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Progress in understanding the role of auxin in lateral organ development in plants
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ScienceDirect Progress in understanding the role of auxin in lateral organ development in plants Marcus G Heisler and Mary E Byrne Plants continuously produce lateral organs from the shoot apex such as leaves and flowers, providing an excellent opportunity to study their development. The plant hormone auxin plays a central role in this process by promoting organ formation where it accumulates due to polar auxin transport. Recently, the use of live-imaging, fine perturbation techniques and computational modelling has helped researchers make exciting progress in addressing long-standing questions on plant organogenesis, not only regarding the role of auxin in promoting growth but also on the regulation of morphogenesis and transcriptional control. In this review, we discuss a number of recent studies that address these points, with particular reference to how auxin acts in early leaf development and in leaf shape. Address School of Life and Environmental Sciences, University of Sydney, NSW 2006, Australia Corresponding authors: Heisler, Marcus G ([email protected]), Byrne, Mary E ([email protected])
Current Opinion in Plant Biology 2020, 53:73–79 This review comes from a themed issue on Growth and development Edited by Marcus Heisler and Alexis Maizel
https://doi.org/10.1016/j.pbi.2019.10.007 1369-5266/ã 2019 Published by Elsevier Ltd.
Introduction Leaves and flowers initiate on the flanks of the shoot meristem, with successive leaves arising in the position of an auxin maximum [1,2]. In turn, the localized maxima are positioned by the auxin efflux carrier PIN1, which directs auxin towards initiation sites according to a feedback loop between auxin signalling and PIN1 polarity [1,3]. However, while auxin can induce organ growth on the flanks of the meristem, tissues at the apex or further down the flank are nonresponsive [2]. What defines the auxin responsive peripheral zone? What does auxin actually do to promote tissue outgrowth and how is the shape of the resulting organ controlled? Below we discuss several recent studies that address these questions. www.sciencedirect.com
Adaxial-abaxial patterning and organ positioning: several models Leaves initiate in the peripheral region of the shoot apical meristem (SAM) and establish along three axes: an adaxialabaxial (top to bottom), proximal-distal (base to tip) and mediolateral (middle to margin). The adaxial-abaxial axis forms two opposing sides of the leaf primordium, which differentiate into distinct tissues and both tissues are required for lamina development [4–6]. The tissues closest to the shoot axis are called adaxial while the tissues further away are called abaxial. Three competing models have been proposed to explain how leaf adaxial/abaxial polarity arises or is potentially maintained (Figure 1). In potato and tomato, wounding experiments that physically separate the initiating leaf primordium from the meristem develop as radial abaxial leaves. This has been suggested as evidence that leaf adaxial fate and therefore adaxial-abaxial polarity, requires a signal, often called the Sussex signal, from the meristem to leaf (Figure 1a) [7,8]. A second model proposes that leaves are pre-patterned according to tissue types in the SAM where leaves originate (Figure 1b). Supporting this, the expression of polarity defining transcription factors in the SAM, such as REVOLUTA (REV) and KANADI1 (KAN1), form concentric non-overlapping domains within the SAM that form a pre-pattern with respect to initiating organs [9,10]. Leaf primordia are centered specifically on the boundary region of these gene expression patterns such that adjacent adaxial and abaxial cell types are also incorporated into the primordia, thereby establishing tissue polarity and the propagation of this SAM peripheral region boundary into the developing organ. What limits leaf primordia initiation to this boundary? Manipulation of REV and KAN1 expression within the SAM indicates that these transcription factors repress auxin-dependent organogenesis; hence, only cells in between their expression domains, that is, at the boundary, are capable of responding to auxin, for instance by expressing boundary genes such as PRESSED FLOWER (PRS) and WUSCHEL RELATED HOMEOBOX1 (WOX1) [9]. The third model proposes that flow of auxin from the initiating leaf primordium to the meristem creates a region of low auxin that promotes adaxial fate (Figure 1c) [11]. This model is largely based on the finding that application of auxin to the adaxial side of leaf primordia in tomato results in leaves that are abaxialized and that auxin levels appear to be lower in adaxial tissues, as judged using the DII auxin sensor [11]. Extending this idea, the authors propose that auxinresponsive genes expressed in the middle domain of the leaf such as WOX1 and PRS are not expressed adaxially because of a lack of auxin while in abaxial tissues they are Current Opinion in Plant Biology 2020, 53:73–79
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Figure 1
(a)
Mobile signal from meristem centre that promotes adaxial identity Direction of movement
(b)
HD-ZIPIII expression Boundary region in between HD-ZIPIII and KAN domains KANADI gene expression
(c)
High auxin levels Low auxin levels Direction of auxin transport
Initiating leaf primordium Current Opinion in Plant Biology
Alternative hypotheses for explaining the establishment of adaxial-abaxial patterning in developing leaf primordia. (a) Model, in which a morphogen is produced by the meristem that promotes adaxial cell identity, thereby conferring polarity to newly developing leaf primordia [8]. (b) Pre-pattern model, in which organ position along the radial axis of the shoot is controlled by the same transcription factors (HD-ZIPIII and KANADI) that specify adaxial and abaxial tissue identity. These factors make sure organs arise at the boundary between their expression domains such that adaxial-abaxial patterning is transferred from meristem to leaf primordium [9]. (c) Auxin depletion model, in which, due to PIN1 mediated auxin transport, auxin is depleted in adaxial cells of new leaf primordia. This low level of auxin helps maintain or establish adaxial identity [11].
