Signal transduction in plant cells

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Signal transducti0n in C e r t a i n critical features of the plant lifestyle are relevant to understanding molecular signalling in plants. Probably the primary one is the lack of obvious movement. Coordination between tissues and cells is much less obvious in plants than in animals, and plants also lack the pronounced differentiation between sensory and responding cells and tissues found in mammals. Some plant tissues, such as stomata and pulvini (the leaf stalk), do show limited movement, and signalling between the major parts of plants can also be detected, but parallels with nerve-muscle coordination or longrange coordination by specific hormones are not easily made. The signals - between root or shoot, for example - are usually a complex mixture reflecting the major functions of these tissues and the different environments in which they grow. Plant growth and development may be considered the equivalent of movement. Growing plants are permanently embryogenic, and many plant cells in the mature plant body remain unspecialized, accounting for their remarkable regenerative capabilities. Plants can grow rather than move into the more equable parts of their local environment. The developmental plasticity of higher plants 1 necessitates an accurate sensing and integration of the incoming direction of environmental 1.00 .(a) o-o-43--o--o,

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PTGll Variation in [Ca2+]iwith stomatal aperture as the Ca2+ concentration is artificiallyelevated in single guard cells. Measurement of fluorescent light emission from a fluorescent Ca2+ indicator can be used to determine [Ca2+]i.In (b) and (c), respectively, IP3 or Ca2÷were released from chemically caged forms by 10 s ultraviolet photolysis at arrow. (a) Unphotolysed control. The aperture is half the width between two guard cells. Further details are in Ref. 18.

plant cells A. TREWAVASAND S. GIlllOY

In plants, unUke animals, signal transduction studies are in their infancy. While intracellular Ca2+appears to have second messenger functions, attempts to show that protein kinases, tnositol phosphates and cyclic AMP are involved in signal transduction in plants have run into considerable di~cu~ty. signals such as light (of various wavelengths), gravity, wind, minerals, water, gases and soil structure 2. Each growing part has a semiautonomous capability both to perceive and to act upon the information that impinges upon it. Indeed individual plant cells (e.g. guard cells) can both directly sense and respond to environmental stimuli. This external information must be transduced and integrated with internal signals and with a continuously changing cell and tissue ontogeny to generate optimal growth and development patterns, but how plants manage this difficult feat is only dimly perceived. This short article outlines our present state of knowledge concerning signal transduction systems in plants. Understanding is rapidly accelerating, and a considerably clearer picture seems certain to emerge within the next few years. CaZ+-based signal transduction All the elements of a Ca2+-based transduction system are found in plant cells. The evidence may be simply listed. (1) Many measurements have shown that the basal concentration of intracellular calcium, [Ca2+]i, in different plant cells is less than 200 nM3,4, and in a limited number of cases this level has been shown to increase upon stimulation of the cell (see below). (2) Calmodulin has been isolated from numerous plant sources, and the gene cloned and sequenced 5,6. There is a strong conservation of sequence, trypsin lability and structure between plant and animal calmodulins. Calmodulin is found in both membrane and soluble fractions and the plasma membrane is particularly enriched in an EGTA-stable, bound state 7. Other Ca2+-binding proteins have also been detected 8. (3) A number of Ca2+/calmodulin-dependent enzymes have been detected, including protein kinase, Ca 2+ ATPase and NAD kinase 8. Ca 2+ has also been reported to regulate enzymes involved in turgor generation, sucrose synthesis and cytoskeletal structures 9,1°. (4) The presence of Ca 2+ channels has been established in both lower and higher plants n. There is evidence that both phosphorylation and inositol trisphosphate (IP,) modify channel activity, and recently the plant growth substance abscisic acid has been shown to activate channels, allowing Ca 2+ influx to the cytosol across the plant plasma membrane 12. (5) Cae+ ATPases have been characterized in the membrane of a number of plant cells4. A calmodulinstimulated ATPase has been described which shows sequence and structural conservation with erythrocyte

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FIGIi Variation of intraceUularCa2÷ concentration in a single guard cell 0, 2, 5, 10 and 20 min after closure was initiated by lowering the intracellular K÷ concentration. Different [Ca2+]i measured by fluorescence ratio imaging are shown as different colours. Detectable aperture closure starts at about 10 min. Further details are in Ref. 21. Ca 2+ ATPase and may be located in the plasma membrane. Accumulation of Ca 2+ into numerous plant organelles has been observed, with the vacuole probably being the major intracellular storeg. (6) Proteins binding to GTP with strong affinity and specificity have been reported to be present in membranes of several plant cells 13,14, and there is evidence that GTP can cause Ca 2÷ release from membrane vesicles 15. GTP-binding proteins may therefore regulate Ca 2+ movement. Proteins from several plants cross-react with antibodies to the ct subunit of mammalian G proteins and a gene for this protein has been found in Arabidopsis thaliana 16. However, we are still far from defining a role for G proteins in signalling in plant cells.

