVp::REV-2×YPet; PIN1p::PIN1-CFP transgenic line.
DR5−3 × VENUS v2 reporter gene (Liao et al., 2015) was a generous gift from Prof Dolf Weijers (Wageningen University). The line R2D2 PIN1p::PIN1-GFP was described previously (Bhatia et al., 2016). The seeds for pin1-4 mutant were a kind gift from Prof. David Smyth (Monash University) and the heterozygote plants were crossed wild type plants carrying pREV::REV-2xVENUS and pKAN::KAN-2xGFP. pCLV3::GFP-ER has been described previously (Gordon et al., 2009).
For inducible gene perturbations in the vegetative SAM, seeds were germinated directly on GM medium containing 10 μM Dexamethasone (Sigma, stock solution was prepared in Ethanol). Seedlings were then dissected for imaging at 4 DAS, 5 DAS or 7 DAS depending on the experiment. For DEX induction in the IM, 10 μM DEX solution containing 0.015% Silwet L-77 was applied to the IM every second day three times. Inflorescences were then dissected and imaged. The number of T2 inducible transgenic lines that exhibit the presented phenotypes and the frequencies of phenotypes amongst imaged plants is shown in Supplementary file 1B and associated caption.
Plants were dissected for imaging as previously described (Bhatia et al., 2016; Heisler and Ohno, 2014) and imaged with a Leica SP5 Upright confocal microscope using an Argon laser. The objective used was a water-immersion HCX IRAPO L25x/0.95 W (Leica). Excitation for CFP is 458 nm, GFP is 488 nm, YFP (YPet and VENUS) is 514 nm and tdTomato is 561 nm. Emission signal were collected at 460–480 nm for CFP, 490–512 nm for GFP, 516–550 nm for YFP (YPet and VENUS), and 570–620 nm for tdTomato. The resulting z-stacks were processed using the Dye Separation (Channel mode or automatic mode) function available in the LAS AF program in order to separate the GFP channel from the YFP (YPet or VENUS) channel. Three software packages: LAS AF from Leica, Imaris 8.0.2 by Bitplane and FIJI (https://fiji.sc) were used for data analysis. Ratios for R2D2 were calculated as described previously (Bhatia et al., 2016).
For distance measurements between oppositely positioned leaves on plants transgenic for inducible KAN1 (CLV3 >>KAN1-2×GFP – see above) the measurement tool from Imaris 8.0.2 (Bitplane) was used. For comparisons to control, untreated seedlings grown on GM were compared to seedlings grown on DEX for 5 days. t-Test was performed using Excel.
500 mM stock solutions of auxinole, yucasin and L-Kynurenine were prepared separately in DMSO. The stocks were diluted in 1 mL 0.1M phosphate buffer in sterile water to make a working solution containing all the three drugs (0.2 μL of each stock) to a final concentration of 100 mM each. The final concentration of DMSO in the working solution containing all the three drugs was 0.06%.
Treatments were carried out on the inflorescence meristems of whole plants transplanted from soil to boxes containing GM medium supplemented with vitamins. The older flowers were removed as described (Heisler and Ohno, 2014). The plants were chosen such that the stem of the meristem was a few millimeters above the rosette to prevent the drug solution from dispersing into the surrounding medium. After imaging, the meristems were carefully dried using a thin strip of sterile filter paper to remove excess water. Approximately 50 μL of the drug solution was added directly to the meristem, drop-wise. The meristems were treated only once in a time course of 12–18 hr.
Seedling aged 3 DAS were dissected to expose the meristem and the first leaf pair as described (Bhatia et al., 2016). After imaging, seedlings were transferred to new GM medium containing plates and blotted dry with thin strips of sterile paper. 5–10 μL of 100 μM NPA in sterile water (100 mM stock in DMSO) was added directly to the dissected seedlings every twenty-four hours for three days in total.
Seedlings aged 4DAS (days after stratification) were dissected to expose the meristem and the first leaf pair as described (Bhatia et al., 2016). 5 mM NAA (1M stock in 1M KOH) solution was prepared in liquid GM medium. Seedlings were then immersed in 100 μL of NAA containing medium in individual wells in 96 well plate and grown under continuous light without shaking for 12 hr.
