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Regulation of shoot branching by auxin
Review
TRENDS in Plant Science
Vol.8 No.11 November 2003
541
Regulation of shoot branching by auxin Ottoline Leyser Department of Biology, University of York, Heslington, York, UK YO10 5YW
The idea that apically derived auxin inhibits shoot branching by inhibiting the activity of axillary buds was first proposed 70 years ago, but it soon became clear that its mechanism of action was complex and indirect. Recent advances in the study of axillary bud development and of auxin signal transduction are allowing a better understanding of the role of auxin in controlling shoot branching. These studies have identified a new role for auxin early in bud development as well as some of the second messengers involved in mediating the branch-inhibiting effects of auxin. Plant development is continuous and modular through the action of meristems. The timing and the frequency with which the meristems produce each module, including the production of new meristems, is regulated by the genetic makeup of the plant, the developmental stage of the plant, and by the environment in which the plant is growing. The integration of genetic, developmental and environmental information during the decision to produce each new module allows plants to achieve an astonishing level of developmental plasticity, with the body plan of plants of a single species varying radically depending on genotype, age and the prevailing environmental conditions. This feat of signal integration is likely to be dependent on the action of plant hormones. As hormone signalling pathways become better understood, there is now an unprecedented opportunity to investigate their roles in mediating developmental plasticity. The degree of shoot branching is an excellent system in which to study this problem. In the shoot, the developmental module consists of a stem segment (an internode), a leaf, and usually at least one shoot meristem laid down in the leaf axil. These axillary meristems can activate to give rise to an entire secondary shoot, consisting of similar modules, or they can remain dormant. The activity of axillary meristems is regulated by a wide range of genetic, developmental and environmental factors and hormones play central roles in mediating these diverse influences. For example, more than 70 years of research have clearly demonstrated a role for auxin in inhibiting shoot branching [1]. A major site for auxin synthesis is at the primary shoot apex, particularly in young leaves, from which it is transported down the plant in the polar transport stream [2]. Auxin moving in this way can inhibit shoot branching because either blocking polar auxin transport or removing Corresponding author: Ottoline Leyser ([email protected]).
the shoot apex promotes shoot branching, with the branchpromoting effect of removing the apex being negated by the addition of auxin to the decapitated stump [3]. This role for the shoot apex, mediated at least partly by auxin, has been termed apical dominance. In spite of its long history, the mechanism of auxin action in the control of shoot branching is still poorly understood and hence its potential role as a signal integrator cannot easily be assessed. Furthermore, there are multiple levels during the development of a branch at which auxin could influence events. Axillary shoot development is divisible into at least three stages: axillary meristem initiation, meristem dormancy and active meristem growth. Recent genetic approaches in tomato, pea and Arabidopsis have begun to improve our understanding of the role of auxin at these various stages significantly. Axillary meristem initiation There are two theories about the initiation of axillary meristems. The detached meristem hypothesis proposes that axillary meristems are derived directly from cells of the shoot apical meristem, which never lose their meristematic identity [4,5]. The alternative model proposes that axillary meristems initiate de novo from cells in the leaf axil [6]. These two models have persisted because histologically, axillary meristem origins appear to be different in different species. In many plants, such as potato, meristematic cells are obvious at the base of each leaf at the time of leaf inception on the flanks of the primary shoot apical meristem. These meristems are apparently not derived from the cells of their subtending leaves because they have been shown to come from the same population of cells as the leaf above them, rather than from the leaf below [7,8]. By contrast, the axillary meristems of vegetative Arabidopsis plants are not visible until long after leaf initiation [9– 11]. Both histological and clonal analysis indicates that they are derived from cells at the base of the leaf petiole [9 –13]. This apparent interspecific variation in the provenance of axillary meristems has fuelled the longrunning debate between the reserved and de novo axillary meristem hypotheses. As usual, the answer appears to be somewhere in between the two. The idea that different species have different mechanisms for axillary meristem initiation is weakened by the observation that there is dramatic variation through the life cycle of a plant in the size of the axillary meristems at their inception, relative both to the subtending leaf and to the primary shoot apical meristem (Figure 1). For example,
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SAM
Foral node
Primordium AM component Leaf component
Cauline node
Rosette node
TRENDS in Plant Science
Figure 1. Primordia produced at the flanks of the Arabidopsis shoot apical meristem (SAM) produce a leaf and an axillary meristem (AM, shown in red). The distribution of the primordium between these two structures varies through the life cycle. During vegetative growth (rosette nodes), no histologically distinct axillary meristems are observed in the leaf axils at the time of leaf inception, but a small number of cells on the adaxial side of the primordium express the LAS gene and have the potential to become axillary meristems. The axillary meristems of the leaves that are made immediately before floral transition (cauline nodes) develop rapidly into a histologically distinct dome while the leaf is still at an early stage of development, and the flowers (floral nodes) are produced from axillary meristems, with the development of their subtending leaf being completely suppressed.
