Patterning of development in Dictyostelium discoideum: factors regulating growth, differentiation, spore dormancy, and germination.

Patterning of development in Dictyostelium discoideum: factors regulating growth, differentiation, spore dormancy, and germination DAVIDA. COTTER,TODDW. SANDS,AND KIRANJ . VIRDY Department of Biology, University of Windsor, Windsor, Ont., Canada N9B 3P4 MICHAELJ. NORTH Department of Biological and Molecular Sciences, School of Natural Sciences, University of Stirling, Stirling FK9 4LA, Scotland

Biochem. Cell Biol. 1992.70:892-919. Downloaded from www.nrcresearchpress.com by Calif Dig Lib - Davis on 01/04/15. For personal use only.

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GERARDKLEINAND MICHELSATRE Laboratoire de Biologie Cellulaire (URA 1130 CNRS), DPpartement de Biologie MolPculaire et Structurale, Centre d J ~ t u d eNuclPaires s de Grenoble, 38041 Genoble CEDEX, France Received April 6 , 1992 D. A., SANDS, T. W., VIRDY,K. J., NORTH,M. J., KLEIN,G., and SATRE,M. 1992. Patterning of development COT~ER, in Dictyostelium discoideum: factors regulating growth, differentiation, spore dormancy, and germination. Biochem. Cell Biol. 70: 892-919. Cellular communication dictates all stages of growth and development in the cellular slime molds. Dictyostelium discoideum utilizes a number of signal molecules for cell-to-cell communication, including growth and density factors, CAMP, ammonia, differentiation-inducing factor, discadenine, and spore autoactivator. A source and sink model is presented in which the assimilation of ammonia plays a major role in determining cell fate and pattern formation. This model emphasizes a recycling of ammonia by prespore cells, the accumulation of free hydrophilic and neutral amino acids, and their incorporation into proteins associated with sporulation and (or) germination. If spore cAMP signalling is regulated by the relative concentrations of discadenine and autoactivator, and its disruption triggers the initiation of the spore germination cascade, then the accumulation of intracellular cAMP may be necessary for both sporulation and dormancy maintenance. Key words: Dictyostelium, development, ammonia, CAMP, discadenine, spore autoactivator, differentiation-inducing factor, glutamine, amino acids. COTTER,D. A., SANDS,T. W., VIRDY,K. J., NORTH,M. J., KLEIN,G., et SATRE,M. 1992. Patterning of development in Dictyostelium discoideum: factors regulating growth, differentiation, spore dormancy, and germination. Biochem. Cell Biol. 70 : 892-919. La communication entre cellules est essentielle pour toutes les Ctapes de la croissance et du dkveloppement des myxomycktes. Dictyostelium discoideum utilise pour cela plusieurs signaux moleculaires qui comprennent des facteurs de croissance, des facteurs de rtponse a la densite cellulaire, I'AMP cyclique, I'ammoniaque, le facteur d'induction de la differenciation, la discadenine et I'autoactivateur des spores. Un modkle de production-elimination est propose dans lequel l'assimilation de l'ammoniaque joue un r61e majeur dans le destin individuel des cellules et dans l'organisation multicellulaire. Le modkle met en relief le recyclage de l'ammoniaque par les cellules prespores et I'accumulation d'acides aminks libres hydrophiles et neutres qui s'incorporent dans des protkines impliquees dans la sporulation et (ou) la germination. La communication entre les spores contr81Ce par I'AMP cyclique pourrait Ctre regulee par les concentrations relatives de discadenine et d'autoactivateur, et une perturbation de cet equilibre declencherait la cascade de reactions qui conduit a la germination des spores. De ce fait, l'accumulation d'AMP cyclique intracellulaire serait necessaire a la fois pour la sporulation et le maintien de la dormance. Mots c l k : Dictyostelium, differenciation, arnrnoniaque, AMP cyclique, discadenine, autoactivateur des spores, facteur d'induction de la differenciation, glutamine, acides aminks.

Introduction The cellular slime molds are ideal organisms for studying evolutionarily conserved signal transduction and metabolic pathways occurring in such diverse eucaryotes as fungi,

protozoa, and mammals. For instance, the asexual life cycle of Dictyostelium discoideum has been particularly useful in examining the basic regulatory mechanisms controlling eucaryotic cell type differentiation and pattern formation (Bonner 1967; Loomis 1975, 1982; Raper 1984). This organism has been frequently chosen because vegetative cells ABBREVIATIONS: DIF, differentiation-inducing factor; DGF, density growth factor; PSF, prestarvation factor; Ins(1,4,5)P3, terminally differentiate into two main cell types, i.e., dead inositol 1,4,5-trisphosphate; CMF-L and CMF-H, low and high stalk cells and viable dormant spores (Fig. I), and vegetative molecular weight forms of conditioned medium factor; TCA, growth is temporally separated from the other stages of the tricarboxylic acid; pst, prestalk; alk, anterior like; CABP, CAMPlife cycle. binding protein; pk-A, CAMP-dependent protein kinase; Although the cellular slime molds are believed to possess R, regulatory; C, catalytic; DMO, 5,s-dimethyl-2,4-oxazolionly 28 000 single copy genes, they are capable of entering dinedione; PsA, matrix protein; SDS-PAGE, sodium dodecyl three separate developmental pathways (Firtel and Bonner sulfate - polyacrylamide gel electrophoresis; CBF, cAMP binding 1972; Loomis 1982; Raper 1984). The simplest of these, the factor; CAE, CA-rich element(s); Br-CAMP, 8-bromo-CAMP; microcyst pathway in Polysphondylium pallidum, is initiated DTT, dithiothreitol; DMSO, dimethyl sulfoxide; HPLC, high by an increase in osmotic pressure which, in turn, induces pressure liquid chromatography; SAAR, signature amino acid an individual amoeba to cease feeding, round up, secrete residue(s); CREB, CAMP-response element-binding factor. Pr~nledIn Canada / Imprime au Canada