For treatments on the vegetative meristems, 3DAS old seedlings expressing REV, KAN and PIN1 markers were dissected and imaged first. Samples were then treated with a combined solution of 100 μM NPA and 5 mM NAA (both diluted in water) or 100 μM NPA alone or 5 mM NAA alone. Samples were imaged and treated every 24 hr for 72 hr in total. For treatments on the infloresecnce meristems, flowering plants expressing REV, KAN1 and PIN1 markers and CLV3 GFP-ER marker were transferred from soil to boxes containing GM vit medium as described above. The infloresecnce meristems were dissected to remove older flower buds covering the meristem. The meristems were imaged, dried and treated with 10 μL of combined solution of 100 μM NPA and 5 mM NAA. Plants were imaged at the required timepoints (6 hr, 15 hr and 48 hr). Plants were treated with NAA and NPA after every timepoint imaging.
pin1-4 mutant plants expressing REV and KAN1 markers with visibly smooth pin meristems were transferred from soil to boxes containing GM-vitamin medium. Leaves covering the meristems were dissected away. Control paste was applied locally to follow the meristem orientation over time. The meristems were imaged, dried and lanolin paste carrying 5 mM IAA (3-Indoleacetic acid) was administered locally at the meristem periphery. Plants were then imaged 15–17 hr later to follow REV response at the site of auxin paste application. For this, first a z-stack representing an overview of the meristem was recorded. The paste was then scrapped off using a microfiber of the blotting paper carefully without damaging the meristem. A z stack was recorded again to follow REV and KAN expression at the site of paste application. A separate set of treated plants were imaged 24 hr later to follow REV and KAN expression in the grown out primordia at the site of auxin paste application.
Laser ablations on the inflorescence meristems were carried out using the Mai Tai multi-photon laser from Spectra Physics, which was controlled with LEICA SP5 confocal software. Z-stacks were acquired prior to ablation. Single cells were targeted one after the other using bleach point mode. Ablations were carried out at 800 nm with an output power of ~3W. Each pulse was shot for 1–15 milliseconds. Usually ablations were accomplished within 1–3 bursts of the laser. Ablated cells could be visually identified as their nuclei exploded resulting in unusual auto fluorescence. Z stacks were acquired immediately after the ablations.
After ablations, the meristems were carefully blotted dry using thin strips of sterile filter paper. 20 μL 5 mM NAA in sterile water (0.5M stock in 1M KOH) or 20 μL of a solution containing 5 mM NAA and 100 μM NPA in sterile water (100 mM NPA stock in DMSO) or mock solution were added directly to the meristems every 24 hr for 48 hr in total.
We developed a computational model to understand how interplay between the HD-ZIPIII and KAN pattern and PIN1 dynamics can influence auxin transport and primordia initiation. The model introduces a dependence on KANADI and REVOLUTA into previous models describing PIN1 and auxin dynamics (Bhatia et al., 2016; Heisler et al., 2010; Heisler and Jönsson, 2006; Jönsson et al., 2006; Sahlin et al., 2009). In the model, auxin resides in cell compartments and is able to move between cells either via active transport mediated by PIN1 proteins or passively via a diffusion-like process. PIN1 proteins cycle between cytosol and membrane compartments and a quasi-equilibrium model is used for determining its membrane localization at any time point. Auxin generates a signal able to polarize PIN1 in neighboring cells, i.e. a high auxin concentration increases the amount of PIN1 proteins in the neighboring cell membrane facing that cell. The molecule X in our model acts as a mediator of the signalling between auxin and PIN1, and the signal has previously been interpreted as a molecular (Jönsson et al., 2006), or mechanical stress signal (Heisler et al., 2010). In the model, the signal X is activated by auxin, and repressed by KAN and REV. The equations governing the dynamics of the molecules are
where is the auxin concentration and is the level of the signalling molecule in cell is the set of cells neighboring cell is the cell volume of cell , and is the cell wall area for the wall section between cells and is the total PIN1concentration in the cytosol and membrane compartments of cell . Membrane-bound PIN1 appears in the equation as , which is the PIN1 concentration in the membrane compartment of cell i that faces cell . A simple linear feedback between the signal and is used and a quasi-stable assumption, introduced in Jönsson et al. (2006), leads to
where is the ratio of endocytosis and exocytosis rates and sets the relation between symmetric and polarized exocytosis (, where is the PIN1 in the cytosol compartment). The feedback between auxin and PIN1 is identical to previous models if the KAN and REV factors in the equation are 1 (e.g. by setting KAN and REV to zero in all cells), while the polarising signal becomes reduced in regions where KAN and/or REV are expressed. This effect tunes the interaction between the dorsoventral patterning and PIN1/auxin dynamics (cf. Figure 8B and D).