actively repressed by repressor-type Auxin Response Factors (ARFs) [12]. Hence, this model contrasts with the second model in terms of regulation of WOX1 and PRS in adaxial tissues (Figure 2). How does support for these competing models compare? An important piece of evidence supporting the prepattern model is that experimental alteration of this pre-pattern, solely in the meristem, is sufficient to cause major changes in leaf polarity, demonstrating its functional significance [9]. Furthermore, the abaxialization response to wounds, which inspired the meristem derived signal model [7] can also be explained by the meristem pre-pattern model since at least in Arabidopsis, wounding within the meristem results in auxin depletion [13] and this promotes ectopic KAN1 expression around the wound at the expense of REV, indicating abaxialization of the wounded tissue [9]. Supporting a causal role of auxin depletion, exogenous auxin treatment at the time of wounding can inhibit such a response [9]. However, Current Opinion in Plant Biology 2020, 53:73–79
although wounding Arabidopsis meristem tissues results in changes to adaxial and abaxial gene expressions, loss of leaf polarity was not observed. Instead, the Arabidopsis meristem bifurcates into two or more meristems and the leaves that form subsequently are normal. Hence an important task is to repeat the wounding experiments in potato and tomato and to test whether auxin can prevent polarity disruptions. What about the auxin depletion model, which also proposes that low auxin levels restrict WOX1 and PRS expression? This model is not necessarily mutually exclusive to the meristem pre-pattern model if the auxin depletion mechanism is required only for polarity maintenance after leaf initiation. However, beyond that, there are several inconsistencies between the auxin depletion and pre-pattern models. One major inconsistency is that auxin promotes HD-ZIPIII expression and restricts KAN1 expression [9], which is opposite to what one would expect if high auxin levels disrupt adaxial www.sciencedirect.com
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Figure 2
auxin
Absence of auxin (depletion model) or HD-ZIPIII expression (pre-pattern model)
Adaxial
PRS WOX1
Leaf margin
KAN ARF3/4
Abaxial Current Opinion in Plant Biology
Implications of alternative adaxial-abaxial hypotheses for the regulation of PRS and WOX1. Both the pre-pattern and auxin depletion models support a role for KANADI and ARF3/4 transcription factors in repressing PRS and WOX1 in abaxial leaf tissues. However the auxin depletion model proposes that it is a lack of auxin in adaxial tissues that prevents adaxial PRS and WOX1 expression while the pre-pattern model proposes that WOX1 and PRS are repressed adaxially directly or indirectly by the HD-ZIPIII proteins.
identity. A second inconsistency is that if auxin is applied exogenously to leaf primordia, WOX1 and PRS expression does not extend into adaxial leaf tissues, indicating that a lack of auxin is not restricting their expression domains [9] as has been proposed [12] (Figure 2). Finally, the finding that auxin levels are lower in adaxial leaf tissues has also come under dispute [14,15]. Despite these discrepancies however, no other model can explain why tomato leaf primordia become abaxialized upon adaxial auxin application. Do tomato leaves develop differently to Arabidopsis? Ultimately additional experiments in tomato may be needed that monitor auxin levels and adaxial and abaxial tissue identity in response to global and localized auxin manipulations. During the early stages of leaf growth the adaxial/middle (boundary)/abaxial pattern present within the founder cell population must be maintained during leaf development. A recent study has revealed how small RNAs act in a surprising way to help achieve this. Regulation of adaxial HD-ZIPIII and abaxial ARF genes is accomplished by small RNAs mir165/166 and ta-siARF, respectively [16] and since these small RNAs are known to move cell to cell from their sites of transcription in the adaxial or abaxial epidermis [16], they are anticipated to form a gradient across the leaf. In contrast, the defined expression of target genes to specific mesophyll cell layers suggest a response to threshold levels of these regulatory small RNA [17]. How might miRNA gradients be established? Although it remains to be established whether or not miRNA gradients are functional in early organ patterning, within the SAM, a miRNA and corresponding target reporter system has shown discrete domains of directional miRNA movement that is distinct from that of protein cell-to-cell movement [18]. miRNA distribution in the shoot meristem may include movement from sites of synthesis through plasmodesmata www.sciencedirect.com
as in the root where callous deposition and plasmamembrane plasmodesmata-associated receptor-like kinases BAM1 and BAM2 effect the spatial distribution of mir165/166 [19–21]. In the context of the three models proposed for establishing leaf adaxial/abaxial polarity, small RNAs may represent the Sussex signal and be sufficient for early patterning of the leaf. Alternatively, small RNAs may reinforce or act together with the other proposed mechanisms defining leaf adaxial/abaxial polarity.
Downstream of auxin Which genes apart from WOX1 and PRS are known to act downstream of auxin to promote organ growth? Several transcription factors are induced by auxin at primordial positions via Auxin Response Factor 5 (ARF5)/MONOPTEROS (MP) and function to promote flower development, through induction of genes including LEAFY, AINTEGUMENTA, AINTEGUMENTA-LIKE6 and FILAMENTOUS FLOWER (FIL) [22,23]. More recently, the LEAFLESS (LFS) gene from tomato, encoding a gene closely related to the Arabidopsis DORNRONSCHEN (DRN) and DRN-like (DRNL) genes, was found to be essential for leaf formation [24]. LFS is induced by auxin and is expressed at sites of auxin maxima in the shoot meristem. Loss of LFS results in a pin-like shoot where the shoot meristem fails to initiate organs-resembling shoot apices treated with auxin transport inhibitors. Like mp mutants, exogenous application of auxin is not sufficient to induce organ outgrowth from lfs apices [1,24]. As double mutants of DRN and DRNL in Arabidopsis show a similar phenotype to lfl in tomato, these results indicate LFL/DRN/ DRNL act as an essential transducer of the auxin signal leading to organ formation [24]. Another important transcriptional event downstream of auxin is downregulation of SHOOT MERISTEMLESS (STM). Recently it was Current Opinion in Plant Biology 2020, 53:73–79
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found that MP acts indirectly to achieve this by activating FIL. FIL forms a protein complex with ETT and ARF4, which directly represses STM, as well as the closely related BREVIPEDICELLUS (BP), through recruitment of the histone deactylase HDAC19 and subsequent transcriptional silencing [25].
Figure 3
Organ outgrowth
Leaf morphogenesis So far, we have discussed recent work focused on genetic regulatory networks which pattern cell types and regulate and are regulated by auxin. But how does auxin function at a cellular level to promote organ outgrowth and how is the shape of new organs determined? Recent work has provided important new insights into both these questions. Previous work has shown that auxin acts to promote organ formation at least in part through the local de-polymerization of microtubules. Since interphase microtubule arrangements help orient cellulose deposition in the cell wall, which helps orient cellular growth, this finding implied that a loss of growth anisotropy is sufficient to initiate organogenesis [26]. Is growth anisotropy the only relevant process triggered by microtubule de-polymerization? Using computer simulations, a more recent study concluded this cannot be the case [27]. Rather than increasing growth rates, as has been measured to occur at sites of organogenesis, computer simulations of growth using a mechanical model of the shoot meristem showed that local reductions in cell wall anisotropy should result in a decreased growth rate. This suggested that in addition to cell wall anisotropy, microtubule depolymerization likely influences other cell wall properties that increase growth rates. To investigate further, the authors documented the expression patterns of many genes encoding cell wall modifying enzymes and found several to be expression in the SAM and floral organ primordia. Interestingly, several of these genes were also upregulated in the organ-like outgrowths formed from the apex of double mutants of PIN1 and BOTERO1/KATANIN, encoding a microtubule associated protein, suggesting that auxin build up may not be required for organogenesis. This hypothesis was further supported by an absence of DR5 expression in organs induced by oryzalin application to NPA-induced ‘pin’ meristems. Lastly, since it had been shown that local application of cell wall modifying enzymes such as pectin methylesterase to NPA-pins can also induce outgrowths [28], the authors tested whether such treatments also induced a loss of microtubule anisotropy and found that they do, although the response was slower compared to auxin treatments [27]. Altogether these results reveal that several distinct processes are coupled during organ formation. High auxin concentrations result in a loss of microtubule array anisotropy and this is accompanied by the local upregulation of cell wall modifying enzymes. The latter two processes positively regulate each other and promoting either can trigger organ outgrowth (Figure 3). Whether the loss of microtubule anisotropy Current Opinion in Plant Biology 2020, 53:73–79
Expression of cell wall modifying enzymes
Microtubule depolymerisation
Auxin Current Opinion in Plant Biology
The role of feedback loops between the activity of cell wall modifying enzymes and microtubule depolymerization in mediating auxininduced organ formation. Feedback between microtubule depolymerization and the expression of cell wall enzymes is indicated by the solid arrows. Although auxin promotes both processes, it is not yet clear whether auxin promotes both independently or not (dashed arrows). It also remains unclear whether both microtubule depolymerization and cell wall enzyme expression promote organ outgrowth independently or not (dashed arrows).