a microcyst wall, and enter dormancy; a decrease in osmotic pressure results in microcyst germination (Raper 1984; North et al. 1991a; Klein et al. 1992). The second, more studied developmental pathway is initiated by starvation. In Dictyostelium, it involves secretion of CAMP,which induces a cascade of cellular events ending with the production of a fruiting body (see Fig. 1; and Loomis 1982). The third developmental pathway involves genetic recombination between starved cells of opposite mating types, which meet in an environment containing free c a 2 + . During aggregation, mediated by CAMP, a pair of cells fuses to form a zygote, which then engulfs the rest of the haploid cell population (Blaskovics and Raper 1957; Clark et al. 1973; Raper 1984; Amagai and Filosa 1984; Lydan and O'Day 1990; North et al. 1990b). While the cells are being engulfed, they begin to synthesize a tripartite wall to become dormant macrocysts (West and Erdos 1990). The macrocysts mature, meiosis and mitosis occur, and a mass of amoebae emerges from the cysts completing the sexual life cycle (Raper 1984; Abe and Maeda 1986). Because such alternative life cycles occur, they should also be considered in any theoretical modelling of the developmental processes in D. discoideum. The cellular and molecular biology of development in D. discoideum has been primarily studied using six variations of culture techniques: (i) growth with bacteria or axenic liquid media, (ill growth and multicellular development on moist surfaces, (iil) development of submerged cell masses in unshaken or slowly shaken conditions, (iv) slow shake cultures allowing cell-to-cell contact, (v) fast shake cultures preventing cell-to-cell contact, and (vi) development of noncontiguous wild type and sporogenous mutants as individual cells at very low cell densities on moist surfaces (monolayer cultures). Other studies have involved grafting segments of pseudoplasmodia (slugs) and (or) the analysis of complete slugs and fruiting bodies. Each of the above techniques provides complementary information about the various stages of the D. discoideum life cycle (see Fig. 1). Numerous studies have examined the following developmental life cycle stages: (i)vegetative growth and preaggregation; (ii) aggregation; (iii) prespore cell, prestalk cell, and tip development; (iv) pseudoplasmodial formation and migration, and pattern maintenance; and (v) culmination, and terminal stalk and spore differentiations. Such studies have considerably advanced our knowledge of the regulation of cell type divergence and pattern formation; however, few studies have focused on regulatory controls in the stages of (vi) spore dormancy and (vii) germination. Although the regulatory controls governing these latter two stages remain the least understood, new insight is being provided from research concerning terminal cell differentiation. Recent reviews have detailed some of the important roles of CAMP, DIF, and ammonia in D. discoideum cell differentiation and pattern formation (Weeks and Gross 1991; Schaap 1991). In addition to our attempt to model the regulation of dormancy and germination, we will also describe the potential regulatory roles of five morphogens (i.e., CAMP, NH3, DIF, discadenine and autoactivator) and their control functions in the various stages of the life cycle of D. discoideum. Vegetative growth and preaggregation A single amoeba is capable of growing and dividing at the expense of nutrients to produce a large population of

FIG. I. The life cycle of Dictyostelium discoideum. The following stages are diagrammed: (a) spore dispersal; (b)spore germination; (c) vegetative growth; (d) nutrient starvation to preaggregation, time 0-5 h; (e) beginning of aggregation, time 6 h; (f) middle of aggregation, time 8 h; (g) late aggregation, time 9 h; (h) tipped aggregation, time 1 1 h; (i) standing slug, time 13 h; (j)initiation of pseudoplasmodium (slug) migration, time 16 h; (k) end of short slug migration period, time 18 h; (I) reestablishment of vertical polarity in preparation for culmination, time 19 h; (m) initiation of culmination, time 20 h; (n) early culmination, time 21 h; (0) middle culmination, time 22 h; @) culmination complete, time 24 h; and (q) fruiting body and spore maturation, time 1-10 days.