The model was simulated using the in-house developed software tissue (available at the gitlab repository https://gitlab.com/SLCU/TeamHJ/tissue; please see README.txt file in Supplementary file 2 for details). A copy is archived at https://github.com/elifesciences-publications/TeamHJ-tissue. Files defining the model with parameter values (Supplementary file 1C) and initial configuration of (static) cell geometries and KAN and REV expression domains are provided in Supplementary file 2. The simulations use a 5th order Runge-Kutta solver with adaptive step size (Press et al., 2007), and initial auxin, PIN1 and X concentrations are set to zero in all compartments.
The model defined above was run on a template containing a predefined KAN/REV pattern (provided as Supplementary file 1C, Figure 9C). The geometry of the template was generated by a combination of cell/wall growth and mechanical interactions together with a shortest path division rule (Sahlin and Jönsson, 2010). A KAN/REV pattern was generated by the equations
This system was run to equilibrium on the above-mentioned template. In the above equations ri is the distance of cell i from the center of the template. The parameters were set to VK = VR = dK=dR = 1, KK = 30, KR = 25, nK = nR = 20. These parameters are set such that two distinct domains are created, with a small overlap of low KAN and REV concentrations in the boundary between these regions. To make the transition of KAN concentrations between domains sharper, KAN concentrations were set to 0 (1) if Ki ≤0.5 (>0.5).
4 day-old wild-type Ler seedlings were immersed into 5 mM NAA solution in liquid GM medium, and grown under continuous light without shaking for 15 hr. Cotyledons, hypocotyl, and roots were removed under dissecting scope, and only shoot meristem and the first pair of leaves were collected and immediately frozen in liquid nitrogen. Each biological replicate, represents tissue from 10 to 15 individual seedlings. Five biological replicates were collected for both mock and NAA treatment. RNeasy Mini kit (Qiagen) was used according to manufacturer’s instruction for RNA extraction. 1 microgram of RNA was used for cDNA preparation using Super script III reverse transcriptase for Q-PCR analysis. Q-PCRs were performed in a StepOne Plus Real Time PCR system thermo cycler (The applied bio systems) using 20 μl of PCR reaction containing 10 μL of SYBR Green mix (Roche), 1 μl of primer mix (10 μm), 2 μl of 1:10 diluted cDNA and 7 μl of water. Transcript levels were normalized to ACTIN2 transcript levels. Data was analyzed using the 2−ΔΔCT method. A freely available online tool was used for analysis using an unpaired t-test of the RT-PCR results: http://graphpad.com/data-analysis-resource-center/. For p-value calculation, data entry format with mean, SD and N was used. Measurements and calculations for all replicates are provided in Figure 8—source data 1.
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Dominique C BergmannReviewing Editor; Stanford University/HHMI, United States
In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.
[Editors’ note: this article was originally rejected after discussions between the reviewers, but the authors were invited to resubmit after an appeal against the decision.]
Thank you for submitting your work entitled "Cell type boundaries organize plant development" for consideration by eLife. Your article has been reviewed by three peer reviewers, one of whom is a member of our Board of Reviewing Editors and the evaluation has been overseen a Senior Editor. The reviewers have opted to remain anonymous.
Our decision has been reached after consultation between the reviewers. Based on these discussions and the individual reviews below, we regret to inform you that your work will not be considered further for publication in eLife.
The reviewers and editors all agreed that this work focused on a long–standing issue of organ formation in plants and that the manuscript, using beautiful imaging and side–specific markers, presented convincing and high quality evidence for the presence of the boundary within the SAM and the influence of the location and form of this boundary on the position of leaf initiation, leaf polarity and the resulting leaf form. While this was appreciated, there was also a strong feeling from several reviewers that much of this, beautiful as it is, was largely confirmatory. In addition, there were concerns that some manipulations had technical issues (notably the long treatment with cocktails of auxin drugs) or use of multiple types of meristems that made the results somewhat ambiguous.