or the induction of cell wall enzymes is sufficient to trigger outgrowth in the absence of the other remains to be determined (although simulations suggest loss of anisotropy is not sufficient), as do the molecular mechanisms through which this mutual regulation occurs. Given that organ initiation is accompanied by a loss of mechanical anisotropy, how is organ shape determined? The available data suggest that the loss of anisotropy only occurs during the initial outgrowth process and is restricted to the central point of auxin maximum [26]. Microtubule orientations otherwise are generally observed to be circumferentially oriented on the surface of organ primordia, consistent with the known role of mechanical stress in regulating their orientation [29]. These microtubule orientations would be expected to promote growth along the proximo-distal axis. Cell polarity, as marked by PIN1, also correlates with microtubule orientations linking polarity to growth direction and supporting the proposal that mechanical stresses also regulate cell polarity patterns [30]. Consistent with this scenario, a two-dimensional ‘polarity field’, (although in this case polarity is suggested to be patterned by a hypothetical diffusing signal), has been proposed to regulate lamina growth for the simple leaves of Arabidopsis [31]. This polarity field was proposed to promote www.sciencedirect.com
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growth both parallel and perpendicular to the field axis depending on position within the primordium, which in turn caused deformations in the field itself. This meant that although the field was initially oriented proximo-distally, it later diverged more laterally [31]. The observed growth directions were found to be consistent with this proposal, being mainly oriented proximo-distally early in development but later diverging towards the margin and becoming more isotropic [31]. Evidence that leaf cells are indeed polarized in a corresponding pattern has now been reported, based on the cellular localization patterns of the BASL protein. BASL is a novel protein that is asymmetrically localized in stomatal epidermal cell linages of the leaf [32]. Using an inducible GFP-BASL in a mutant background that lacks stomata, ectopic BASL expresses asymmetrically all epidermal cells of the leaf thus marking leaf polarity fields [33]. At early stages of leaf development, BASL polarity patterns exhibit a distal to proximal pattern along the same axis as distally oriented PIN1 but at late stages, it’s pattern diverges laterally as predicted [33]. However, further evidence that this polarity pattern is regulated by mechanics rather than a diffusible factor comes from the finding that BRXL2, a protein with asymmetric localization similar to BASL in the leaf, re-orients its polarity in response to mechanical perturbations [34]. The mechanism by which mechanical stress orients polarity over such large distances however remains speculative [33,35]. Given that PIN1 polarities and interphase microtubule array orientations are known to correlate in the SAM during organ initiation [30] and BASL also correlates with PIN1 polarity [33], it would seem likely that ectopic BASL polarity patterns are marking microtubule orientations during leaf development. A recent pre-print however indicates that while this is likely true early in leaf development, later in leaf development, as the leaf blade broadens, microtubule orientations become isotropic at the leaf surface, somewhat consistent with observed growth patterns but inconsistent with BASL polarity patterns [36]. The authors also find that while mechanical stress patterns can account for the observed microtubule orientations early on, the switch to a more isotropic pattern does not fit their model predictions. The authors conclude that at later developmental stages microtubules must become decoupled from stress patterns (although previous laser ablation experiments indicate microtubules do remain stress-responsive at least until pavement cells have formed [37]). In accordance with their modelling, they also predict that the switch to isotropy promotes leaf blade expansion [36]. All together these results indicate that at later stages of leaf development when the tissue differentiates, microtubule orientations and cell polarity patterns may become uncoupled and this may be important for leaf blade expansion. How do leaves become flat from a more symmetrical starting point? The same preprint mentioned above also investigates microtubule and cellulose patterns in the internal cell layers of the leaf. It reports that during both www.sciencedirect.com
early and late development, microtubules and cellulose fibrils within the anticlinal walls of leaf cells are oriented anticlinally, that is, perpendicular to the leaf surface and that according to model predictions, these orientations promote leaf flatness and follow the predicted stress patterns for the tissue (as mircrotubules are known to do elsewhere) [36]. How the primordium initially becomes asymmetric remains unclear. Although mechanical differences in wall stiffness on different sides of the leaf and have been proposed to account for initial leaf flattening [38], this hypothesis is under dispute [39,40]. Finally, what about complex leaves? Arabidopsis thaliana has simple leaves with an undivided leaf lamina and small serrations, or outgrowths, along the leaf margin. Leaf shape varies considerably in different plant species and can be simple, as in A. thaliana or may be complex, where the leaf lamina is divided into leaflets, as in Cardamine hirsute a close relative of A. thaliana. A feedback regulatory network involving auxin and the transcription factor CUC2 act along the leaf margin to control the formation of serrations in A. thaliana [41]. Furthermore, both auxin and CUC2 are required for development of distinct leaflets in compound leaves [42,43]. It has been less clear how other regulators of leaf shape are involved in compound leaf development. A recent study combining quantitative live-imaging, computational modelling and genetics provides a detailed look at how growth patterns are modified to go from simple leaves to complex [44]. The authors start by quantitatively comparing leaf growth patterns in A. thaliana and C. hirsute and with modelling and genetic manipulation find that the differences in morphology can largely be explained by the action of two homeobox genes, SHOOT MERISTEMLESS (STM) and REDUCED COMPLEXITY (RCO). In C. hirsute, the expression of STM (which is absent in A. thaliana leaves) acts broadly to slow growth but also extends the period of time, in which the leaf primordium remains undifferentiated. This is especially relevant to the distal and marginal part of the primordium where STM prolongs the growth phase in Cardamine and enables leaflets to form respectively [44]. In contrast, RCO acts locally to inhibit growth and is normally expressed at the base of Cardamine leaflets. By expressing both STM and RCO in A. thaliana leaves, the authors were able to create complex leaves in Arabidopsis that resembled those from Cardamine, thereby demonstrating how key genes enable morphological diversity [44].