vegetative cells. Growth is controlled by several secreted polypeptide hormones. For instance, DGF, a putative growth factor, is secreted by growing D. discoideum cells and may regulate the mitogenic pathway (Whitbread et al. 1991). In addition, vegetatively growing cells continuously secrete PSF protein in a density-dependent manner (Clarke et al. 1987, 1988). Prior to starvation and the initiation of development, PSF reaches a critical threshold concentration, which then triggers the synthesis of certain lysosomal enzymes (cr-mannosidase-1 and 0-galactosidase-2) and regulates transcription of other early developmental genes (e.g., discoidin 1) (Rathi et al. 1991). Earlier studies implicated a small, heat-stable, secreted compound as an effector of the expression of early developmental genes (Grabel and Loomis 1977, 1978). The increase in this yet uncharacterized effector is also proportional to the exponential growth of cells; at a cell titer of 5 x l o S / m ~the , effector reaches a critical threshold to induce the genes of early development, even in the presence of excess nutrients (Loomis 1989). Other low molecular weight factors are also secreted for the initiation of multicellular development (Hanna and Cox 1978; Klein 1992). Thus, it appears that amoeboid growth and the induction of early development are exquisitely regulated by a number of secretory molecules,


894

BIOCHEM. CELL BIOL. VOL. 70, 1992

which signal both the cell density and the remaining level of nutrients (Klein and Darmon 1976; Darmon and Klein 1978; Fontana and Price 1989; Gomer et al. 1991). The ability of cells to monitor their population density, by the concentration of secreted molecules, ensures that sufficient cells are in proximity for development (Loomis 1989). Levels of amino acids may be the key factor initiating development (Lee 1972; Marin 1976, 1977; Franke and Kessin 1977; Darmon and Klein 1978; Murphy and Klein 1979; Margolskee et al. 1980). When the amino acid starvation threshold is reached, a portion of the cells initiate the preaggregation stage of development. From this point onward, cells do not feed and the dry weight, as well as the protein content, fall by 50%; however, the carbohydrate content of the cell mass remains relatively constant. Therefore, protease activity controlling amino acid catabolism may provide a major source of energy for the multicellular developmental program (see Loomis 1975). The proteases and amino acid transaminases involved in early development have been investigated (Wright and Anderson 1960; Wright 1963; Wright and Bard 1963; Bruhmuller and Wright 1963; Krivanek and Krivanek 1965; Cleland and Coe 1969; Firtel and Brackenbury 1972; Pong and Loomis 1971; Fong and Rutherford 1978; North and Harwood 1979). Recently, an NADH-dependent glutamate dehydrogenase, present in both vegetative cells and early developing cells, has been shown to be activated by 5'-AMP and ADP, suggesting that a low energy charge in starving cells contributes to amino acid turnover (Pamula and Weldrake 1990, 1991, 1992). Protein catabolism produces excess ammonia; however, the intracellular level of free glutamate has been reported to increase during the 24-h developmental process (Wright 1963). A central problem in developmental biology concerns cell fate determination in multicellular animals. In D. discoideum, several researchers agree that the initiation of cell fate is determined at that point in the vegetative cell cycle when cells are starved and challenged to differentiate (McDonald and Durston 1984; Sharpe and Watts 1985; Gomer and Firtel 1987; Maeda et al. 1989). Another preaggregative condition known to influence cell fate is the age of the cells (Leach et al. 1973; McDonald 1980, 1984; McDonald and Durston 1984). Cells in the early portion of the cell cycle or those starved for long periods of time (aged cells) sort preferentially to the anterior (prestalk region) of the slug to become stalk cells of the sorocarp. These two conditions may both be related to the metabolic status of the cells, since cells in the early phase of the cell cycle will contain approximately half of the energy reserves of those in the late phase, and of course, starved cells will have lower energy reserves than unstarved cells (Weijer et al. 1984). Schaap (1986) has further suggested that such differentiation may be specifically related to intracellular carbohydrate reserves, since amoebae grown in glucose containing media (G cells) formed spores in preference to amoebae grown in the absence of glucose (NS cells), when the two were mixed and allowed to form slugs and fruiting bodies. To further emphasize the importance of carbohydrate reserves in cell fate determination, it has been shown that when the prestalk area (from slugs containing G cells) was grafted to the prespore area (from slugs containing NS cells), the cells did not remain in place, but sorted out, such that