during vegetative growth of Arabidopsis, no histologically distinct axillary meristems are observed in the leaf axils until late in leaf development when meristems apparently develop from a small number of cells at the petiole base. However, the axillary meristems of the leaves that are made immediately before floral transition develop much more rapidly while the leaf primordia are still at an early stage. The flowers of Arabidopsis are produced from axillary meristems that appear to be derived more directly from the primary shoot apical meristem at the same time as their subtending leaves, with the development of the leaf being completely suppressed [9]. Hence, the primary meristem of Arabidopsis can be considered to produce primordia that will give rise to a leaf and to an axillary meristem. During vegetative development, the primordium is used almost entirely to make the leaf, but after floral transition the primordium is used almost entirely to make the axillary meristem (Figure 1). Similar variation can be seen in tomato, where the size of the axillary meristem also increases through the life cycle of the plant. At floral transition a sympodial branching habit is observed in which the primary shoot apical meristem appears to bifurcate because the axillary meristem of the last initiated leaf consumes half the primary shoot apical meristem [14]. The primary meristem becomes determinate and floral, and the axillary meristem assumes the role of the primary meristem. This wide range of observed origins for axillary meristems within a single plant suggests that the opposing models of detached versus de novo are opposite ends of a continuum rather than distinct and different mechanisms. A recent exciting advance in this area has been the molecular identification of mutationally defined tomato genes that are required for axillary meristem initiation. http://plants.trends.com
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The best characterized of these is the lateral suppressor gene, Ls, which encodes a GRAS domain protein that is likely to be a transcription factor [15]. Loss of Ls function in tomato results in loss of axillary meristems in the vegetative nodes, but sympodial branching post-floral transition is unaffected. Recently, an Arabidopsis orthologue of Ls named LAS has been identified [16]. Loss of LAS function results in absence of axillary meristems in most of the vegetative nodes of Arabidopsis with only the most apical nodes producing branches after floral transition. A genomic clone of the Arabidopsis orthologue, when introduced into ls mutant tomato, can completely restore a wild-type phenotype, demonstrating conservation of the mechanism of axillary meristem formation in these two species with apparently different axillary meristem biology. An orthologous gene has also been discovered in rice [17]. Furthermore, in situ hybridization experiments demonstrate that, even in vegetative Arabidopsis plants, LAS is expressed in the primary shoot apical meristem at the adaxial margins of initiating leaf primordia, where it appears to act to protect primordial cells from differentiation, supporting the reserve meristem hypothesis [16]. Auxin and axillary meristem initiation In addition to contributing to our understanding of the origins of axillary meristems, analysis of Arabidopsis las mutants has revealed a previously unsuspected role for auxin in axillary meristem initiation. Characterization of the auxin-resistant Arabidopsis mutant axr1 had suggested that the effect of auxin on shoot branching is entirely after axillary meristem initiation, through the inhibition of bud out growth [11]. As predicted for a mutant with reduced auxin response, the axr1 mutant is highly branched. However, no histological differences in the timing of axillary meristem initiation could be detected in axr1 mutants, leading to the conclusion that the bushy phenotype was entirely the result of an effect on bud outgrowth [11]. If this were the case, then one would predict that las should be completely epistatic to axr1 because the effect of the axr1 mutation should be entirely after that of the las mutation. However, the axr1 mutation is able weakly to suppress the phenotype conferred by las [16]. In the las single mutant, only 0 –2 side shoots develop from rosette nodes, but in the axr1 las double mutant, up to five such branches were observed. This interesting new role for auxin needs to be investigated further. In the mean time, most attention has focused on the more dramatic later role of auxin in regulating bud outgrowth. Auxin and bud outgrowth After initiation, axillary meristems produce a few leaves to form a bud. The buds can then become dormant or they can continue growth to produce a side shoot. Furthermore, dormant buds can later reactivate to produce a side shoot (see [18] and the Review article by David Horvath et al. in this of Trends in Plant Science [19]). ‘Dormant’ is a somewhat misleading term for the non-growing bud because such buds are highly metabolically active producing a characteristic set of transcripts and proteins. The switch to activity is accompanied by a change in this pattern but,
Review
TRENDS in Plant Science
within a small time window, activation can be reversed and dormancy can be reimposed, so that axillary buds can cycle between the dormant and active states (see [18] and Horvath et al. in this issue of Trends in Plant Science [19]). It has been known for many decades that auxin travelling in the polar transport stream promotes the dormant state in axillary buds. However, its mode of action is obscure because for almost as long it has been known that auxin does not act directly to suppress bud growth. Direct application of auxin to buds does not inhibit their outgrowth [20] and levels of auxin in buds rise as they activate [21]. Furthermore, apically applied auxin is not transported into the buds [22]. For example, in Arabidopsis, an assay has been developed in which an excised stem segment, bearing a single leaf, is inserted between two agar blocks in a Petri dish [23]. The axillary bud in the leaf axil activates and grows out to produce a side shoot. Addition of auxin to the apical, but not to the basal block, inhibits the outgrowth of the bud. Inhibitors of polar transport block the effect of apical auxin and buds of the axr1 mutant are resistant to the effect of apically applied auxin. However, when radio-labelled auxin is used in this assay, less accumulates in the bud when the auxin is applied apically than when it is applied basally [24]. This strongly supports the idea that auxin acts indirectly, which might be predicted given that to enter a bud, auxin would have to move acropetally from the primary axis into the leaf petiole, against the flow of the polar transport stream. This indirect mode of action for auxin has led to the hypothesis that a second messenger carries the auxin signal into the bud. Several candidates have been proposed, with the strongest contender being cytokinin. Cytokinin can promote the outgrowth of buds when directly applied to them, and cytokinin levels rise in buds as they are activated [3,25]. Furthermore, cytokinin supplied through the basal cut surface of an excised Arabidopsis nodal segment can overcome the inhibitory effects of apically applied auxin on the growth of axillary buds [23]. The link between auxin and cytokinin comes from two sources. First, it has been shown that in bean (Phaseolus vulgaris), decapitation results in an increase in export of cytokinin from roots in the xylem [26]. This increase can be partly blocked by applying auxin to the decapitated stump. Second, it has been shown that auxin can down-regulate cytokinin synthesis [27,18]. These data suggest that one mode of action for auxin could be to down-regulate both local cytokinin synthesis and cytokinin export from roots and that this might prevent the activation of buds. Novel second messengers Although action through cytokinins probably accounts for part of the effect of apical auxin, it appears not to be the only mode of auxin action. There is growing evidence for a second long-range signalling system that interacts with auxin to inhibit shoot branching. In recent years, evidence has accumulated for the involvement of an entirely novel signal in relaying the auxin message into the bud. This has come from the analysis of a series of bushy mutants in pea and Arabidopsis, called the rms and max mutants, respectively [28 – 31]. The mutants are characterized by increased http://plants.trends.com