Evidence that changes in [Ca2+]itransduce signals Many studies have used calmodulin-binding inhibitors, channel blockers and other inhibitors of Ca 2+ transport to demonstrate that light, gravity or growth regulator signals are transduced through [Ca2+]i (Refs 8, 9, 17). Direct corroboration by measurement of [Ca2+]i has lagged far behind. The development of a range of fluorescent Ca 2+ indicators by Tsien and colleagues has made detection of changes in [Ca2+]i relatively straightforward in animal cells. However, a series of often crippling problems has emerged with their use in plants. There are serious difficulties with dye loading, organelle localization and leakage. Consequently, very few reliable measurements

of [Ca2+]i have been published. Fluorescent dyes have been used to observe increases in [Ca2+]i during stomatal closure (Figs 1, 2) and pollen tube growthlS, 19. Ca 2+sensitive microelectrodes have revealed an auxininduced [Ca2+]i change and a localized high [Ca2+]i in rhizoid tips of Fucus 19. Inhibition of cytoplasmic streaming is accompanied by increased [Ca2+]i (Ref. 9). An entirely new procedure for measuring [Ca2+]i is based on the expression of transgenic aequorin. This Ca2+-dependent luminescent protein is cytoplasmic, nontoxic and can potentially be expressed in any viable cell of a transgenic plant. Indeed, Ca2+-induced luminescence can be measured from whole seedlings 2°. Unlike the fluorescent dyes, transgenic aequorin requires no complex uptake protocol and is not relocalized within the cytoplasm. Use of this procedure has shown that [Ca2+li in whole tobacco seedlings is increased by much, cold shock, fungal elicitors and salination 2°. Calmodulin gene expression is also greatly increased by touch 6. Even though a range of signals elevate [Ca2+]i, the problems have not yet diminished. Elevation of [Ca2+]i in guard cells, either by release of Ca 2+ from a chemically caged form inside the cell (Fig. 1), or by modifying extracellular Ca 2+ or K+ (Fig. 2), initiates stomatal closure 21. However, when closure is initiated by the growth regulator abscisic acid, the change in [Ca2+]i is highly variable, ranging from no change to a sustained increase with possible superimposed oscillations.

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]REVIEWS Similar variability is observed in the Ca 2+ channel activity of the plasma membrane in response to abscisic acid 12. These results are just indications of the complex responses we can expect from a Ca2+-based signal transduction system in plant cells.

proteins, ions, Ca2+-binding constituents and outwardly facing plasma membrane phospholipids will contribute to [Ca2+]w control. The regulation of free [Ca2+lw is clearly a critical area for further study.