the G cells became spores (Garrod and Ashworth 1973; Tasaka and Takeuchi 1981). Regardless of the specific nutritional conditions determining cell fate, preaggregation events commence prior to the complete exhaustion of nutrients from the medium. Earliest events include the secretion of a soluble cAMP phosphodiesterase to degrade background levels of cAMP in the medium (Riedel and Gerisch 1971; Gerisch et a/. 1972; Malkinson and Ashworth 1973; Tsang and Coukell 1977). Following nutrient exhaustion, specific classes of vegetative genes are no longer transcribed (McPherson and Singleton 1992) and a phosphodiesterase inhibitory protein is released to inhibit the activity of the soluble cAMP phosphodiesterase (Gerisch et al. 1972). The single copy phosphodiesterase gene @dsA) is then transcribed to synthesize a cell surface form of phosphodiesterase, resistant to the secreted phosphodiesterase inhibitor (Franke et al. 1991). Other events including the secretion of lysosomal enzymes, such as trehalase-1 and various cysteine proteases (e.g., CP5l and CP42), begin approximately at this time (Dimond et al. 1981; Seshadri et al. 1986; North et al. 1990~). The synthesis of two cAMP cell surface receptors, CARl and CAR3, and two internal and adaptable signal transduction systems occurs in the majority of cells, and this allows them to receive and relay nanomolar cAMP pulses secreted every 6 min by a small group of more advanced cells (Devreotes and Sherring 1985; Klein, P.S., et al. 1988; Firtel et a/. 1989; Saxe et al. 1991). The reception of cAMP pulses results in an autocatalytic advancement in the developmental programs of both advanced cells and relaying cells. These advanced cells can be compared to the embryonic organizers in mammalian tissues, and their removal allows others to quickly replace them. The signal transduction pathway linked by CARl to the adenyl cyclase for aggregation relays the nanomolar cAMP pulses (Devreotes 1982, 1989). After the detection of a cAMP pulse, a fourfold increase in internal c a 2 + levels occurs within 30 s (Abe et al. 1988); this increase may result both from the external uptake and the release from internal c a 2 + stores (Bumann et al. 1984; Coukell and Cameron 1988; Blumberg et al. 1989; Milne and Coukell 1991). A pulse of cAMP is then secreted from the relaying cell; however, not all of the newly synthesized cAMP is secreted, since some is retained to serve a second messenger function(s). Alterations in this pathway could disrupt normal aggregation (Coukell and Chan 1980). For instance, ammonia is known to delay aggregation in aggregation competent cells of Dictyostelium by changing the levels of intercellular cAMP (Thadani et al. 1977; Schindler and Sussman 1977; Feit 1988). In addition, stimulatory and inhibitory G-proteins may serve important regulatory function(s) in this pathway (Newell et a/. 1988; Firtel et a/. 1989; Saxe et al. 1991; Van Haastert et al. 1991). The chemotaxis/differentiation signal transduction pathway most likely involves CAR3 coupled to Ga2 protein, guanylate cyclase, and phospholipase C. The generated second messengers such as cGMP, diacylglycerol, Ins(1 ,4,5)P3, and c a 2 + regulate actin-myosin polymerization and result in chemotaxis (see Saxe et a/. 1991; Van Haastert et al. 1991). Therefore, second messengers such as diacylglycerol and C a 2 + ,either present alone or in various combinations with and without internal CAMP, are


REVIEWS / SYNTHESES

involved in the regulation of gene expression during development (Van Haastert et al. 1991). Further cellular differentiation in the life cycle is thought to be mediated by sustained (nonpulsatile), high levels of cAMP (Gross et al. 1981; Mehdy et al. 1983; Chisholm et al. 1984; Riley et al. 1989; Peters et al. 1991). These late developmental pathways also seem to require cAMP cell surface receptors, CAR2 and CAR4, which may be linked to the as yet uncharacterized, nonadaptive signal transduction pathway(s) involving protein kinase A (Schaap and Van Driel 1985; Gomer et al. 1986; Haribabu and Dottin 1986; Oyama and Blumberg 1986a; Kimmel 1987; Kwong et a/. 1988b; Simon et al. 1989; Firtel and Chapman 1990; Saxe et al. 1991). Aggregation During early aggregation, cells migrate toward high cAMP concentrations and bind to one another in a head-to-tail arrangement which is mediated by specific CAMP-induced cell surface binding proteins such as gp24, followed later by gp80 (CsA) (Gerisch 1968; Beug and Gerisch 1972; Lam and Siu 1981; Loomis 1988). After establishing contact, cells then enter an aggregate in streams to eventually form a tight aggregate or mound containing up to lo5 cells (see Loomis 1982). During this stage of development, cells expressing aggregation-specific genes secrete a class of CMFs. These factors, in addition to CAMP, initiate differentiation, triggering the expression of cell-type-specific genes (Gomer et al. 1991). The low molecular weight CMF forms (CMF-L) are much more potent in inducing differentiation than the high molecular weight form (CMF-H) (Yuen et al. 1991). We suggest that the breakdown of CMF-H into CMF-L may be mediated by concurrently secreted cysteine proteases (e.g., CP51 or CP42). Further development involving the formation of a tip and a surface sheath requires an air-water interface and thus, does not occur under submerged conditions (Gerisch 1968; Beug and Gerisch 1972). Prespore cells, prestalk cells, and tip formation We have found that development can be arrested at the mound stage by allowing aggregations to occur in 96-well microtiter plates containing 40 pL of cell suspension/well. Normal development proceeding to tip and sheath formation will occur if the well buffer volume is reduced from 40 to 20 pL at any time up to 24 h after the inhibition has been observed. The developmental CP48 and CP43 proteases may correspond to cprA and (or) cprB product(s) and are synthesized before the transition from the mound to the tipped mound stage (Cotter et al. 1992). As development proceeds further, it appears that they become increasingly cryptic in prespore cells, but not prestalk cells (T. Sands and D. Cotter, unpublished). The transition to the tipped mound stage also involves the synthesis of many other spore specific proteins and the appearance of spore-associated enzyme activities and organelles such as the prespore vacuoles (Hohl and Hamamoto 1969; Gregg and Badman 1970; Krefft et al. 1984; Morrissey et al. 1984; Miiller and Hohl1973; Cardelli et al. 1985,1990; Lam and Siu 1981; Devine et al. 1983; Wilkinson et al. 1985; West and Erdos 1990). There is also