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branching, reduced stature and in most cases shorter leaves. The axillary buds of all the rms and max mutants tested to date show at least a reduced response to apically applied auxin following decapitation (rms mutants) or in the isolated node assay (max mutants) [29,32]. Thus, these genes are required for a wild-type auxin-mediated inhibition of bud outgrowth. A sub-set of the RMS and MAX genes are excellent candidates for involvement in the production of the predicted auxin second messenger. The highly branched phenotype of mutant shoots of max1, max3, max4, rms1, rms2 and rms5 can be restored to wild type by grafting to wild-type root stocks [29,30,33– 35]. Thus, these genes are probably required for the production of a long-range graft-transmissible, branch-inhibiting signal. Hormonal measurements suggest that the rms1 phenotype is not associated with either reduced auxin or increased cytokinin, implicating a novel factor [32]. Because all the mutants tested to date have auxin-resistant bud growth in decapitation assays, this long-range signal must be required for auxin-mediated apical dominance. Further grafting studies have revealed other properties of this signal [29,30,33– 35] (Figure 2). For example, the reciprocal experiment in which wild-type shoots are grafted to mutant roots also results in wild-type shoot branching. This demonstrates that the signal can be synthesized all along the plant axis, and, just as production in only the root is sufficient to restore wild-type branching, so is production in only the shoot. Small wild-type interstock segments grafted between mutant tissues in the shoot are also sufficient to restore wild-type shoot branching to the shoot system. This suggests that the signal is either potent or can be made in large quantities. The use of a different grafting techniques, so-called Y-grafted, suggests that the signal can only move up plants [30,34]. In these experiments, a wild-type shoot is grafted into the root– shoot junction of a mutant plant, but the mutant shoot is not restored to wild-type, indicating that the signal cannot move down the plant from the wild-type shoot and then back up to restore a wild-type branching habit to the mutant shoot system. What then is this potent upwardly mobile signal? Some hints have come from the molecular analysis of the MAX4 and RMS1 genes, which encode orthologous members of the carotenoid cleaving dioxygenase family [29]. The MAX4 and RMS1 proteins are predicted to be plastidic because they contain potential plastid-targeting sequences. Plastids are the site of carotenoid synthesis in plants. Hence, an attractive hypothesis is that the longrange branch-inhibiting signal is derived by cleavage of a carotenoid. However, other members of this diverse family have non-carotenoid substrates and some do not catalyse cleavage reactions and instead are involved in isomerization [36]. However, if the hypothesis were correct it would add to the growing number of plant growth regulating compounds derived from carotenoids. Apart from the wellcharacterized hormone abscisic acid, there is some evidence that b-ionone, a carotenoid cleavage product, has hormonal activity [37,38]. The molecular characterization of the MAX4 and RMS1 genes has also allowed an investigation of the interaction
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Wild-type shoot Wild-type root
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Mutant shoot Wild-type root
Wild-type shoot Mutant root
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Mutant shoot Wild-type segment Mutant root TRENDS in Plant Science
Figure 2. Summary of the results of grafting experiments between the rms1, rms2 and rms5 mutants (green) of pea with their wild types (purple) and the max1, max3 and max4 mutants (green) of Arabidopsis with their wild type (purple). Self grafts of the wild types have wild-type branching patterns (a) and self grafts of the mutants result in typical mutant bushy plants (b). Wild-type roots can restore wild-type branching to mutant shoots (c) and wild-type shoots show wild-type branching patterns when grafted to mutant roots (d). For the genotypes tested to date, small interstock wild-type grafts restore wild-type branching to nodes above the graft (e).
of auxin and the MAX/RMS-dependent signal [29]. Interestingly, the pea and Arabidopsis systems diverge here. In pea, there is evidence that auxin can rapidly up-regulate the transcription of RMS1 in the stem. In this case, auxin could inhibit bud growth by locally increasing the amount of RMS1, and hence presumably the product of the reaction it catalyses. By contrast, neither PCR-based approaches nor promoter-GUS reporter constructs have shown evidence for auxin regulating the transcription of MAX4 at the node. Instead, some auxin-induced MAX4 expression was observed behind the root tip, but this only occurs 24 h after auxin treatment. Hence, in Arabidopsis, a post-transcriptional mode of action for auxin in regulating the MAX pathway is likely. Transcription of MAX4 does not appear to be a limiting step in branch inhibition because plants overexpressing MAX4 from the cauliflower mosaic virus 35S promoter are as branched as wild-type plants are. Once synthesized, the mechanism of action by which the MAX/RMS-dependent signal inhibits buds is unknown. Some clues come from the molecular analysis of the MAX2 gene. The max2 mutants are phenotypically near identical to the max1, max3 and max4 mutants. However, the max2 shoot phenotype cannot be restored to wild type by grafting to wild-type roots (J. Booker, C. Turnbull and O. Leyser, unpublished). Therefore, MAX2 acts in the shoot to regulate branching and could be involved in the perception of the MAX-dependent signal. Similar conclusions have been drawn from grafting experiments carried out with the rms3 and rms4 mutants [39,28]. Consistent with this idea, MAX2 encodes an F-box protein. F-box proteins are involved in the selection of specific substrates for ubiquitin-mediated degradation [40]. There are ,700 F-box proteins in the Arabidopsis genome [41], and several of the better-characterized family members have roles in hormone signalling [40]. For example, an F-box protein, TIR1, mediates the auxin-induced degradation of the members of the Aux/IAA family of transcriptional repressors, thus controlling auxin-induced gene expression [42]. http://plants.trends.com
A similar system could operate in the transduction of the MAX/RMS-dependent signal. Conclusions It was first shown in 1933 that apically derived auxin could inhibit axillary bud growth. More than 70 years later the mechanism by which auxin regulates shoot branching is still unknown. Recent results show that auxin can influence various stages of bud development, from a minor role in bud initiation through to a major but indirect effect on bud activity involving at least two different hormonal second messengers. It will be particularly exciting to find out the identity of the novel unknown signal that acts downstream of auxin. In the future, the hormonal control of shoot branching should provide an excellent model for the study of the integration of environmental and genetic controls in plant development. Here, the comparison of the regulation of branching in different species with different regulatory balances should be particularly informative, and already RMS1 and MAX4 provide an example of orthologous genes that are regulated differently. These recent advances have come from the powerful combination of physiological, genetic, biochemical and genomic resources, which are now available to researchers in this field, promising rapid continued progress. References 1 Thimann, K. and Skook, F. (1933) Studies on the growth hormone of plants III: the inhibitory action of the growth substance on bud development. Proc. Natl. Acad. Sci. U. S. A. 19, 714 – 716 2 Ljung, K. et al. (2001) Sites and homeostatic control of auxin biosynthesis in Arabidopsis during vegetative growth. Plant J. 28, 465 – 474 3 Cline, M.G. (1991) Apical dominance. Bot. Rev. 57, 318 – 358 4 Garrison, R. (1955) Studies in the development of axillary buds. Am. J. Bot. 42, 257 – 266 5 Sussex, I.M. (1955) Morphogenesis in Solanum tuberosum L.: experimental investigation of leaf dorsoventrality and orientation in the juvenile shoot. Phytomorphology 5, 286 – 300 6 Snow, M. and Snow, R. (1942) The determination of axillary buds. New Phytol. 41, 13 – 22