Signal-responsive protein ktnase in plant cells Ca2+homeostasis and regulation in the cell wall The possession of a wall is one of the more obvious ways in which plant cells differ from the typical animal cell. The composition, structure and stiffness of the wall change during cell development. There is a constant interchange of information between the wall and the cell, and if the cell wall is damaged, for example by pathogen-produced cell wall hydrolases, the wall fragments induce host cell defence responses. Ca 2+ is a major structural constituent of the wall, where it is bound (often strongly) by polygalacturonic acid (pectic acid). Until recently it was assumed that wall Ca >, [Ca>lw, only functioned to maintain cell wall stiffness, but this view is changing. The cell maintains its [Ca2+]w between narrow limits and adjusts this value when growth and development changes. Cell walls even contain calmodulin. There is now a growing number of observations indicating that the concentration of free [Ca2+]w may act to regulate both growth and development. Exogenous Ca 2+, [Ca2+]e, inhibits shoot growth at concentrations below 0.1 mM. When shoots are gravitropically stimulated to bend, [Ca2+]w increases in shoot regions where growth is inhibited and, significantly, accumulation precedes bending 8,9. Similar growthinhibiting accumulations occur during gravitropic bending of roots. Epidermal cells, which are believed to constrain and regulate shoot growth, have a higher [cae+lw. Lowering of epidermal [Cae+]~,. during incubation of tissues in water may induce anomalous growth changes. Loss of [Ca>]w may be the primary event in both fruit ripening and leaf abscission. When plant tissues are fed Ca 2+, it accumulates in specialized cells, often as calcium oxalate crystals in vacuoles. (The shape of the crystal is, intriguingly, genetically determined and can be used for taxonomic purposes.) The number of these cells is determined by the amount of Ca 2+ provided 22. If tissues are placed in medium free of Ca 2+, this internally accumulated Ca e+ is lost in a few hours. Since the loss occurs first from the wall, the cell perceives the altered level of wall Ca 2+ and pumps out Ca 2+ to compensate. In insectivorous pitcher plants the pitcher liquid is maintained at about 0.01 mM Ca 2+ (Ref. 23); if Ca 2+ is added to the liquid the concentration decreases, if taken away it increases. Mobile [Ca2+],,. may also act to signal stress responses. Stomata can be closed by elevating [Ca2*]e (Ref. 21) and a release of Ca 2+ from stressed root and leaf cells may act as a physiologically relevant message. What is the concentration of free [Ca2+]w? When protoplasts were titrated with [Ca2+]e, from 0.01 gM to 1 mM, the [Ca2+]i increased only threefold between external Ca 2+ concentrations of 0.1 and 1 ~tM, and was thereafter constant 2q. On this basis free [Ca2+lw may be 1 btM or less. The concentration of free [Ca2+]w will be dependent on wall pH, which in turn is determined by ontogeny and stress and uptake of Ca 2+ from the wall into the cell. Other wall materials, polysaccharides and

Protein kinases are ubiquitous elements in signal transduction. The first reports of protein kinase, in plant cells go back nearly 20 years but the last half decade has seen a real acceleration of interest1°, 25,26. Ca2+-dependent protein kinase was first detected in plants in 198227. Many enzymes and proteins in plants are now known to be phosphorylated, including ATPase, RNA polymerase, phosphoenolpyruvate carboxylase, malate synthase, pyruvate dehydrogenase, sucrose phosphate synthase, histone H1 and numerous chloroplast proteins. The activity of many of these proteins has been shown to be modulated by phosphorylation 25. Phosphorylated proteins are found in all cell fractions. Thus, the metabolic system of plant cells is as dependent upon phosphorylation as the equivalent animal cell. Direct mediation of signalling by protein kinase has been established in a number of significant examples 10. (1) Tobacco mosaic virus stimulates phosphorylation of host-encoded protein via a protein kinase stimulated by double-stranded RNA; enzyme activity peaks during replication of plant viruses or viroids. (2) Phytochrome, a well-characterized and primary light receptor, either is a protein kinase or is intimately associated with kinase activity. Phytochrome has an ATP-binding site, and conversion between the active (PFR) and inactive (PR) forms modifies its apparent autophosphorylation sites. (3) Blue light modifies the phosphorylation of a single plasma membrane protein, which is probably itself a protein kinase and may be the blue light receptor. (4) Fungal elicitors modify the phosphorylation of a single plasma membrane protein in vitro. (5) A membrane-spanning protein kinase has been cloned in which the extracellular domain is very similar to the Brassica S locus incompatibility glycoprotein2~. This protein kinase may represent the initial transduction step of signalling between pollen and stigma which destroys incompatible pollen. Other protein kinases responding to heat shock or pH have been detected. However, our understanding of the role of such kinases in the transduction of these signals awaits identification of their substrates in vivo. Regulation of metabolism by phosphorylation also requires an active phosphatase to reverse the effects induced by protein kinase action. Specific protein phosphatases types 1 and 2A have been detected in plant cells 29, although once again their substrates in vivo remain largely unknown. This is a burgeoning research field, and real and exciting progress in many different areas of metabolic regulation is anticipated.

lnositol lipids and phospholipases Considerable attention has been focused on plant inositol phospholipids as potential sources of second messengers such as IP3 and diglyceride ~<3°. Although the picture is now becoming clearer, there have been considerable difficulties. A primary problem has been the extremely low level of detectable phosphatidyl-