Caggiano et al., address a long–standing issue of organ formation in plants. How do leaves acquire their characteristic arrangement around the shoot meristem and their flattened shape with different ad/abaxial domains? Is there a signal from the meristem, or are organs pre–patterned based on information in the meristem? Using detailed live–cell imaging and a number of genetic and experimental manipulations, the authors argue for pre–patterning in the meristem of ad/ab genes (primarily HD–ZIPs and Kanadi's) and for in instructive role of auxin in defining cell identity, potentially by influencing the expression domains of REV and KAN.
Overall, this work was interesting, the imaging of gene expression patterns was impressive and the overall view of pattern formation seems reasonable. I particularly like the extension of models from their previous SAM work to accommodate the new data provided here. Also, the phenotypes and diagrams explaining their potential origin in Figure 6 are quite cool! The authors point out alternative conclusions about D/V pattern that come from recently published work in tomato. I am afraid I don't have the background to evaluate the difference between these studies.
Organ polarity is influenced by the organ placement relative to the shoot apical meristem (SAM). Several hypotheses have been proposed with respect to the nature of the positional information that directs organ polarity specification. One model suggests that pre–patterning in the SAM peripheral zone specifies the dorsal and ventral (abaxial and adaxial) sides of the leaf. In the current manuscript, using beautiful imaging and side–specific markers, the authors clearly show the existence of polarity within the peripheral zone of the SAM. They further show that the site of organ initiation is determined by the location of this dorsal–ventral boundary within the SAM. Alteration of this boundary by manipulation of the expression domain of KANADI, which promotes ventral identity, influences the initiation site. They further analyze the effect of auxin treatment and wounding on the placement of the boundary.
The manuscripts presents convincing and high quality evidence for the presence of the boundary within the SAM and the influence of the location and form of this boundary on the position of leaf initiation, leaf polarity and the resulting leaf form. The question is whether these findings are mainly confirmatory or bring new biological insights. The dependence of organ initiation and blade outgrowth on dorsal–ventral polarity has been shown before. It also has been shown that there is preexisting polarity within the peripheral zone of the SAM, but this manuscript shows it more clearly, including the pre–existing polar expression of ventral and dorsal determinants. The causal effect of auxin on this boundary is less convincing, as it is hard to tell how a combined NPA and auxin treatment or a treatment of a combination of 3 auxin biosynthesis and response inhibitors affects the different aspects of the process. The effect of these treatments is shown after a relatively long time, and these effects on the localization of the examined markers could result from an effect on the domain patterning within the SAM rather than an effect on REV expression.
Caggiano et al., address the question, how the dorso–ventral (DV) pattern of leaves is established. Using fluorescent protein reporters they show that DV patterning of leaves correlates with central–peripheral patterning of the shoot apical meristem (SAM). The experimental data related to this paragraph are shown in 5 figures (Figures 1–5), of which many details are not addressed in the text. The novelty of this part is limited, because similar data have been published earlier (e.g. McConnell et al., 2001; reviewed by Barton et al., 2009).
Subsequently, the authors provide evidence that HD–ZIPIII and KAN genes repress leaf primordium initiation in the vegetative SAM. However, the question whether this is a direct influence on organ initiation or an indirect influence on SAM maintenance was not answered, because SAM organization in the arrested plants was not analyzed. Similar results were obtained by Kerstetter et al., 2001.
In the next paragraph, the consequences of ectopic KAN expression in the central domain of the SAM are studied. Most seedlings initiate several abnormal organs before their growth is arrested. The distinct organ phenotypes observed can be explained by the orientation of boundaries within organ founder cells.
Furthermore, the manuscript focuses on the interplay between auxin, HD–ZIPIII–expression and KAN–expression. Experiments using either NAA and NPA or a combination of auxinole, yuccasin and kyn led to the conclusion that high levels of auxin promote REV expression, whereas depletion of auxin promotes KAN expression. In this context, the treatment with a combination of three different drugs, which is necessary to see the described extension of KAN expression, may also have a different effect on gene expression than the one mediated through the observed depletion of auxin. The interplay of auxin, HD–ZIPIII expression and KAN–expression has also been addressed in previous studies (e.g. Izhaki and Bowman, 2007; Ilegems et al., 2010) with similar results.
In the last paragraph, the authors show that wounding in inflorescence meristems (IMs) leads to a depletion of auxin around the wound followed by an accumulation of KAN, which could be eliminated by an NAA + NPA treatment. Again, part of these findings were already known (Heisler et al., 2010; Landrein et al., 2015). In addition, IMs may react different from vegetative meristems.