Conclusion It is an exciting time for understanding plant organ development. The work discussed here represents significant progress in addressing questions that have been highlighted in the field for decades, facilitated by the use of quantitative live-imaging and modelling. Nevertheless, several findings remain under dispute and it will be important to resolve these disagreements Current Opinion in Plant Biology 2020, 53:73–79
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in order to make further progress and achieve a cohesive understanding across the field. In addition to the studies cited here, there have been many other important contributions to the literature on plant organ development recently and we regret that due to space constraints our review could not be exhaustive.
Conflict of interest statement Nothing declared.
Acknowledgement Funding is gratefully acknowledged by MH from the Australian Research Council (DP180101149).
References and recommended reading Papers of particular interest, published within the period of review, have been highlighted as: of special interest of outstanding interest 1.
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coupled to microtubule-driven growth isotropy at the shoot apex in Arabidopsis. Development 2018, 145. In this paper a feedback loop between the expression of cell remodelling genes and microtubule-depolymerization is discovered. 28. Peaucelle A, Braybrook SA, Le Guillou L, Bron E, Kuhlemeier C, Hofte H: Pectin-induced changes in cell wall mechanics underlie organ initiation in Arabidopsis. Curr Biol 2011, 21:1720-1726. 29. Hamant O, Heisler MG, Jonsson H, Krupinski P, Uyttewaal M, Bokov P, Corson F, Sahlin P, Boudaoud A, Meyerowitz EM et al.: Developmental patterning by mechanical signals in Arabidopsis. Science 2008, 322:1650-1655. 30. Heisler MG, Hamant O, Krupinski P, Uyttewaal M, Ohno C, Jonsson H, Traas J, Meyerowitz EM: Alignment between PIN1 polarity and microtubule orientation in the shoot apical meristem reveals a tight coupling between morphogenesis and auxin transport. PLoS Biol 2010, 8:e1000516. 31. Kuchen EE, Fox S, de Reuille PB, Kennaway R, Bensmihen S, Avondo J, Calder GM, Southam P, Robinson S, Bangham A et al.: Generation of leaf shape through early patterns of growth and tissue polarity. Science 2012, 335:1092-1096. 32. Dong J, MacAlister CA, Bergmann DC: BASL controls asymmetric cell division in Arabidopsis. Cell 2009, 137:1320-1330. 33. Mansfield C, Newman JL, Olsson TSG, Hartley M, Chan J, Coen E: Ectopic BASL reveals tissue cell polarity throughout leaf development in Arabidopsis thaliana. Curr Biol 2018, 28:26382646 e2634. This study reveals that cells remain polarized in the epidermis of leaves until late developmental stages. The polarity pattern resembles the pattern predicted through modelling in an earlier study. 34. Bringmann M, Bergmann DC: Tissue-wide mechanical forces influence the polarity of stomatal stem cells in Arabidopsis. Curr Biol 2017, 27:877-883. 35. Bhatia N, Heisler MG: Self-organizing periodicity in development: organ positioning in plants. Development 2018, 145. This review summarizes our current understanding of how cell polarity patterns are regulated in relation to phyllotaxis. 36. Zhao F, Du F, Oliveri H, Zhou L, Ali O, Chen W, Feng S, Wang Q, Lu¨ S, Long M et al.: A microtubule-mediated mechanical feedback controls leaf blade development in three dimensions. bioRxiv 2019:604710. This pre-print reveals patterns of microtubule orientations during leaf development and shows that some observed patterns match predictions
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based on mechanical stresses while others do not. The authors also propose that a loss of anisotropy in the epidermis during leaf development promotes leaf flattening. 37. Sampathkumar A, Krupinski P, Wightman R, Milani P, Berquand A, Boudaoud A, Hamant O, Jonsson H, Meyerowitz EM: Subcellular and supracellular mechanical stress prescribes cytoskeleton behavior in Arabidopsis cotyledon pavement cells. eLife 2014, 3:e01967. 38. Qi J, Wu B, Feng S, Lu S, Guan C, Zhang X, Qiu D, Hu Y, Zhou Y, Li C et al.: Mechanical regulation of organ asymmetry in leaves. Nat Plants 2017, 3:724-733. This study measures the mechanical properties of adaxial and abaxial leaf tissues using Atomic Force Microscopy. Using modelling and measurements, the authors propose that the differences they observe can account for leaf flattening. 39. Feng S, Zhou L, Lu S, Long M, Jiao Y: Reply to ‘Early shaping of a leaf’. Nat Plants 2018, 4:620-621. This is a reply to Ref. [40]. 40. Coen E, Kennaway R: Early shaping of a leaf. Nat Plants 2018, 4:618-619. In this commentary, the authors raise important issues with a previously proposed mechanical model used to simulate leaf growth (see Ref. [38]). 41. Bilsborough GD, Runions A, Barkoulas M, Jenkins HW, Hasson A, Galinha C, Laufs P, Hay A, Prusinkiewicz P, Tsiantis M: Model for the regulation of Arabidopsis thaliana leaf margin development. Proc Natl Acad Sci U S A 2011, 108:3424-3429. 42. Barkoulas M, Hay A, Kougioumoutzi E, Tsiantis M: A developmental framework for dissected leaf formation in the Arabidopsis relative Cardamine hirsuta. Nat Genet 2008, 40:1136-1141. 43. Blein T, Pulido A, Vialette-Guiraud A, Nikovics K, Morin H, Hay A, Johansen IE, Tsiantis M, Laufs P: A conserved molecular framework for compound leaf development. Science 2008, 322:1835-1839. 44. Kierzkowski D, Runions A, Vuolo F, Strauss S, Lymbouridou R, Routier-Kierzkowska AL, Wilson-Sanchez D, Jenke H, Galinha C, Mosca G et al.: A Growth-based framework for leaf shape development and diversity. Cell 2019, 177:1405-1418 e1417. This paper provides a dynamic and quantitative analysis of leaf growth, focusing on the influence of different genes in regulating leaf morphology. By manipulating the expression of two genes, the authors demonstrate the ability to create Cardamine-like complex leaf morphologies in Arabidopsis.