895

some evidence that at this time the a-mannosidase (synthesized after starvation) may be transferred to the prespore vesicle, and then later secreted along with spore wall components after culmination (Lenhard et al. 1989). West and Erdos (1990) have concluded that while the prespore vesicle contains some lysosomal enzymes, the vesicle is not acidic and coexists with typical lysosomes in the prespore cells at this time. cAMP is required for the expression of both prespore and some prestalk specific genes (Mehdy et al. 1983; Chisholm et al. 1984). It was suggested earlier that the advanced cells at the center of the aggregation simply continue on to become the tip cells at the top of the mound (Takeuchi 1969; Loomis 1975; Schaap 1986). However, this has not been unambiguously verified using cells transformed with vectors containing ecmA and ecmB prestalk regulatory gene sequences coupled to reporter genes (Jermyn et al. 1989). Perhaps prestalk cells arise at random within the mound and then migrate independently to the tip (Jermyn et al. 1989). At any rate, some cells have entered the prestalk pathway by this stage (Williams et al. 1989a). It is still possible that there is a continuous developmental process from the more advanced cell to the prestalk cell and then on to the terminal stalk cell (Schaap 1986). The apparent random differentiation of prestalk cells in the mound leading to tip formation may result from a mixing of nascent prestalk cells with relaying cells during late aggregation. This mixing may be facilitated during a switch from one class of cell surface binding protein@)to another; proteins such as gp24 and gp80, among others, might be involved. Therefore, the 20% of the mound cell population normally fated to become stalk cells (ecmA- and ecmBexpressing cells; see Table 1) may move as individual cells up to the tip or down to the base. If our hypothesis for tip formation is tenable, then it becomes necessary to merge the theoretical models describing cell fate based on position effect with those based on cell sorting (Schaap 1986, 1991; Weeks and Gross 1991; Williams et al. 1989a; Williams 1991). During prestalk differentiation, ammonia is produced abundantly and may accumulate up to 12 mM in the extracellular media (Schindler and Sussman 1977; Sternfeld and David 1979). It would appear that prestalk cells are metabolically more active than prespore cells (Bonner et al. 1955; Takeuchi 1969). This idea is reasonable, since the fate of prestalk cells is to pass through a series of transitions to express ecmA and ecmB, synthesize cellulose, lose nitrogen (as ammonia), and then swell and die; alternatively, the fate of prespore cells is to accumulate reserves (lipid and trehalose), become nitrogen rich (as amino acids), condense, encapsulate, and enter dormancy (Williams et al. 1989b; Klein, G., et al. 1988, 1990; Jermyn and Williams 1991; Weeks and Gross 1991). It is possible that these two cell fate pathways may have been initiated during starvation prior to the start of aggregation. Schaap (1986) suggests that the preferential expression of aggregation-specific components is probably responsible for the sorting behaviour of prestalkpreferring cells. If amino acid starvation triggers the aggregation process, then we suggest that cells entering the prestalk-stalk differentiation pathway will degrade their proteins in preference to carbohydrates, thus releasing CO, and NH3. Alternatively,


896

TABLE1. SAAR compositional analysis of Dictyostelium discoideum proteins


Life cycle stage of the gene expression

Time of expression"

Gene and (or) protein description

Morphogen regulation

% signature amino acid residues (SAAR)/ polypeptideC

Ref. Singleton 1989; Singleton et al. 1989; Singleton et al. 1991

Postamoebae emergence, vegetative growth

6-7 h, after spore activation; primarily vegetative growth

-

V1, ribosomal protein

Postamoebae emergence, vegetative growth

6-7 h, after spore activation; primarily vegetative growth

-

V14

Singleton 1989; Singleton et a/. 1989; Singleton et al. 1991

Entire life cycle

All stages

Some cAMP +

Actins

Vanderchove and Weber 1980; Romans and Firtel 1985

Entire life cycle

All stages

-

Cap 32/34, actin capping protein

Hartmann et al. 1990

Entire life cycle

All stages

-

Cyclophilin

Bariscic et al. 1991

Preaggregation, 0 h, aggregation, 5-7 h, pseudoplasmodium 10-24 h to late culmination (prestalk cell specific)

CAMP

pdsA, CAMP phosphodiesterase

Podgorski et al. 1989

Growth and preaggregation

0-8 h

CAMP-

DdrasG

Robbins et al. 1991

Preaggregation to aggregation

2-3 h 6-8 h

CAMP-

pdiA, CAMP phosphodiesterase inhibitor

Wu and Franke 1990

Aggregation, 4-14 h early development, pseudoplasmodium (prestalk/prespore ratio, 3-4)

CAMP

carA, CAMP cellsurface receptor (CAR1)

Saxe et al. 1988, 1991; Klein, P.S., et al. 1988

Aggregation, later stages

CAMP

aca, adenyl cyclase aggregation (ACA)