Taken together, the manuscript presents a nice set of experiments supporting the view that DV organization of leaves is pre–patterned by the expression domains of REV and KAN in the SAM. However, previous studies revealed many similar results. Therefore, the study by Caggiano et al., presents only an incremental advance in our understanding.
Aiming for an explanation of the origin of DV patterning of leaves, it seems problematic to mix results from experiments using vegetative meristems and using inflorescence meristems, because a flower primordium is not a structure showing a DV pattern like a leaf primordium.
The wounding experiments offer an alternative explanation for the observations made by I. Sussex, but we think they do not rule out the existence of a signal from the main meristem to the developing leaf primordium.
[Editors’ note: what now follows is the decision letter after the authors submitted for further consideration.]
Thank you for resubmitting your work entitled "Cell type boundaries organize plant development" for further consideration at eLife. Your revised article has been favorably evaluated by Christian Hardtke (Senior editor) and three reviewers (mix of new and previous), one of whom is a member of our Board of Reviewing Editors.
There was, as in the first version, considerable appreciation of the imaging quality and evidence for a prepattern mechanism. There was also concern that the role of auxin in the process was not strongly supported by the provided data. Specifically, there was continued concern about the length of auxin treatments and the potential for KAN and REV responses to be secondary. Additionally, because the highest levels of auxin–signaling activity are found in the boundary region, but REV is precisely not expressed there, it was unclear what REV induction there by NPA+NAA treatment means. Moreover, how would this observation relate to a potential auxin–mediated maintenance of REV expression in the central region of the meristem, where there appears to be very little auxin–signaling activity?
After considerable deliberation, we have decided to consider a revised manuscript, IF it can address this critical concern about the proposed mechanism of auxin–mediated pre–patterning via REV activation and KAN repression. The misexpression of boundary genes (reviewer 1) could be one way to add more evidence, but alternative experiments would also be acceptable.
A revised version should also take care to tone down the discussion related to the "Sussex signal" because those experiments were done in floral meristems where the D/V polarity is limited. Ideally the issue would be addressed experimentally (e.g. using tomato apices), but the authors could also consider simply rewriting this section.
In this revision, the authors have addressed my concerns about data presentation and have clarified many of the confusing passages of the previous version.
In this current version, I see that they have generated a very high resolution description of organ boundary gene expression and have a plausible model for auxin (though like most auxin models, there are enough feedbacks that it’s hard to find a way to experimentally disprove it). To my mind a manipulation in which they misexpress boundary genes in such a way that polarity becomes locally reoriented (something like making fly wing clones that create artificial boundaries) would be the real test of their models. It seems to me that reagents to do these experiments should be available, and I would be very supportive of this manuscript if such data could be added.
In the revised manuscript, Caggiano et al., address some of the concerns raised by the reviewers and explain and clarify the new findings presented in this paper. As before, the imaging is impressive and convincing with respect to the pre–patterning of in the polarity in the SAM and the causative role of the boundary in organ initiation.
The authors better clarify the reason for the use of multiple drugs to assess the role of auxin in the patterning. However, I still think that the presented data on the effect of auxin does not support the interpretation on the role of auxin in specifying the spatial patterning in the SAM periphery. The time from application to effect is still long, and the effect on KAN and REV may be secondary to other effects. Combined with the relatively broad application, the results in my opinion do not support the proposed mechanism of auxin–mediated pre–patterning via REV activation and KAN repression.
This manuscript addresses a long–standing question in plant biology, the acquisition of dorso–ventral (DV; or adaxial/abaxial) polarity in plant leaves. The authors use fluorescence live imaging combined with transgenic manipulation of gene–expression domains and pharmacological treatments to characterize the role of the boundary region between dorsal and ventral gene–expression domains in the meristem in leaf–polarity acquisition. They confirm and provide further detail on the pre–pattern in the shoot meristem involving centrally expressed HD–ZIPIII and peripherally expressed KANADI1 genes, and show that the boundary between the two domains, where neither gene is expressed strongly, is the site of auxin convergence driven by PIN1 and the highest auxin–signaling activity. Overexpressing either REV or KAN1 throughout the meristem epidermis abolishes organ initiation, consistent with their documented role in suppressing auxin activity. Together this suggests that the auxin maxima can only form in the boundary domain to trigger organ outgrowth there. Forcing KAN1 expression in the meristem centre and thus creating a new DV boundary there can lead to leaf radialization or even polarity reversal, indicating that the pre–pattern in the meristem can have an instructive function for leaf DV polarity. Having demonstrated that the pattern of REV and KAN1 expression determines the pattern of auxin responsiveness in the meristem, the authors then ask whether auxin in turn influences the expression pattern of these genes. Treating meristems with NAA and NPA causes an increase in the REV expression domain into the boundary region and can prevent the re–establishment of KAN1 expression in the periphery separating an outgrowing primordium from the meristem; conversely, inhibiting auxin signaling leads to an encroachment of KAN1 expression towards the meristem centre. Lastly, the authors show that the previously demonstrated auxin depletion around wound sites leads to an upregulation of KAN1 expression around the wound. This latter observation is offered as an explanation for the leaf radialization seen after separating a leaf primordium from the meristem in Sussex' classical experiments.