Current Opinion in Plant Biology 2020, 53:73–79
ScienceDirect Progress in understanding the role of auxin in lateral organ development in plants Marcus G Heisler and Mary E Byrne Plants continuously produce lateral organs from the shoot apex such as leaves and flowers, providing an excellent opportunity to study their development. The plant hormone auxin plays a central role in this process by promoting organ formation where it accumulates due to polar auxin transport. Recently, the use of live-imaging, fine perturbation techniques and computational modelling has helped researchers make exciting progress in addressing long-standing questions on plant organogenesis, not only regarding the role of auxin in promoting growth but also on the regulation of morphogenesis and transcriptional control. In this review, we discuss a number of recent studies that address these points, with particular reference to how auxin acts in early leaf development and in leaf shape. Address School of Life and Environmental Sciences, University of Sydney, NSW 2006, Australia Corresponding authors: Heisler, Marcus G ([email protected]), Byrne, Mary E ([email protected])
Current Opinion in Plant Biology 2020, 53:73–79 This review comes from a themed issue on Growth and development Edited by Marcus Heisler and Alexis Maizel
https://doi.org/10.1016/j.pbi.2019.10.007 1369-5266/ã 2019 Published by Elsevier Ltd.
Introduction Leaves and flowers initiate on the flanks of the shoot meristem, with successive leaves arising in the position of an auxin maximum [1,2]. In turn, the localized maxima are positioned by the auxin efflux carrier PIN1, which directs auxin towards initiation sites according to a feedback loop between auxin signalling and PIN1 polarity [1,3]. However, while auxin can induce organ growth on the flanks of the meristem, tissues at the apex or further down the flank are nonresponsive [2]. What defines the auxin responsive peripheral zone? What does auxin actually do to promote tissue outgrowth and how is the shape of the resulting organ controlled? Below we discuss several recent studies that address these questions. www.sciencedirect.com
Adaxial-abaxial patterning and organ positioning: several models Leaves initiate in the peripheral region of the shoot apical meristem (SAM) and establish along three axes: an adaxialabaxial (top to bottom), proximal-distal (base to tip) and mediolateral (middle to margin). The adaxial-abaxial axis forms two opposing sides of the leaf primordium, which differentiate into distinct tissues and both tissues are required for lamina development [4–6]. The tissues closest to the shoot axis are called adaxial while the tissues further away are called abaxial. Three competing models have been proposed to explain how leaf adaxial/abaxial polarity arises or is potentially maintained (Figure 1). In potato and tomato, wounding experiments that physically separate the initiating leaf primordium from the meristem develop as radial abaxial leaves. This has been suggested as evidence that leaf adaxial fate and therefore adaxial-abaxial polarity, requires a signal, often called the Sussex signal, from the meristem to leaf (Figure 1a) [7,8]. A second model proposes that leaves are pre-patterned according to tissue types in the SAM where leaves originate (Figure 1b). Supporting this, the expression of polarity defining transcription factors in the SAM, such as REVOLUTA (REV) and KANADI1 (KAN1), form concentric non-overlapping domains within the SAM that form a pre-pattern with respect to initiating organs [9,10]. Leaf primordia are centered specifically on the boundary region of these gene expression patterns such that adjacent adaxial and abaxial cell types are also incorporated into the primordia, thereby establishing tissue polarity and the propagation of this SAM peripheral region boundary into the developing organ. What limits leaf primordia initiation to this boundary? Manipulation of REV and KAN1 expression within the SAM indicates that these transcription factors repress auxin-dependent organogenesis; hence, only cells in between their expression domains, that is, at the boundary, are capable of responding to auxin, for instance by expressing boundary genes such as PRESSED FLOWER (PRS) and WUSCHEL RELATED HOMEOBOX1 (WOX1) [9]. The third model proposes that flow of auxin from the initiating leaf primordium to the meristem creates a region of low auxin that promotes adaxial fate (Figure 1c) [11]. This model is largely based on the finding that application of auxin to the adaxial side of leaf primordia in tomato results in leaves that are abaxialized and that auxin levels appear to be lower in adaxial tissues, as judged using the DII auxin sensor [11]. Extending this idea, the authors propose that auxinresponsive genes expressed in the middle domain of the leaf such as WOX1 and PRS are not expressed adaxially because of a lack of auxin while in abaxial tissues they are Current Opinion in Plant Biology 2020, 53:73–79
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Figure 1
(a)
Mobile signal from meristem centre that promotes adaxial identity Direction of movement
(b)
HD-ZIPIII expression Boundary region in between HD-ZIPIII and KAN domains KANADI gene expression
(c)
High auxin levels Low auxin levels Direction of auxin transport
Initiating leaf primordium Current Opinion in Plant Biology
Alternative hypotheses for explaining the establishment of adaxial-abaxial patterning in developing leaf primordia. (a) Model, in which a morphogen is produced by the meristem that promotes adaxial cell identity, thereby conferring polarity to newly developing leaf primordia [8]. (b) Pre-pattern model, in which organ position along the radial axis of the shoot is controlled by the same transcription factors (HD-ZIPIII and KANADI) that specify adaxial and abaxial tissue identity. These factors make sure organs arise at the boundary between their expression domains such that adaxial-abaxial patterning is transferred from meristem to leaf primordium [9]. (c) Auxin depletion model, in which, due to PIN1 mediated auxin transport, auxin is depleted in adaxial cells of new leaf primordia. This low level of auxin helps maintain or establish adaxial identity [11].