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7 Johri, M.M. and Coe, E.H. (1983) Clonal analysis of corn plant development. 1. The development of the tassel and the ear shoot. Dev. Biol. 97, 154 – 172 8 McDaniel, C.N. and Poethig, R.S. (1988) Cell-lineage patterns in the shoot apical meristem of the germinating maize embryo. Planta 175, 13 – 22 9 Long, J. and Barton, M.K. (2000) Initiation of axillary and floral meristems in Arabidopsis. Dev. Biol. 218, 341 – 353 10 Gribic, V. and Bleecker, A.B. (2000) Axillary meristem development in Arabidopsis thaliana. Plant J. 21, 215 – 223 11 Stirnberg, P. et al. (1999) AXR1 acts after lateral bud formation to inhibit lateral bud growth in Arabidopsis. Plant Physiol. 121, 839– 847 12 Irish, V.F. and Sussex, I.M. (1992) A fate map of the Arabidopsis embryonic shoot apical meristem. Development 115, 745 – 753 13 Furner, I.J. and Pumfrey, J.E. (1992) Cell fate in the shoot apical meristem of Arabidopsis thaliana. Development 115, 755– 764 14 Schmitz, G. and Theres, K. (1999) Genetic control of branching in Arabidopsis and tomato. Curr. Opin. Plant Biol. 2, 51– 55 15 Schumacher, K. et al. (1999) The Lateral suppressor (Ls) gene of tomato encodes a new member of the VHIID protein family. Proc. Natl. Acad. Sci. U. S. A. 96, 290 – 295 16 Greb, T. et al. (2003) Molecular analysis of the LATERAL SUPPRESSOR gene in Arabidopsis reveals a conserved control mechanism for axillary meristem formation. Genes Dev. 17, 1175– 1187 17 Li, X. et al. (2003) Control of tillering in rice. Nature 422, 618– 621 18 Shimizu-Sato, S. and Mori, H. (2001) Control in outgrowth and dormancy in axillary buds. Plant Physiol. 127, 1405 – 1413 19 Horvarth, D.P. et al. (2003) Knowing when to grow: signals regulating bud dormancy. Trends Plant Sci. 8(11) (2003), this issue. doi:10.1016/ j.tplants.2003.09.013 20 Cline, M.G. (1996) Exogenous auxin effects on lateral bud outgrowth in decapitated shoots. Ann. Bot. 78, 255 – 266 21 Hillman, J.R. et al. (1977) Apical dominance and levels of IAA in Phaseolus lateral buds. Planta 134, 191 – 193 22 Morris, D.A. (1977) Transport of exogenous auxin in two-branched dwarf pea seedlings (Pisum sativum L.). Planta 136, 91 – 96 23 Chatfield, S.P. et al. (2000) The hormonal regulation of axillary bud growth in Arabidopsis. Plant J. 24, 159 – 169 24 Booker, J. et al. (2003) Auxin acts in xylem associated/medullary cells to mediate apical dominance. Plant Cell 15, 495 – 507 25 Turnbull, C.N.G. et al. (1997) Rapid increase in cytokinin concentration in lateral buds of chick pea (Cicer anerinum) during release of apical dominance. Planta 202, 271 – 276
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26 Bangerth, F. (1994) Response of cytokinin concentration in xylem exude of beans (Phaseolus vulgaris L.) plants to decapitation and auxin treatment and relationship to apical dominance. Planta 194, 439– 442 27 Eklof, S. et al. (1997) Auxin – cytokinin interactions in wild-type and transgenic tobacco. Plant Cell Physiol. 38, 225 – 235 28 Beveridge, C.A. (2000) Long distance signalling and a mutational analysis of branching in pea. Plant Growth Regul. 32, 193 – 203 29 Sorefan, K. et al. (2003) MAX4 and RMS1 are orthologous dioxygenase-like genes that regulate shoot branching in Arabidopsis and pea. Genes Dev. 17, 1469 – 1474 30 Turnbull, C.N.G. et al. (2002) Micrografting techniques for testing long distance signalling in Arabidopsis. Plant J. 32, 255 – 262 31 Stirnberg, P. et al. (2002) MAX1 and MAX2 control shoot lateral branching in Arabidopsis. Development 129, 1131 – 1141 32 Beveridge, C. et al. (2000) Auxin inhibition of decapitation-induced branching is dependent of graft-transmissible signals regulated by genes rms1 and rms2. Plant Physiol. 123, 689– 697 33 Beveridge, C.A. et al. (1997) RMS mutant has increased IAA and decreased root sap zeatin riboside content but increased branching, controlled by graft transmissible signal(s). Plant Physiol. 115, 1251– 1258 34 Foo, E. et al. (2001) Long distance signalling and the control of branching in the rms1 mutant of pea. Plant Physiol. 126, 203 – 209 35 Morris, S.E. et al. (2001) Mutational analysis of branching in pea (Pisum sativum L.): evidence that Rms1 and Rms5 regulate the same novel signal. Plant Physiol. 126, 1205– 1213 36 Schwartz, S.H. et al. (2001) Characterization of a novel carotenoid cleavage dioxygenase from plants. J. Biol. Chem. 276, 25208 – 25211 37 Redmond, T.M. et al. (1998) Rpe65 is necessary for the production of 11-cis-vitamin A in the retinal visual cycle. Nat. Genet. 20, 344– 351 38 Katonoguchi, H. (1994) Occurrence of a growth inhibitor, 3-hydroxybeta-ionone, in 7 cultivars of Phaseolus vulgaris and its role in lightinduced growth-inhibition. Phytochemistry 36, 273 – 275 39 Beveridge, C.A. et al. (1996) Branching in pea – action of genes rms3 and rms4. Plant Physiol. 110, 859 – 865 40 Hellmann, H. and Estelle, M. (2002) Plant development: regulation by protein degradation. Science 297, 793 – 797 41 Gagne, J.M. et al. (2002) The F-box subunit of the SCF E3 complex is encoded by a diverse superfamily of genes in Arabidopsis. Proc. Natl. Acad. Sci. U. S. A. 99, 11519 – 11524 42 Gray, W.M. et al. (2001) Auxin regulates SCFTIR1-dependent degradation of Aux/IAA proteins. Nature 414, 271 – 276
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Regulation of shoot branching by auxin Ottoline Leyser Department of Biology, University of York, Heslington, York, UK YO10 5YW
The idea that apically derived auxin inhibits shoot branching by inhibiting the activity of axillary buds was first proposed 70 years ago, but it soon became clear that its mechanism of action was complex and indirect. Recent advances in the study of axillary bud development and of auxin signal transduction are allowing a better understanding of the role of auxin in controlling shoot branching. These studies have identified a new role for auxin early in bud development as well as some of the second messengers involved in mediating the branch-inhibiting effects of auxin. Plant development is continuous and modular through the action of meristems. The timing and the frequency with which the meristems produce each module, including the production of new meristems, is regulated by the genetic makeup of the plant, the developmental stage of the plant, and by the environment in which the plant is growing. The integration of genetic, developmental and environmental information during the decision to produce each new module allows plants to achieve an astonishing level of developmental plasticity, with the body plan of plants of a single species varying radically depending on genotype, age and the prevailing environmental conditions. This feat of signal integration is likely to be dependent on the action of plant hormones. As hormone signalling pathways become better understood, there is now an unprecedented opportunity to investigate their roles in mediating developmental plasticity. The degree of shoot branching is an excellent system in which to study this problem. In the shoot, the developmental module consists of a stem segment (an internode), a leaf, and usually at least one shoot meristem laid down in the leaf axil. These axillary meristems can activate to give rise to an entire secondary shoot, consisting of similar modules, or they can remain dormant. The activity of axillary meristems is regulated by a wide range of genetic, developmental and environmental factors and hormones play central roles in mediating these diverse influences. For example, more than 70 years of research have clearly demonstrated a role for auxin in inhibiting shoot branching [1]. A major site for auxin synthesis is at the primary shoot apex, particularly in young leaves, from which it is transported down the plant in the polar transport stream [2]. Auxin moving in this way can inhibit shoot branching because either blocking polar auxin transport or removing Corresponding author: Ottoline Leyser ([email protected]).