How DV polarity in leaves is determined is a question of fundamental importance and broad interest in plant development. As the ongoing interest in the quest for the proposed Sussex signal indicates, the work presented is clearly highly timely. The manuscript provides a mix of rather confirmatory and novel results. Important novel aspects are the demonstration that disturbing the DV expression pre–pattern in the meristem can lead to leaf–polarity defects; the finding that auxin in turn can modulate this DV expression pre–pattern; and the observation that peripheral/abaxial identity is induced surrounding wounds in the meristem.
I have the following comments:
1) The finding of a likely causal role of the pre–pattern on leaf polarity is an important advance. Also, the observation that auxin can modulate the DV expression pattern is interesting, yet its biological significance appears somewhat unclear. As the authors show in Figure 1 and in their model in Figure 9, the highest levels of auxin–signaling activity are found in the boundary region. However, it is precisely there that REV is not expressed. Thus, what does it mean that REV can be induced there by NPA+NAA treatment? And how does this observation relate to a potential auxin–mediated maintenance of REV expression in the central region of the meristem, where there appears to be very little auxin–signaling activity? Have the authors attempted to incorporate the auxin effect on REV expression into their model? Does this provide any information about the possible significance of this interaction?
2) The alternative explanation offered for the effects of wounds separating a primordium from the meristem is novel, interesting and potentially of broad significance. However, its impact appears limited, as the KAN1 upregulation around the entire wound is currently only shown clearly for Arabidopsis inflorescence meristems (while the seedling meristems undergo more dramatic re–organization, making their behavior more difficult to interpret), where the action of a Sussex signal is hard to demonstrate due to the limited DV polarity of floral meristems. This means that the alternative explanation is not backed up by functional experiments. Ideally, the authors would directly test in a different system, such as the tomato apex, whether NAA+NPA treatment after appropriate wounding could prevent organ abaxialization.https://doi.org/10.7554/eLife.27421.030
- Marcus G Heisler
- Pia Sappl
- Elliot M Meyerowitz
- Henrik Jönsson
- Henrik Jönsson
The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.
We thank M E Byrne for helpful feedback and ideas on the manuscript. We thank J Bowman for helpful discussions as well as for plasmids containing REVr and PHVr cDNAs. We thank D Weijers and C Y Liao for a plasmid containing pDR5v2::ntdTomato; K Nakajima for the T-DNA construct of miR166Ap::GFPER, Dr Atsushi Miyawaki from RIKEN Brain Science Institute for the VENUS fluorescent protein which was obtained through an MTA. We thank Professor Ken-ichiro Hayashi, Okayama University of Science for providing Auxinole. The EMM laboratory is supported by funds from the Howard Hughes Medical Institute and the Gordon and Betty Moore Foundation (through grant GBMF3406). The research leading to these results received funding from the Australian Research Council (MGH) and European Research Council under the European Union’s Seventh Framework Programme (FP/2007–2013)/ERC Grant Agreement n. 261081 (MGH), as well as the People Programme (Marie Curie Actions) under REA grant agreement n. 255089 (PS). The work was also supported by: the European Molecular Biology Laboratory (XY, MPC, CO, PS, NB, HR and MGH); the EMBL International PhD Programme (XY, NB and MPC); Gatsby Charitable Foundation (GAT3395/PR4) (HJ) and Swedish Research Council (VR2013-4632) (HJ). The authors declare no competing financial interests. Supplement contains additional data and movie. We declare there are no competing interests.
- Dominique C Bergmann, Reviewing Editor, Stanford University/HHMI, United States
© 2017, Caggiano et al.
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