actively repressed by repressor-type Auxin Response Factors (ARFs) [12]. Hence, this model contrasts with the second model in terms of regulation of WOX1 and PRS in adaxial tissues (Figure 2). How does support for these competing models compare? An important piece of evidence supporting the prepattern model is that experimental alteration of this pre-pattern, solely in the meristem, is sufficient to cause major changes in leaf polarity, demonstrating its functional significance [9]. Furthermore, the abaxialization response to wounds, which inspired the meristem derived signal model [7] can also be explained by the meristem pre-pattern model since at least in Arabidopsis, wounding within the meristem results in auxin depletion [13] and this promotes ectopic KAN1 expression around the wound at the expense of REV, indicating abaxialization of the wounded tissue [9]. Supporting a causal role of auxin depletion, exogenous auxin treatment at the time of wounding can inhibit such a response [9]. However, Current Opinion in Plant Biology 2020, 53:73–79
although wounding Arabidopsis meristem tissues results in changes to adaxial and abaxial gene expressions, loss of leaf polarity was not observed. Instead, the Arabidopsis meristem bifurcates into two or more meristems and the leaves that form subsequently are normal. Hence an important task is to repeat the wounding experiments in potato and tomato and to test whether auxin can prevent polarity disruptions. What about the auxin depletion model, which also proposes that low auxin levels restrict WOX1 and PRS expression? This model is not necessarily mutually exclusive to the meristem pre-pattern model if the auxin depletion mechanism is required only for polarity maintenance after leaf initiation. However, beyond that, there are several inconsistencies between the auxin depletion and pre-pattern models. One major inconsistency is that auxin promotes HD-ZIPIII expression and restricts KAN1 expression [9], which is opposite to what one would expect if high auxin levels disrupt adaxial www.sciencedirect.com
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Figure 2
auxin
Absence of auxin (depletion model) or HD-ZIPIII expression (pre-pattern model)
Adaxial
PRS WOX1
Leaf margin
KAN ARF3/4
Abaxial Current Opinion in Plant Biology
Implications of alternative adaxial-abaxial hypotheses for the regulation of PRS and WOX1. Both the pre-pattern and auxin depletion models support a role for KANADI and ARF3/4 transcription factors in repressing PRS and WOX1 in abaxial leaf tissues. However the auxin depletion model proposes that it is a lack of auxin in adaxial tissues that prevents adaxial PRS and WOX1 expression while the pre-pattern model proposes that WOX1 and PRS are repressed adaxially directly or indirectly by the HD-ZIPIII proteins.
identity. A second inconsistency is that if auxin is applied exogenously to leaf primordia, WOX1 and PRS expression does not extend into adaxial leaf tissues, indicating that a lack of auxin is not restricting their expression domains [9] as has been proposed [12] (Figure 2). Finally, the finding that auxin levels are lower in adaxial leaf tissues has also come under dispute [14,15]. Despite these discrepancies however, no other model can explain why tomato leaf primordia become abaxialized upon adaxial auxin application. Do tomato leaves develop differently to Arabidopsis? Ultimately additional experiments in tomato may be needed that monitor auxin levels and adaxial and abaxial tissue identity in response to global and localized auxin manipulations. During the early stages of leaf growth the adaxial/middle (boundary)/abaxial pattern present within the founder cell population must be maintained during leaf development. A recent study has revealed how small RNAs act in a surprising way to help achieve this. Regulation of adaxial HD-ZIPIII and abaxial ARF genes is accomplished by small RNAs mir165/166 and ta-siARF, respectively [16] and since these small RNAs are known to move cell to cell from their sites of transcription in the adaxial or abaxial epidermis [16], they are anticipated to form a gradient across the leaf. In contrast, the defined expression of target genes to specific mesophyll cell layers suggest a response to threshold levels of these regulatory small RNA [17]. How might miRNA gradients be established? Although it remains to be established whether or not miRNA gradients are functional in early organ patterning, within the SAM, a miRNA and corresponding target reporter system has shown discrete domains of directional miRNA movement that is distinct from that of protein cell-to-cell movement [18]. miRNA distribution in the shoot meristem may include movement from sites of synthesis through plasmodesmata www.sciencedirect.com
as in the root where callous deposition and plasmamembrane plasmodesmata-associated receptor-like kinases BAM1 and BAM2 effect the spatial distribution of mir165/166 [19–21]. In the context of the three models proposed for establishing leaf adaxial/abaxial polarity, small RNAs may represent the Sussex signal and be sufficient for early patterning of the leaf. Alternatively, small RNAs may reinforce or act together with the other proposed mechanisms defining leaf adaxial/abaxial polarity.
Downstream of auxin Which genes apart from WOX1 and PRS are known to act downstream of auxin to promote organ growth? Several transcription factors are induced by auxin at primordial positions via Auxin Response Factor 5 (ARF5)/MONOPTEROS (MP) and function to promote flower development, through induction of genes including LEAFY, AINTEGUMENTA, AINTEGUMENTA-LIKE6 and FILAMENTOUS FLOWER (FIL) [22,23]. More recently, the LEAFLESS (LFS) gene from tomato, encoding a gene closely related to the Arabidopsis DORNRONSCHEN (DRN) and DRN-like (DRNL) genes, was found to be essential for leaf formation [24]. LFS is induced by auxin and is expressed at sites of auxin maxima in the shoot meristem. Loss of LFS results in a pin-like shoot where the shoot meristem fails to initiate organs-resembling shoot apices treated with auxin transport inhibitors. Like mp mutants, exogenous application of auxin is not sufficient to induce organ outgrowth from lfs apices [1,24]. As double mutants of DRN and DRNL in Arabidopsis show a similar phenotype to lfl in tomato, these results indicate LFL/DRN/ DRNL act as an essential transducer of the auxin signal leading to organ formation [24]. Another important transcriptional event downstream of auxin is downregulation of SHOOT MERISTEMLESS (STM). Recently it was Current Opinion in Plant Biology 2020, 53:73–79
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found that MP acts indirectly to achieve this by activating FIL. FIL forms a protein complex with ETT and ARF4, which directly represses STM, as well as the closely related BREVIPEDICELLUS (BP), through recruitment of the histone deactylase HDAC19 and subsequent transcriptional silencing [25].
Figure 3
Organ outgrowth
Leaf morphogenesis So far, we have discussed recent work focused on genetic regulatory networks which pattern cell types and regulate and are regulated by auxin. But how does auxin function at a cellular level to promote organ outgrowth and how is the shape of new organs determined? Recent work has provided important new insights into both these questions. Previous work has shown that auxin acts to promote organ formation at least in part through the local de-polymerization of microtubules. Since interphase microtubule arrangements help orient cellulose deposition in the cell wall, which helps orient cellular growth, this finding implied that a loss of growth anisotropy is sufficient to initiate organogenesis [26]. Is growth anisotropy the only relevant process triggered by microtubule de-polymerization? Using computer simulations, a more recent study concluded this cannot be the case [27]. Rather than increasing growth rates, as has been measured to occur at sites of organogenesis, computer simulations of growth using a mechanical model of the shoot meristem showed that local reductions in cell wall anisotropy should result in a decreased growth rate. This suggested that in addition to cell wall anisotropy, microtubule depolymerization likely influences other cell wall properties that increase growth rates. To investigate further, the authors documented the expression patterns of many genes encoding cell wall modifying enzymes and found several to be expression in the SAM and floral organ primordia. Interestingly, several of these genes were also upregulated in the organ-like outgrowths formed from the apex of double mutants of PIN1 and BOTERO1/KATANIN, encoding a microtubule associated protein, suggesting that auxin build up may not be required for organogenesis. This hypothesis was further supported by an absence of DR5 expression in organs induced by oryzalin application to NPA-induced ‘pin’ meristems. Lastly, since it had been shown that local application of cell wall modifying enzymes such as pectin methylesterase to NPA-pins can also induce outgrowths [28], the authors tested whether such treatments also induced a loss of microtubule anisotropy and found that they do, although the response was slower compared to auxin treatments [27]. Altogether these results reveal that several distinct processes are coupled during organ formation. High auxin concentrations result in a loss of microtubule array anisotropy and this is accompanied by the local upregulation of cell wall modifying enzymes. The latter two processes positively regulate each other and promoting either can trigger organ outgrowth (Figure 3). Whether the loss of microtubule anisotropy Current Opinion in Plant Biology 2020, 53:73–79
Expression of cell wall modifying enzymes
Microtubule depolymerisation
Auxin Current Opinion in Plant Biology
The role of feedback loops between the activity of cell wall modifying enzymes and microtubule depolymerization in mediating auxininduced organ formation. Feedback between microtubule depolymerization and the expression of cell wall enzymes is indicated by the solid arrows. Although auxin promotes both processes, it is not yet clear whether auxin promotes both independently or not (dashed arrows). It also remains unclear whether both microtubule depolymerization and cell wall enzyme expression promote organ outgrowth independently or not (dashed arrows).