the shoot apex promotes shoot branching, with the branchpromoting effect of removing the apex being negated by the addition of auxin to the decapitated stump [3]. This role for the shoot apex, mediated at least partly by auxin, has been termed apical dominance. In spite of its long history, the mechanism of auxin action in the control of shoot branching is still poorly understood and hence its potential role as a signal integrator cannot easily be assessed. Furthermore, there are multiple levels during the development of a branch at which auxin could influence events. Axillary shoot development is divisible into at least three stages: axillary meristem initiation, meristem dormancy and active meristem growth. Recent genetic approaches in tomato, pea and Arabidopsis have begun to improve our understanding of the role of auxin at these various stages significantly. Axillary meristem initiation There are two theories about the initiation of axillary meristems. The detached meristem hypothesis proposes that axillary meristems are derived directly from cells of the shoot apical meristem, which never lose their meristematic identity [4,5]. The alternative model proposes that axillary meristems initiate de novo from cells in the leaf axil [6]. These two models have persisted because histologically, axillary meristem origins appear to be different in different species. In many plants, such as potato, meristematic cells are obvious at the base of each leaf at the time of leaf inception on the flanks of the primary shoot apical meristem. These meristems are apparently not derived from the cells of their subtending leaves because they have been shown to come from the same population of cells as the leaf above them, rather than from the leaf below [7,8]. By contrast, the axillary meristems of vegetative Arabidopsis plants are not visible until long after leaf initiation [9– 11]. Both histological and clonal analysis indicates that they are derived from cells at the base of the leaf petiole [9 –13]. This apparent interspecific variation in the provenance of axillary meristems has fuelled the longrunning debate between the reserved and de novo axillary meristem hypotheses. As usual, the answer appears to be somewhere in between the two. The idea that different species have different mechanisms for axillary meristem initiation is weakened by the observation that there is dramatic variation through the life cycle of a plant in the size of the axillary meristems at their inception, relative both to the subtending leaf and to the primary shoot apical meristem (Figure 1). For example,
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Review
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SAM
Foral node
Primordium AM component Leaf component
Cauline node
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TRENDS in Plant Science
Figure 1. Primordia produced at the flanks of the Arabidopsis shoot apical meristem (SAM) produce a leaf and an axillary meristem (AM, shown in red). The distribution of the primordium between these two structures varies through the life cycle. During vegetative growth (rosette nodes), no histologically distinct axillary meristems are observed in the leaf axils at the time of leaf inception, but a small number of cells on the adaxial side of the primordium express the LAS gene and have the potential to become axillary meristems. The axillary meristems of the leaves that are made immediately before floral transition (cauline nodes) develop rapidly into a histologically distinct dome while the leaf is still at an early stage of development, and the flowers (floral nodes) are produced from axillary meristems, with the development of their subtending leaf being completely suppressed.