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Is cyclic AMP a messenger in plant cells? The first attempts to demonstrate a functional role for cyclic AMP in plant ceils started in the late 1960s and early 19"v0s,~,~. M'ter an initial flush ()f enthusiasnl and some not very careful work, interest all but disappeared. The field fl~undcred on two (tifficuhies: an apparent lack of a biologically reasonable level of cAMP (a rectlrring problem, as evidenced in recent studies ~) and the difficuhy of detecting a cAMP-dependent prorein kinase. Recent molecular approaches suggest that both cAMP-binding proteins and a cAMP-dependent protein kinase may, however, be present ~s,:<. Plant cAMP has retained a small coterie of supporters 33 against a rather larger n u n l l ) e r o f convinccd sceptics~L Neither side has, in ()ur view, yet proved its case, Reporls of detection of adenylyl cyclase by

TIG NOVEMBI!R/I)ECI~IM~ER1991 vol.. 7 N(). t 1,12

r~EVIEWS protagonists have not been followed up by exacting purification but, on the other hand, the antagonists have not appreciated the possibilities of transduction system adaptation. The difficulties attending the demonstration of IP 3 in plant cells should act as a cautionary tale and, indeed, there are remarkable similarities between the problems of detection of both IP 3 and cAMP in plant cells. If, for example, cAMP were confined to guard cells, leaf homogenates would contain only tiny amounts, in agreement with observation. Cyclic AMP is definitely present in algae33; it may just be proving coy in higher plants.

What does the future hold? Within the space of five or six years, plant cell signal transduction has grown from a relatively obscure study involving some dozen or so scientists to a rapidly accelerating area involving many hundreds of individuals. Their endeavour has highlighted at least one unusual characteristic of plant development: an apparent redundancy in the regulating signals 1. Many different chemical and environmental signals can elicit the same physiological response. Plants are sedentary autotrophs, collecting their resources from a local but varying environment, and they modify their development to exploit the changing circumstances. Redundancy in signalling is essential to this lifestyle and helps provide fail-safe possibilities. An equivalent redundancy in transduction pathways now seems likely. Artificial elevation of guard cell a variety of means (Fig. 1) causes stomata to close, but increased [Ca2+]iis not obligatory for closure even though Ca 2÷ channel blockers and calmodulin-binding inhibitors can impede the functioning of closing signals. As knowledge of transduction pathways and their numbers increases so will the apparent complexities of pathway interaction. Such observations will challenge our concepts of causality and question the oversimplified perception/transduction schemes quoted in many textbooks. Understanding can be sought in network or systems behaviour, as already described for plant development 1, but an open mind is going to be essential. Transduction chain redundancy may also help to explain the difficulties with IP3 and cAMP noted previously. The presence of an unstirred layer of free wall Ca 2+ adjacent to but outside the plasma membrane has no precedent in mammalian cells. Ca 2+ has a constrained mobility in both the cell wall and the cytoplasm. Spatial inhomogeneities in [Ca2+]ishould induce equivalent adjacent 'islands' of free [Ca2+]w which in turn should generate inhomogeneities in the [Ca2+]i of adjacent cells; these fluctuations could modify the cytoskeleton and thus transmit [Ca2+]i inhomogeneities onwards to the cell wall and cytoplasm of other cells. This mechanism may explain how plants respond to gradients of gravity, light or pressure. Perception of these signals is known to be limited to a few cells in shoots or roots but growth responses occur throughout extensive regions of these tissues. The 'to and fro' transfer of Ca 2+ between wall and cytoplasm provides a simple mechanism for building spatially defined gradients or bands of Ca 2÷ well beyond perception sites. [Ca2÷l~ can thus form the basis of a simple extracellular signalling/transmission system perhaps analogous to

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Dictyostelium, which uses cAMP, normally an intracellular messenger, as an extracellular aggregating signal. Pattern specification, cell differentiation and tissue morphogenesis are areas of research that will advance dramatically when the technology for imaging free wall Ca 2+ in living tissues becomes available. With present technological progress 2° this is not far off.