or the induction of cell wall enzymes is sufficient to trigger outgrowth in the absence of the other remains to be determined (although simulations suggest loss of anisotropy is not sufficient), as do the molecular mechanisms through which this mutual regulation occurs. Given that organ initiation is accompanied by a loss of mechanical anisotropy, how is organ shape determined? The available data suggest that the loss of anisotropy only occurs during the initial outgrowth process and is restricted to the central point of auxin maximum [26]. Microtubule orientations otherwise are generally observed to be circumferentially oriented on the surface of organ primordia, consistent with the known role of mechanical stress in regulating their orientation [29]. These microtubule orientations would be expected to promote growth along the proximo-distal axis. Cell polarity, as marked by PIN1, also correlates with microtubule orientations linking polarity to growth direction and supporting the proposal that mechanical stresses also regulate cell polarity patterns [30]. Consistent with this scenario, a two-dimensional ‘polarity field’, (although in this case polarity is suggested to be patterned by a hypothetical diffusing signal), has been proposed to regulate lamina growth for the simple leaves of Arabidopsis [31]. This polarity field was proposed to promote www.sciencedirect.com
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growth both parallel and perpendicular to the field axis depending on position within the primordium, which in turn caused deformations in the field itself. This meant that although the field was initially oriented proximo-distally, it later diverged more laterally [31]. The observed growth directions were found to be consistent with this proposal, being mainly oriented proximo-distally early in development but later diverging towards the margin and becoming more isotropic [31]. Evidence that leaf cells are indeed polarized in a corresponding pattern has now been reported, based on the cellular localization patterns of the BASL protein. BASL is a novel protein that is asymmetrically localized in stomatal epidermal cell linages of the leaf [32]. Using an inducible GFP-BASL in a mutant background that lacks stomata, ectopic BASL expresses asymmetrically all epidermal cells of the leaf thus marking leaf polarity fields [33]. At early stages of leaf development, BASL polarity patterns exhibit a distal to proximal pattern along the same axis as distally oriented PIN1 but at late stages, it’s pattern diverges laterally as predicted [33]. However, further evidence that this polarity pattern is regulated by mechanics rather than a diffusible factor comes from the finding that BRXL2, a protein with asymmetric localization similar to BASL in the leaf, re-orients its polarity in response to mechanical perturbations [34]. The mechanism by which mechanical stress orients polarity over such large distances however remains speculative [33,35]. Given that PIN1 polarities and interphase microtubule array orientations are known to correlate in the SAM during organ initiation [30] and BASL also correlates with PIN1 polarity [33], it would seem likely that ectopic BASL polarity patterns are marking microtubule orientations during leaf development. A recent pre-print however indicates that while this is likely true early in leaf development, later in leaf development, as the leaf blade broadens, microtubule orientations become isotropic at the leaf surface, somewhat consistent with observed growth patterns but inconsistent with BASL polarity patterns [36]. The authors also find that while mechanical stress patterns can account for the observed microtubule orientations early on, the switch to a more isotropic pattern does not fit their model predictions. The authors conclude that at later developmental stages microtubules must become decoupled from stress patterns (although previous laser ablation experiments indicate microtubules do remain stress-responsive at least until pavement cells have formed [37]). In accordance with their modelling, they also predict that the switch to isotropy promotes leaf blade expansion [36]. All together these results indicate that at later stages of leaf development when the tissue differentiates, microtubule orientations and cell polarity patterns may become uncoupled and this may be important for leaf blade expansion. How do leaves become flat from a more symmetrical starting point? The same preprint mentioned above also investigates microtubule and cellulose patterns in the internal cell layers of the leaf. It reports that during both www.sciencedirect.com
early and late development, microtubules and cellulose fibrils within the anticlinal walls of leaf cells are oriented anticlinally, that is, perpendicular to the leaf surface and that according to model predictions, these orientations promote leaf flatness and follow the predicted stress patterns for the tissue (as mircrotubules are known to do elsewhere) [36]. How the primordium initially becomes asymmetric remains unclear. Although mechanical differences in wall stiffness on different sides of the leaf and have been proposed to account for initial leaf flattening [38], this hypothesis is under dispute [39,40]. Finally, what about complex leaves? Arabidopsis thaliana has simple leaves with an undivided leaf lamina and small serrations, or outgrowths, along the leaf margin. Leaf shape varies considerably in different plant species and can be simple, as in A. thaliana or may be complex, where the leaf lamina is divided into leaflets, as in Cardamine hirsute a close relative of A. thaliana. A feedback regulatory network involving auxin and the transcription factor CUC2 act along the leaf margin to control the formation of serrations in A. thaliana [41]. Furthermore, both auxin and CUC2 are required for development of distinct leaflets in compound leaves [42,43]. It has been less clear how other regulators of leaf shape are involved in compound leaf development. A recent study combining quantitative live-imaging, computational modelling and genetics provides a detailed look at how growth patterns are modified to go from simple leaves to complex [44]. The authors start by quantitatively comparing leaf growth patterns in A. thaliana and C. hirsute and with modelling and genetic manipulation find that the differences in morphology can largely be explained by the action of two homeobox genes, SHOOT MERISTEMLESS (STM) and REDUCED COMPLEXITY (RCO). In C. hirsute, the expression of STM (which is absent in A. thaliana leaves) acts broadly to slow growth but also extends the period of time, in which the leaf primordium remains undifferentiated. This is especially relevant to the distal and marginal part of the primordium where STM prolongs the growth phase in Cardamine and enables leaflets to form respectively [44]. In contrast, RCO acts locally to inhibit growth and is normally expressed at the base of Cardamine leaflets. By expressing both STM and RCO in A. thaliana leaves, the authors were able to create complex leaves in Arabidopsis that resembled those from Cardamine, thereby demonstrating how key genes enable morphological diversity [44].