during vegetative growth of Arabidopsis, no histologically distinct axillary meristems are observed in the leaf axils until late in leaf development when meristems apparently develop from a small number of cells at the petiole base. However, the axillary meristems of the leaves that are made immediately before floral transition develop much more rapidly while the leaf primordia are still at an early stage. The flowers of Arabidopsis are produced from axillary meristems that appear to be derived more directly from the primary shoot apical meristem at the same time as their subtending leaves, with the development of the leaf being completely suppressed [9]. Hence, the primary meristem of Arabidopsis can be considered to produce primordia that will give rise to a leaf and to an axillary meristem. During vegetative development, the primordium is used almost entirely to make the leaf, but after floral transition the primordium is used almost entirely to make the axillary meristem (Figure 1). Similar variation can be seen in tomato, where the size of the axillary meristem also increases through the life cycle of the plant. At floral transition a sympodial branching habit is observed in which the primary shoot apical meristem appears to bifurcate because the axillary meristem of the last initiated leaf consumes half the primary shoot apical meristem [14]. The primary meristem becomes determinate and floral, and the axillary meristem assumes the role of the primary meristem. This wide range of observed origins for axillary meristems within a single plant suggests that the opposing models of detached versus de novo are opposite ends of a continuum rather than distinct and different mechanisms. A recent exciting advance in this area has been the molecular identification of mutationally defined tomato genes that are required for axillary meristem initiation. http://plants.trends.com
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The best characterized of these is the lateral suppressor gene, Ls, which encodes a GRAS domain protein that is likely to be a transcription factor [15]. Loss of Ls function in tomato results in loss of axillary meristems in the vegetative nodes, but sympodial branching post-floral transition is unaffected. Recently, an Arabidopsis orthologue of Ls named LAS has been identified [16]. Loss of LAS function results in absence of axillary meristems in most of the vegetative nodes of Arabidopsis with only the most apical nodes producing branches after floral transition. A genomic clone of the Arabidopsis orthologue, when introduced into ls mutant tomato, can completely restore a wild-type phenotype, demonstrating conservation of the mechanism of axillary meristem formation in these two species with apparently different axillary meristem biology. An orthologous gene has also been discovered in rice [17]. Furthermore, in situ hybridization experiments demonstrate that, even in vegetative Arabidopsis plants, LAS is expressed in the primary shoot apical meristem at the adaxial margins of initiating leaf primordia, where it appears to act to protect primordial cells from differentiation, supporting the reserve meristem hypothesis [16]. Auxin and axillary meristem initiation In addition to contributing to our understanding of the origins of axillary meristems, analysis of Arabidopsis las mutants has revealed a previously unsuspected role for auxin in axillary meristem initiation. Characterization of the auxin-resistant Arabidopsis mutant axr1 had suggested that the effect of auxin on shoot branching is entirely after axillary meristem initiation, through the inhibition of bud out growth [11]. As predicted for a mutant with reduced auxin response, the axr1 mutant is highly branched. However, no histological differences in the timing of axillary meristem initiation could be detected in axr1 mutants, leading to the conclusion that the bushy phenotype was entirely the result of an effect on bud outgrowth [11]. If this were the case, then one would predict that las should be completely epistatic to axr1 because the effect of the axr1 mutation should be entirely after that of the las mutation. However, the axr1 mutation is able weakly to suppress the phenotype conferred by las [16]. In the las single mutant, only 0 –2 side shoots develop from rosette nodes, but in the axr1 las double mutant, up to five such branches were observed. This interesting new role for auxin needs to be investigated further. In the mean time, most attention has focused on the more dramatic later role of auxin in regulating bud outgrowth. Auxin and bud outgrowth After initiation, axillary meristems produce a few leaves to form a bud. The buds can then become dormant or they can continue growth to produce a side shoot. Furthermore, dormant buds can later reactivate to produce a side shoot (see [18] and the Review article by David Horvath et al. in this of Trends in Plant Science [19]). ‘Dormant’ is a somewhat misleading term for the non-growing bud because such buds are highly metabolically active producing a characteristic set of transcripts and proteins. The switch to activity is accompanied by a change in this pattern but,
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within a small time window, activation can be reversed and dormancy can be reimposed, so that axillary buds can cycle between the dormant and active states (see [18] and Horvath et al. in this issue of Trends in Plant Science [19]). It has been known for many decades that auxin travelling in the polar transport stream promotes the dormant state in axillary buds. However, its mode of action is obscure because for almost as long it has been known that auxin does not act directly to suppress bud growth. Direct application of auxin to buds does not inhibit their outgrowth [20] and levels of auxin in buds rise as they activate [21]. Furthermore, apically applied auxin is not transported into the buds [22]. For example, in Arabidopsis, an assay has been developed in which an excised stem segment, bearing a single leaf, is inserted between two agar blocks in a Petri dish [23]. The axillary bud in the leaf axil activates and grows out to produce a side shoot. Addition of auxin to the apical, but not to the basal block, inhibits the outgrowth of the bud. Inhibitors of polar transport block the effect of apical auxin and buds of the axr1 mutant are resistant to the effect of apically applied auxin. However, when radio-labelled auxin is used in this assay, less accumulates in the bud when the auxin is applied apically than when it is applied basally [24]. This strongly supports the idea that auxin acts indirectly, which might be predicted given that to enter a bud, auxin would have to move acropetally from the primary axis into the leaf petiole, against the flow of the polar transport stream. This indirect mode of action for auxin has led to the hypothesis that a second messenger carries the auxin signal into the bud. Several candidates have been proposed, with the strongest contender being cytokinin. Cytokinin can promote the outgrowth of buds when directly applied to them, and cytokinin levels rise in buds as they are activated [3,25]. Furthermore, cytokinin supplied through the basal cut surface of an excised Arabidopsis nodal segment can overcome the inhibitory effects of apically applied auxin on the growth of axillary buds [23]. The link between auxin and cytokinin comes from two sources. First, it has been shown that in bean (Phaseolus vulgaris), decapitation results in an increase in export of cytokinin from roots in the xylem [26]. This increase can be partly blocked by applying auxin to the decapitated stump. Second, it has been shown that auxin can down-regulate cytokinin synthesis [27,18]. These data suggest that one mode of action for auxin could be to down-regulate both local cytokinin synthesis and cytokinin export from roots and that this might prevent the activation of buds. Novel second messengers Although action through cytokinins probably accounts for part of the effect of apical auxin, it appears not to be the only mode of auxin action. There is growing evidence for a second long-range signalling system that interacts with auxin to inhibit shoot branching. In recent years, evidence has accumulated for the involvement of an entirely novel signal in relaying the auxin message into the bud. This has come from the analysis of a series of bushy mutants in pea and Arabidopsis, called the rms and max mutants, respectively [28 – 31]. The mutants are characterized by increased http://plants.trends.com