Pitt et al. 1992

CAMP

Ddras

59.1

Reymond et al. 1984; Esch and Firtel 1991

pk-A, CAMPdependent protein kinase A regulatory subunit

57.4

Veron et al. 1988

3-6 h, >16 h

Aggregation to 8-16 h pseudoplasmodium (prestalk enriched) Preaggregation, 3-5 h aggregation to 8-10 h pseudoplasmodium (not cell specific)

+

+

+

+

-

Aggregation to early 8-14 h pseudoplasmodium

CAMP

csaA, cell surface glycoprotein (gp80, CsA)

Late aggregation to 9-16 h pseudoplasmodium (prestalk/prespore ratio, 1.5-2)

CAMP

cprA, cysteine protease I (CPl)

+

+

54.0

65.2 (66.5) 59.1 (60.8) ((60.9))

Siu and Kamboj 1990

Williams et al. 1985

TABLE1. SAAR compositional analysis of Dictyostelium discoideum proteins (continued)



Time of expressiona

Morphogen regulationb

Late aggregation to pseudoplasmodium (prestalk/prespore ratio, 3-4)

cAMP

At tip formation to culmination (prespore cell derived)

cAMP DIF -

Standing finger to late culmination (prestalk cell derived)

cAMP DIF

Pseudoplasmodium to late culmination (prestalk cell derived)

cAMP DIF

+

+

+

+

+

Entire life cycle


070 signature amino acid residues (SAAR)/ polypeptidec

Ref.

cprB, pst-cath, cysteine protease 2 (CP2)

Pears et al. 1985; Presse et al. 1986b; Datta and Firtel 1987

pspA, surface/slime sheath/matrix protein (D19, PsA, SP29)

Early et al. 1988a; Early and Williams 1988

ecmA, slime sheath/ stalk tube/stalk cell wall protein (pD63, ST430)

Williams et al. 1987; Early et al. 1988a; Berks and Kay 1990

ecmB, slime sheath/ stalk tube/stalk cell wall protein (pD56, ST310)

Ceccarelli et al. 1987; Early et al. 1988a; Berks and Kay 1990; Kwong et al. 1990

Annexin VII, synexin, c a 2 + channel protein

Doring et al. 1991; Greenwood and Tsang 1991

capA, CAMP-binding protein (CABPIA)

Grant and Tsang 1990

capA, CAMP-binding protein (CABPlB)

Grant and Tsang 1990

spiA, membrane protein (Dd3 1)

Richardson et al. 1991

Low level in vegetative cells increasing three- to four-fold during development

cAMP

Low level in vegetative cells increasing three- to four-fold during development

cAMP

Culminationsporulation (spore specific)

cAMP

Just after tip formation to midculmination

cAMP ' DIF -

cotC, spore coat protein (SP60)

Fosnaugh and Loomis 1989a, 1991; Widdowson et al. 1990


cAMP DIF -

cotB, spore coat protein (2H3, SP70)

Fosnaugh and Loomis 1989a, 1991


cAMP DIF -

cotA, spore coat protein (SP96)

Fosnaugh and Loomis 1989b. 1991

EB4, spore coat protein?

Hildebrandt et al. 1991; Fosnaugh and Loomis 1991

After aggregation (prespore cells)

>9h (antisense regulation)

After slug formation (spore specific) Low in late culmination and dormant spores, increase during spore swelling

cAMP DIFcAMP

+

+

+

+

+

+

Widdowson et al. 1989

+

celA , cellulase (pRK270-6) during spore germination

Giorda et al. 1990; Ennis et al. 1991

898


TABLE1. SAAR compositional analysis of Dictyostelium discoideum proteins (concluded) -



Time of expressiona

Morphogen regulation

-


070 signature amino acid residues (SAAR)/ polypeptideC

Ref.

-

pRK270-11

79.1 (80.5)

Giorda et al. 1990; Ennis et al. 1991

22-24 h, 0-1.5 h during spore germination

-

pLK 109

55.9 (59.8)

Giorda et al. 1989

24-42 h, 0-2 h during spore germination

-

acg, adenyl cyclase germination (ACG)

60.1

Pitt et al. 1992

Late germination and >3.0 h during vegetative growth spore germination until 0 h of development

-

V4, for transition from vegetative growth to development

63.8

McPherson and Singleton 1992

Low in late culmination and dormant spores, increase during spore swelling

22-24 h, 0-2 h during spore germination

Low in late culmination and dormant spores, increases during spore swelling After sporulation, early germination

"The developmental time of gene and (or) protein expression varies depending upon environmental conditions and the strain used; the values above are from published data and may have been adjusted to a 24-h developmental clock as shown in Fig. 1 . b + . stimulation; - , inhibition; -, not known. 'SAARVo refers to the percent of the amino acid residues in the polypeptide which are Q, E, N, G, A. D, S, T, K, or P; SAARVo in single parentheses refers to the SAARVo of the polypeptide without its signal sequence; SAAR% in double parentheses refers to the SAAR% of the polypeptide without a signal sequence and without (other) N- or C-termini. Some gene designations were selected from Kuspa el al. 1992.