References 1 Trewavas, A.J. (1986) in Plasticity in Plants (Jennings, D.H. and Trewavas, A.J., eds), pp. 31-77, Company of Biologists 2 Gilroy, S. and Trewavas, A.J. (1990) in The Plant Plasma Membrane (Larsson, C. and Moiler, J.M., eds), pp. 204-232, Springer-Verlag 3 Busa, D.S. and Jones, R.L. (1990) Plant Physiol. 93, 841-845 4 Evans, D.E., Briars, S,A. and Williams, L.E. (1991)J. Exp. Bot. 42, 285-303 5 Ling, V. and Zielinski, R.E. (1989) Plant Physiol. 90, 714-719 6 Braam, J. and Davies, R.W. (1990) Cell60, 357-364 7 Collinge, M. and Trewavas, A.J. (1989)J. Biol. Chem. 264, 8865---8872 8 Allan, E. and Trewavas, A.J. (1987) in The Biochemistry of Plants: A Comprehensive Treatise (Vol. 12) (Stumpf, P.K. and Conn, E.E., eds), pp. 117-153, Academic Press 9 Hepler, P.K. and Wayne, R.O. (1985) Annu. Rev. Plant Physiol. 36, 397-436 10 Randall, D.D. and Blevins, D.G. (1990) Curt. Top. Plant Biocbem. Physiol. 9, 1-433 11 Tester, M. (1990) NewPhytol. 114, 305-340 12 Schroeder, J. and Hagiwara, W. (1990) Proc. NatlAcad. Sci. USA 87, 9305-9309 13 Jacobs, M. et al. (1988) Biochem. Biophys. Res. Commun. 155, 1478-1484 14 Einspahr, K.J. and Thompson, G.A. (1990) Plant Physiol. 93, 361-366 15 Allan, E., Dawson, A., Drobak, B. and Roberts, K. (1989) Cell Signalling 1, 29-39 I 6 Ma, H., Yanofsky, M.F. and Meyerowitz, E.M. (1990) Proc. Natl Acad. Sci. USA 87, 3821-3825 17 Trewavas, A.J. (1986) Molecular and Cellular Aspects of Calcium in Plant Development, Plenum Press 18 Gilroy, S., Read, N.D. and Trewavas, A.J. (1990) Nature 346, 769-771 19 Hepler, P. (1990) Curr. Top. Plant Biochem. Physiol. 9, 1-10 20 Knight, M., Campbell, A., Smith, S.M. and Trewavas, A.J. (1991) Nature 352, 524-526 21 Gilroy, S., Fricker, M.D., Read, N.D. and Trewavas, A.J. (1991) Plant Cell 3, 333--344 22 Borchert, K (1990) Planta 182, 339-347 23 Meir, P., Juniper, B.E. and Evans, D.E. Ann. Bot. (in press) 24 Gilroy, S., Hughes, W.A. and Trewavas, A.J. (1989) Plant Physiol. 90, 482-491 25 Ranjeva, R. and Boudet, A.M. (1987) Annu. Rev. P l a n t Physiol. 38, 73-93 26 Budde, R.J.A. and Chollet, R. (1988) Physiol. Plant. 72, 435-439 27 Hetherington, A. and Trewavas, A.J. (1982) FEBSLett. 145, 67-71 28 Walker, J.C. and Zhang, R. (1990) Nature 345, 743-746 29 MacKintosh, C. and Cohen, P. (1989) Biochem.J. 262, 335-339 30 Boss, W.F., Memon, A.R. and Chen, Q. (1989) in Signal Perception and Transduction in Higber Plants (Ranjeva, R. and Boudet, A.M., eds), pp. 161-184, Springer-Verlag

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[~EVIEWS 31 Sanders, D. et al. (1990) Curr. Top. Plant Biochem. Physiol. 9, 20-37 32 Gilroy, S. et al. (1989) in Plant Membrane Transport: The Current Position (Dainty, J., de Michaelis, M.I., Marre, E.

and Rasi-Caldogno, F., eds), pp. 215-225, Elsevier 33 Newton, R.P. and Brown, E.G. (1986) in Hormones, Receptors and Cellular Interaction in Plants (Chadwick, C.M. and Garrod, D.R., eds), pp. 115-154, Cambridge University Press 34 Spiteri, A. et al. (1989) Plant Physiol. 91,624-628

35 Biermann, B., Johnson, E.M, and Feldman, L.J. (1990) Plant Physiol. 94, 1609-1615 36 Kabagiri, F., Lain, E. and Chua, N-H. (1989) Nature 340,

727-730 A. TREWAVAS IS IN THE INSTITUTE OF CELL AND MOLECULAR BIOLOGY, UNIVERSflY OF EDINBURGH, EDINBURGH E H 9 ~]H, U K AND S~ GILROY IS IN THE DEPARTMENT OF PLANT BIOLOGY,

UNWE~rrY OFCAUFORNL4,BERgELEY,CA 94720, USA.