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branching, reduced stature and in most cases shorter leaves. The axillary buds of all the rms and max mutants tested to date show at least a reduced response to apically applied auxin following decapitation (rms mutants) or in the isolated node assay (max mutants) [29,32]. Thus, these genes are required for a wild-type auxin-mediated inhibition of bud outgrowth. A sub-set of the RMS and MAX genes are excellent candidates for involvement in the production of the predicted auxin second messenger. The highly branched phenotype of mutant shoots of max1, max3, max4, rms1, rms2 and rms5 can be restored to wild type by grafting to wild-type root stocks [29,30,33– 35]. Thus, these genes are probably required for the production of a long-range graft-transmissible, branch-inhibiting signal. Hormonal measurements suggest that the rms1 phenotype is not associated with either reduced auxin or increased cytokinin, implicating a novel factor [32]. Because all the mutants tested to date have auxin-resistant bud growth in decapitation assays, this long-range signal must be required for auxin-mediated apical dominance. Further grafting studies have revealed other properties of this signal [29,30,33– 35] (Figure 2). For example, the reciprocal experiment in which wild-type shoots are grafted to mutant roots also results in wild-type shoot branching. This demonstrates that the signal can be synthesized all along the plant axis, and, just as production in only the root is sufficient to restore wild-type branching, so is production in only the shoot. Small wild-type interstock segments grafted between mutant tissues in the shoot are also sufficient to restore wild-type shoot branching to the shoot system. This suggests that the signal is either potent or can be made in large quantities. The use of a different grafting techniques, so-called Y-grafted, suggests that the signal can only move up plants [30,34]. In these experiments, a wild-type shoot is grafted into the root– shoot junction of a mutant plant, but the mutant shoot is not restored to wild-type, indicating that the signal cannot move down the plant from the wild-type shoot and then back up to restore a wild-type branching habit to the mutant shoot system. What then is this potent upwardly mobile signal? Some hints have come from the molecular analysis of the MAX4 and RMS1 genes, which encode orthologous members of the carotenoid cleaving dioxygenase family [29]. The MAX4 and RMS1 proteins are predicted to be plastidic because they contain potential plastid-targeting sequences. Plastids are the site of carotenoid synthesis in plants. Hence, an attractive hypothesis is that the longrange branch-inhibiting signal is derived by cleavage of a carotenoid. However, other members of this diverse family have non-carotenoid substrates and some do not catalyse cleavage reactions and instead are involved in isomerization [36]. However, if the hypothesis were correct it would add to the growing number of plant growth regulating compounds derived from carotenoids. Apart from the wellcharacterized hormone abscisic acid, there is some evidence that b-ionone, a carotenoid cleavage product, has hormonal activity [37,38]. The molecular characterization of the MAX4 and RMS1 genes has also allowed an investigation of the interaction
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Figure 2. Summary of the results of grafting experiments between the rms1, rms2 and rms5 mutants (green) of pea with their wild types (purple) and the max1, max3 and max4 mutants (green) of Arabidopsis with their wild type (purple). Self grafts of the wild types have wild-type branching patterns (a) and self grafts of the mutants result in typical mutant bushy plants (b). Wild-type roots can restore wild-type branching to mutant shoots (c) and wild-type shoots show wild-type branching patterns when grafted to mutant roots (d). For the genotypes tested to date, small interstock wild-type grafts restore wild-type branching to nodes above the graft (e).
of auxin and the MAX/RMS-dependent signal [29]. Interestingly, the pea and Arabidopsis systems diverge here. In pea, there is evidence that auxin can rapidly up-regulate the transcription of RMS1 in the stem. In this case, auxin could inhibit bud growth by locally increasing the amount of RMS1, and hence presumably the product of the reaction it catalyses. By contrast, neither PCR-based approaches nor promoter-GUS reporter constructs have shown evidence for auxin regulating the transcription of MAX4 at the node. Instead, some auxin-induced MAX4 expression was observed behind the root tip, but this only occurs 24 h after auxin treatment. Hence, in Arabidopsis, a post-transcriptional mode of action for auxin in regulating the MAX pathway is likely. Transcription of MAX4 does not appear to be a limiting step in branch inhibition because plants overexpressing MAX4 from the cauliflower mosaic virus 35S promoter are as branched as wild-type plants are. Once synthesized, the mechanism of action by which the MAX/RMS-dependent signal inhibits buds is unknown. Some clues come from the molecular analysis of the MAX2 gene. The max2 mutants are phenotypically near identical to the max1, max3 and max4 mutants. However, the max2 shoot phenotype cannot be restored to wild type by grafting to wild-type roots (J. Booker, C. Turnbull and O. Leyser, unpublished). Therefore, MAX2 acts in the shoot to regulate branching and could be involved in the perception of the MAX-dependent signal. Similar conclusions have been drawn from grafting experiments carried out with the rms3 and rms4 mutants [39,28]. Consistent with this idea, MAX2 encodes an F-box protein. F-box proteins are involved in the selection of specific substrates for ubiquitin-mediated degradation [40]. There are ,700 F-box proteins in the Arabidopsis genome [41], and several of the better-characterized family members have roles in hormone signalling [40]. For example, an F-box protein, TIR1, mediates the auxin-induced degradation of the members of the Aux/IAA family of transcriptional repressors, thus controlling auxin-induced gene expression [42]. http://plants.trends.com