Conclusion It is an exciting time for understanding plant organ development. The work discussed here represents significant progress in addressing questions that have been highlighted in the field for decades, facilitated by the use of quantitative live-imaging and modelling. Nevertheless, several findings remain under dispute and it will be important to resolve these disagreements Current Opinion in Plant Biology 2020, 53:73–79
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in order to make further progress and achieve a cohesive understanding across the field. In addition to the studies cited here, there have been many other important contributions to the literature on plant organ development recently and we regret that due to space constraints our review could not be exhaustive.
Conflict of interest statement Nothing declared.
Acknowledgement Funding is gratefully acknowledged by MH from the Australian Research Council (DP180101149).
References and recommended reading Papers of particular interest, published within the period of review, have been highlighted as: of special interest of outstanding interest 1.
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coupled to microtubule-driven growth isotropy at the shoot apex in Arabidopsis. Development 2018, 145. In this paper a feedback loop between the expression of cell remodelling genes and microtubule-depolymerization is discovered. 28. Peaucelle A, Braybrook SA, Le Guillou L, Bron E, Kuhlemeier C, Hofte H: Pectin-induced changes in cell wall mechanics underlie organ initiation in Arabidopsis. Curr Biol 2011, 21:1720-1726. 29. Hamant O, Heisler MG, Jonsson H, Krupinski P, Uyttewaal M, Bokov P, Corson F, Sahlin P, Boudaoud A, Meyerowitz EM et al.: Developmental patterning by mechanical signals in Arabidopsis. Science 2008, 322:1650-1655. 30. Heisler MG, Hamant O, Krupinski P, Uyttewaal M, Ohno C, Jonsson H, Traas J, Meyerowitz EM: Alignment between PIN1 polarity and microtubule orientation in the shoot apical meristem reveals a tight coupling between morphogenesis and auxin transport. PLoS Biol 2010, 8:e1000516. 31. Kuchen EE, Fox S, de Reuille PB, Kennaway R, Bensmihen S, Avondo J, Calder GM, Southam P, Robinson S, Bangham A et al.: Generation of leaf shape through early patterns of growth and tissue polarity. Science 2012, 335:1092-1096. 32. Dong J, MacAlister CA, Bergmann DC: BASL controls asymmetric cell division in Arabidopsis. Cell 2009, 137:1320-1330. 33. Mansfield C, Newman JL, Olsson TSG, Hartley M, Chan J, Coen E: Ectopic BASL reveals tissue cell polarity throughout leaf development in Arabidopsis thaliana. Curr Biol 2018, 28:26382646 e2634. This study reveals that cells remain polarized in the epidermis of leaves until late developmental stages. The polarity pattern resembles the pattern predicted through modelling in an earlier study. 34. Bringmann M, Bergmann DC: Tissue-wide mechanical forces influence the polarity of stomatal stem cells in Arabidopsis. Curr Biol 2017, 27:877-883. 35. Bhatia N, Heisler MG: Self-organizing periodicity in development: organ positioning in plants. Development 2018, 145. This review summarizes our current understanding of how cell polarity patterns are regulated in relation to phyllotaxis. 36. Zhao F, Du F, Oliveri H, Zhou L, Ali O, Chen W, Feng S, Wang Q, Lu¨ S, Long M et al.: A microtubule-mediated mechanical feedback controls leaf blade development in three dimensions. bioRxiv 2019:604710. This pre-print reveals patterns of microtubule orientations during leaf development and shows that some observed patterns match predictions
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based on mechanical stresses while others do not. The authors also propose that a loss of anisotropy in the epidermis during leaf development promotes leaf flattening. 37. Sampathkumar A, Krupinski P, Wightman R, Milani P, Berquand A, Boudaoud A, Hamant O, Jonsson H, Meyerowitz EM: Subcellular and supracellular mechanical stress prescribes cytoskeleton behavior in Arabidopsis cotyledon pavement cells. eLife 2014, 3:e01967. 38. Qi J, Wu B, Feng S, Lu S, Guan C, Zhang X, Qiu D, Hu Y, Zhou Y, Li C et al.: Mechanical regulation of organ asymmetry in leaves. Nat Plants 2017, 3:724-733. This study measures the mechanical properties of adaxial and abaxial leaf tissues using Atomic Force Microscopy. Using modelling and measurements, the authors propose that the differences they observe can account for leaf flattening. 39. Feng S, Zhou L, Lu S, Long M, Jiao Y: Reply to ‘Early shaping of a leaf’. Nat Plants 2018, 4:620-621. This is a reply to Ref. [40]. 40. Coen E, Kennaway R: Early shaping of a leaf. Nat Plants 2018, 4:618-619. In this commentary, the authors raise important issues with a previously proposed mechanical model used to simulate leaf growth (see Ref. [38]). 41. Bilsborough GD, Runions A, Barkoulas M, Jenkins HW, Hasson A, Galinha C, Laufs P, Hay A, Prusinkiewicz P, Tsiantis M: Model for the regulation of Arabidopsis thaliana leaf margin development. Proc Natl Acad Sci U S A 2011, 108:3424-3429. 42. Barkoulas M, Hay A, Kougioumoutzi E, Tsiantis M: A developmental framework for dissected leaf formation in the Arabidopsis relative Cardamine hirsuta. Nat Genet 2008, 40:1136-1141. 43. Blein T, Pulido A, Vialette-Guiraud A, Nikovics K, Morin H, Hay A, Johansen IE, Tsiantis M, Laufs P: A conserved molecular framework for compound leaf development. Science 2008, 322:1835-1839. 44. Kierzkowski D, Runions A, Vuolo F, Strauss S, Lymbouridou R, Routier-Kierzkowska AL, Wilson-Sanchez D, Jenke H, Galinha C, Mosca G et al.: A Growth-based framework for leaf shape development and diversity. Cell 2019, 177:1405-1418 e1417. This paper provides a dynamic and quantitative analysis of leaf growth, focusing on the influence of different genes in regulating leaf morphology. By manipulating the expression of two genes, the authors demonstrate the ability to create Cardamine-like complex leaf morphologies in Arabidopsis.
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