C e l l interactions direct morphogenesis, cell movement, differentiation and developmental synchrony during fruiting body development of the prokaryote Myxococcus xanthus. Although Myxococcus uses as many as four different intercellular signals during development, this article focuses on one of them, the C factor. The aim is to show how genetic, biochemical and cell mosaic analyses have deepened our understanding of C factor function. Studies of intercellular signaling mechanisms in M. x a n t h u s benefit from the prokaryotic structure and genetics of this organism 1. The growing cells of M. x a n t h u s (Fig. 1, O h) are long, thin Gram-negative rods, 0.5 I.tm by 5 tim, that normally feed in moving contiguous swarms by secreting hydrolytic enzymes that degrade polymeric organic substrates in soil. The rate of growth on a protein substrate increases with cell density, indicating that feeding is cooperative 2. Movement is also cooperative; certain mutant classes move only when cells are close together3, suggesting coordinated signaling between cells. Myxobacteria have a genome of 9500 kb of unique DNA4, only twice the size of the E. coli genome, and are amenable to genetic transduction, integrative plasmid transfer from E. coli, transposon mutagenesis and other genetic techniques applicable to Gram-negative bacteria (reviewed in Ref. 5). Cells may be grown to high density in dispersed culture, have a generation time of 4 h, and mosaic cultures can be formed simply by mixing two genetically distinct strains. When starved for amino acids at high cell density on a solid surface, M. x a n t h u s cells form aggregation centers (Fig. 1, 4 h). The number and density of cells increase at these centers until a mound forms (8 and 12 h), containing about 100000 densely packed cells6. Early in aggregation, ridge-like accumulations of cells (not shown in Fig. 1) move rhythmically and coordinately over the substrate, like ripples on a water surface 7,8. Within a nascent fruiting body the cells differentiate into dormant ovoid spores 9 (Fig. 1). A dense mound of spores about 0.1 mm high, called a fruiting body, forms about 24 h after the removal of nutrient. This orderly change in morphology from an initially undifferentiated sheet of starving cells is also reflected in the temporal pattern of developmental gene expression (Fig. 2) l°,n.

Cell-cell interactions and the developmental program Control of M. x a n t b u s fruiting body development by sequential cell-cell interactions is implied by the

Intercellular signaling in Myxococcusdevelopment: The role of C factor SEUNG K. KIM Cell communicationgoverns differentiation and morphogenesis infruiting body formation by Myxococcus xanthus. Transmission of a small short-range intercellular sigBal by a protein called Cfactor directs multiceUular pattern formation and coordinates the timing of major developmental events. existence of four classes of nonautonomous developmental mutants 12. These mutants were isolated in a genetic screen for strains that were developmentally defective but could be rescued by addition of wildtype cells. The idea was that wild-type cells could provide an extracellular signal that would allow mutants defective in production of that signal to develop normally. In all, 51 mutants were isolated that had a reduced capacity to form spores, but whose sporulation was rescued by mixing with wild-type cells. In addition, some mutant strains were found to produce extracellular factors that could complement other mutant strains for defects in sporulation and fruiting body formation. Other pairs of mutants failed to complement each other extracellularly in cell mixtures and were therefore assigned to the same group. Four extracellular complementation groups - A, B, C and D were identified in this manner. Linked insertions of the Tn5 transposon were used to map mutations belonging to each class of nonautonomous mutants 13. The different groups have mutations in different loci. Group C loci are called csg (for C signal) and groups A, B and D loci are called asg, bsg and dsg, respectively (Table 1). In each group of mutants, development is arrested at a different time and the pattern of developmentally regulated gene expression (monitored by measuring 13-galactosidase activity produced from the lacZ gene fused to developmentally regulated promoters 14) is modified in a different way. The gene expression that is disrupted in asg, bsg and csg mutants is rescued by codevelopment with wild-type cells 14-16. The nature of the intercellular signal defined by the csg class of mutants is the best understood. All members

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