A similar system could operate in the transduction of the MAX/RMS-dependent signal. Conclusions It was first shown in 1933 that apically derived auxin could inhibit axillary bud growth. More than 70 years later the mechanism by which auxin regulates shoot branching is still unknown. Recent results show that auxin can influence various stages of bud development, from a minor role in bud initiation through to a major but indirect effect on bud activity involving at least two different hormonal second messengers. It will be particularly exciting to find out the identity of the novel unknown signal that acts downstream of auxin. In the future, the hormonal control of shoot branching should provide an excellent model for the study of the integration of environmental and genetic controls in plant development. Here, the comparison of the regulation of branching in different species with different regulatory balances should be particularly informative, and already RMS1 and MAX4 provide an example of orthologous genes that are regulated differently. These recent advances have come from the powerful combination of physiological, genetic, biochemical and genomic resources, which are now available to researchers in this field, promising rapid continued progress. References 1 Thimann, K. and Skook, F. (1933) Studies on the growth hormone of plants III: the inhibitory action of the growth substance on bud development. Proc. Natl. Acad. Sci. U. S. A. 19, 714 – 716 2 Ljung, K. et al. (2001) Sites and homeostatic control of auxin biosynthesis in Arabidopsis during vegetative growth. Plant J. 28, 465 – 474 3 Cline, M.G. (1991) Apical dominance. Bot. Rev. 57, 318 – 358 4 Garrison, R. (1955) Studies in the development of axillary buds. Am. J. Bot. 42, 257 – 266 5 Sussex, I.M. (1955) Morphogenesis in Solanum tuberosum L.: experimental investigation of leaf dorsoventrality and orientation in the juvenile shoot. Phytomorphology 5, 286 – 300 6 Snow, M. and Snow, R. (1942) The determination of axillary buds. New Phytol. 41, 13 – 22
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7 Johri, M.M. and Coe, E.H. (1983) Clonal analysis of corn plant development. 1. The development of the tassel and the ear shoot. Dev. Biol. 97, 154 – 172 8 McDaniel, C.N. and Poethig, R.S. (1988) Cell-lineage patterns in the shoot apical meristem of the germinating maize embryo. Planta 175, 13 – 22 9 Long, J. and Barton, M.K. (2000) Initiation of axillary and floral meristems in Arabidopsis. Dev. Biol. 218, 341 – 353 10 Gribic, V. and Bleecker, A.B. (2000) Axillary meristem development in Arabidopsis thaliana. Plant J. 21, 215 – 223 11 Stirnberg, P. et al. (1999) AXR1 acts after lateral bud formation to inhibit lateral bud growth in Arabidopsis. Plant Physiol. 121, 839– 847 12 Irish, V.F. and Sussex, I.M. (1992) A fate map of the Arabidopsis embryonic shoot apical meristem. Development 115, 745 – 753 13 Furner, I.J. and Pumfrey, J.E. (1992) Cell fate in the shoot apical meristem of Arabidopsis thaliana. Development 115, 755– 764 14 Schmitz, G. and Theres, K. (1999) Genetic control of branching in Arabidopsis and tomato. Curr. Opin. Plant Biol. 2, 51– 55 15 Schumacher, K. et al. (1999) The Lateral suppressor (Ls) gene of tomato encodes a new member of the VHIID protein family. Proc. Natl. Acad. Sci. U. S. A. 96, 290 – 295 16 Greb, T. et al. (2003) Molecular analysis of the LATERAL SUPPRESSOR gene in Arabidopsis reveals a conserved control mechanism for axillary meristem formation. Genes Dev. 17, 1175– 1187 17 Li, X. et al. (2003) Control of tillering in rice. Nature 422, 618– 621 18 Shimizu-Sato, S. and Mori, H. (2001) Control in outgrowth and dormancy in axillary buds. Plant Physiol. 127, 1405 – 1413 19 Horvarth, D.P. et al. (2003) Knowing when to grow: signals regulating bud dormancy. Trends Plant Sci. 8(11) (2003), this issue. doi:10.1016/ j.tplants.2003.09.013 20 Cline, M.G. (1996) Exogenous auxin effects on lateral bud outgrowth in decapitated shoots. Ann. Bot. 78, 255 – 266 21 Hillman, J.R. et al. (1977) Apical dominance and levels of IAA in Phaseolus lateral buds. Planta 134, 191 – 193 22 Morris, D.A. (1977) Transport of exogenous auxin in two-branched dwarf pea seedlings (Pisum sativum L.). Planta 136, 91 – 96 23 Chatfield, S.P. et al. (2000) The hormonal regulation of axillary bud growth in Arabidopsis. Plant J. 24, 159 – 169 24 Booker, J. et al. (2003) Auxin acts in xylem associated/medullary cells to mediate apical dominance. Plant Cell 15, 495 – 507 25 Turnbull, C.N.G. et al. (1997) Rapid increase in cytokinin concentration in lateral buds of chick pea (Cicer anerinum) during release of apical dominance. Planta 202, 271 – 276
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26 Bangerth, F. (1994) Response of cytokinin concentration in xylem exude of beans (Phaseolus vulgaris L.) plants to decapitation and auxin treatment and relationship to apical dominance. Planta 194, 439– 442 27 Eklof, S. et al. (1997) Auxin – cytokinin interactions in wild-type and transgenic tobacco. Plant Cell Physiol. 38, 225 – 235 28 Beveridge, C.A. (2000) Long distance signalling and a mutational analysis of branching in pea. Plant Growth Regul. 32, 193 – 203 29 Sorefan, K. et al. (2003) MAX4 and RMS1 are orthologous dioxygenase-like genes that regulate shoot branching in Arabidopsis and pea. Genes Dev. 17, 1469 – 1474 30 Turnbull, C.N.G. et al. (2002) Micrografting techniques for testing long distance signalling in Arabidopsis. Plant J. 32, 255 – 262 31 Stirnberg, P. et al. (2002) MAX1 and MAX2 control shoot lateral branching in Arabidopsis. Development 129, 1131 – 1141 32 Beveridge, C. et al. (2000) Auxin inhibition of decapitation-induced branching is dependent of graft-transmissible signals regulated by genes rms1 and rms2. Plant Physiol. 123, 689– 697 33 Beveridge, C.A. et al. (1997) RMS mutant has increased IAA and decreased root sap zeatin riboside content but increased branching, controlled by graft transmissible signal(s). Plant Physiol. 115, 1251– 1258 34 Foo, E. et al. (2001) Long distance signalling and the control of branching in the rms1 mutant of pea. Plant Physiol. 126, 203 – 209 35 Morris, S.E. et al. (2001) Mutational analysis of branching in pea (Pisum sativum L.): evidence that Rms1 and Rms5 regulate the same novel signal. Plant Physiol. 126, 1205– 1213 36 Schwartz, S.H. et al. (2001) Characterization of a novel carotenoid cleavage dioxygenase from plants. J. Biol. Chem. 276, 25208 – 25211 37 Redmond, T.M. et al. (1998) Rpe65 is necessary for the production of 11-cis-vitamin A in the retinal visual cycle. Nat. Genet. 20, 344– 351 38 Katonoguchi, H. (1994) Occurrence of a growth inhibitor, 3-hydroxybeta-ionone, in 7 cultivars of Phaseolus vulgaris and its role in lightinduced growth-inhibition. Phytochemistry 36, 273 – 275 39 Beveridge, C.A. et al. (1996) Branching in pea – action of genes rms3 and rms4. Plant Physiol. 110, 859 – 865 40 Hellmann, H. and Estelle, M. (2002) Plant development: regulation by protein degradation. Science 297, 793 – 797 41 Gagne, J.M. et al. (2002) The F-box subunit of the SCF E3 complex is encoded by a diverse superfamily of genes in Arabidopsis. Proc. Natl. Acad. Sci. U. S. A. 99, 11519 – 11524 42 Gray, W.M. et al. (2001) Auxin regulates SCFTIR1-dependent degradation of Aux/IAA proteins. Nature 414, 271 – 276
Free journals for developing countries The WHO and six medical journal publishers have launched the Access to Research Initiative, which enables ~70 developing countries to gain free access to biomedical literature through the Internet. The science publishers, Blackwell, Elsevier Science, the Harcourt Worldwide STM group, Wolters Kluwer International Health and Science, Springer-Verlag and John Wiley, were approached by the WHO and the British Medical Journal in 2001. Initially, >1000 journals will be available for free or at significantly reduced prices to universities, medical schools, research and public institutions in developing countries. The second stage involves extending this initiative to institutions in other countries. Gro Harlem Brundtland, director-general for the WHO, said that this initiative was ‘perhaps the biggest step ever taken towards reducing the health information gap between rich and poor countries’. See http://www.healthinternetwork.net for more information. http://plants.trends.com
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