FIG. 2. The ammonia, CAMP, DIF source, and sink model of slug pattern formation and maintenance. A, ammonia; C, CAMP; D, DIF. The dashed line represents the prestalk-prespore boundary. Secretion of CAMP,primarily by prestalk cells, regulates slug motility by chemotaxis and maintains cell-specific gene expression. For such secretion, prestalk cells must eliminate ammonia by the following: evaporation from the prestalk region, secretion of extracellular prestalk proteins, and diffusion to the prespore region. Ammonia is used by the prespore cells to synthesize hydrophilic and neutral amino acids; some of which accumulate in prespore lysosomes and are incorporated into both cellular and secreted prespore or spore proteins. DIF is secreted by prespore cells, but only the prestalk cells respond to and degrade it, resulting in the patterning observed in the migrating slug.

the majority of cells entering the prespore-spore pathway should be metabolically sluggish to conserve their total energy reserves. Therefore, such cells could primarily degrade their carbohydrates, producing glycolytic and TCA intermediates useful in recycling some of the prestalk secreted NH3 in the form of amino acids. Clearly, it is not surprising that NH3 is a prime inducer of microcyst formation in the closely related slime mold Polysphondylium pallidum (Lonski 1976; Choi and O'Day 1982), which like spores of D. discoideum synthesize trehalose and cellulose, and are also very rich in specific

amino acids formed from glycolytic and TCA cycle intermediates (Ennis 1981; Klein et al. 1990, 1992). In this scenario, NADH-dependent glutamate dehydrogenase (Pamula and Wheldrake 1990, 1991) may function primarily to degrade glutamate (E) from protein catabolism to a-ketoglutarate and the waste product NH4+ during prestalk-stalk development. During prespore-spore development, this same enzyme or the NADPH-dependent glutamate dehydrogenase would function to assimilate the secreted NH4+ with a-ketoglutarate (from glycogen catabolism) to produce E (Klein et al. 1990; Pamula and Wheldrake 1990, 1991). Additional amino acids such as aspartate (D) and alanine (A), detectable in dormant spores and microcysts, can be similarly produced (Klein et al. 1990, 1992). Finally, transaminase reactions would allow the synthesis of other amino acids from excess NH4 secreted by the prestalk-stalk cells (Wright and Anderson 1960; Wright 1963; Wright and Bard 1963; Krivanek and Krivanek 1965; Cleland and Coe 1969). This source and sink model (see Fig. 2) is consistent with amino acid analyses using chemical methods, as well as ' 3 ~ -which ~ ~ revealed ~ , that the free amino acid pool of less than 10 mM concentration in D. discoideum and P. pallidum vegetative cells accumulated to over 100 mM in spores and microcysts, respectively (Ennis 1981; Klein et al. 1990, 1992). In D. discoideum, the largest increases occurred in only nine amino acids, with glutamine (Q) increasing 48-fold, followed by high concentrations of E and asparagine (N) (Klein et al. 1990); other amino acids in relatively high concentration in these studies included glycine +


REVIEWS / SYNTHESES

(G), serine (S), threonine (T), lysine (K), A, and D (Ennis 1981; Klein et al. 1990). Ammonia assimilation by prespore cells may lead to increasing concentrations of those amino acids that are most rapidly synthesized from glycolytic and TCA cycle precursors. Amino acids that D. discoideum cannot synthesize are thus spared for turnover into new prespore and spore proteins. Consequently, ammonia recycling may be crucial to establishing dormant spore reserves. These ideas are consistent with prespore cells incorporating 10-fold more radiolabeled amino acids per cell than prestalk cells (Morrissey et al. 1984). Bonner et al. (1984) also reported that glucose utilization and amino acid uptake differed in prestalk and prespore cells of migrating slugs. In general, the prestalk cells closely resemble aggregationcompetent cells and continue to secrete cAMP (and large quantities of ammonia), while prespore cells lose such aggregation-related properties, making them distinct from aggregation-competent cells (Schaap 1986, 1991). Future work is necessary to determine whether the mitochondria1 TCA cycle (Albe and Wright 1992) assumes alternative roles in prestalk and prespore cells. A source and sink model for control of slug formation and patterning During tip development, the cellular mass lengthens and becomes covered with an extracellular sheath composed of cellulose and proteins (Raper 1940; Takeuchi 1969; Loomis 1972; Hohl and Jehli 1973; Durston 1976). The upright slug may then topple over and begin migrating (see Figs. 1 and 2). Siegert and Weijer (1991) concluded that cells rotate around a central core in the aggregate, which eventually forms the tip. The movements of prespore and prestalk cells within the slug are dramatically different. Prestalk cells move perpendicular to the long axis of the slug, while prespore cells move parallel to the long axis. The prestalk zone is thus organized in a scroll wave (Siegert and Weijer 1991). Another model of pattern formation in a migrating slug maintains that the prespore zone is surrounded by a peripheral layer of normal prestalk cells making contact with the slime sheath (Breen and Williams 1988). We hypothesize that the scroll wave gives rise to the peripheral cells which may play a major role in slug motility by moving in a spiral fashion from the tip to the posterior of the slug; this idea implies that the slug mass composed of prestalk and prespore cells rotates as it moves forward within the sheath. The peripheral cells upon reaching the posterior of the slug return to the anterior prestalk region as individual cells by moving through the prespore mass (see Williams 1991). Such returning cells have been called anterior-like cells because they have prestalk characteristics, but they are scattered throughout the prespore zone of the slug (Sternfeld and David 1982; Breen and Williams 1988). These peripheral cells have vital staining patterns similar to prestalk cells (Bonner et al. 1990). As further suggested by Weeks and Gross (1991), the cells typically classified as prestalk (pst) cells and anteriorlike (alk) cells may represent a single class (pst-alk) of highly motile cells with a gradient of shared characteristics. If this is true, then one may envision that the prespore cells are relatively nonmotile and that the motility of the slug is exercised primarily by the highly motile pst-alk cells adjacent to the slime sheath. Furthermore, we suggest that when culmination is triggered, the continual recycling of the pst-alk cells is halted, and any pst-alk cells which happen

899

to be caught in the far posterior region of the slug are then fated to become basal disc cells. For these reasons we propose the presence of only two basic cell types (i.e., prespore and prestalk) in a migrating slug. Ammonia has been well established as an inhibitory morphogen for prestalk and stalk cell differentiation, while still allowing sporulation (Cohen 1953). Concentrations have been shown to be approximately 7.4 x M in the prestalk region of the slug, but 5.7 x lo-' M in the prespore region and 5.8 x M in the far posterior region (destined to become dead basal disc cells) (I.N. Feit, personal communication). The presence of ammonia prolongs slug migration by preventing the transition of prestalk cells into terminally differentiated stalk cells (Sussman and Schindler 1978). The volume of the prestalk region has been shown to decrease by 40.8% within 10-20 min of exposure to NH4CI at M, while the number of anterior-like cells increased by an equivalent amount in the posterior region of the migrating slug (Feit et al. 1990). Research by Feit et al. (1990) is consistent with their hypothesis that the tip produces NH3 which controls the return of anterior-like cells to the prestalk mass. We suggest that when a migrating slug is subjected to a high background level of this volatile morphogen that the most effective arrangement for depletion and recycling NH3 (from prestalk cells into prespore cells) is for the masses of prespore cells to surround individual prestalk cells (returning anterior-like cells). Such an arrangement would trap or retard the return of anterior-like cells to the tip, reducing the tip volume and facilitating the evaporation of NH3 from the tip. Another critical morphogen, CAMP, is initially produced in large quantities by prestalk cells (Pan et al. 1974; Brenner 1977) and studies using sporogenous mutants, cultured to prevent cell-to-cell contact, have shown that cAMP concentrations must decrease to allow terminal stalk cell differentiation (Sobolewski et al. 1983; Berks and Kay 1988, 1990; Kwong and Weeks 1990). This contrasts with the prespore to spore differentiation pathway which is driven by high internal cAMP and ammonia. Some prespore differentiation is also observed when small clumps of cells are submerged in buffer containing cAMP and glucose (Oyama et al. 1983). The cyclic nucleotide is necessary for the induction of all the prespore-specific genes studied to date and that of a major class of prestalk genes (Barklis and Lodish 1983; Chisholm et al. 1984; Mehdy et al. 1983; Weijer and Durston 1985; Oyama and Blumberg 1986a, 1986b, 1986~; Wang et al. 1988; Yamada and Okamoto 1992). There is increasing evidence to suggest that cAMP induction of prespore gene expression is mechanistically distinct from cAMP induction of prestalk cell genes; specifically, the expression of prestalk-specific mRNA requires a three- to four-fold lower cAMP concentration than the expression of prespore specific genes (Kwong and Weeks 1990). During sporulation, it is evident that internal cAMP is required for the continuation of the prespore pathway, possibly by the replacement of the cAMP cell surface receptor (CARI) by CAR2 during slug migration and by CAR4 during culmination (Saxe et al. 1991). cAMP not only stimulates the transcription of cell-specific genes (Oyama and Blumberg 19868; Landfear et al. 1982; Pears and Williams 1988; Datta and Firtel1988; Manrow and Jacobson 1988), but also serves to stabilize many of these transcripts (Mangiarotti et al. 1983; Landfear et al. 1982) (see Table 1).


900


A number of CABPs are expressed throughout the entire life cycle (Tsang and Tasaka 1986; Tsang et al. 1988; Grant and Tsang 1990). The regulation of cAMP directed gene expression may be mediated by the CABP1 protein present in both the cytoplasm and nucleus of developing cells (Kay et al. 1987; Tsang et al. 1988). As in other eukaryotic cells, elevated intracellular cAMP also increases the activity of the CAMP-dependent protein kinase (pk-A) by dissociation of the regulatory (R) subunit from the catalytic (C) subunit (de Gunzburg and Veron 1982; Leichtling et al. 1982; Rutherford et al. 1982; Majerfeld et al. 1984; Mutzel et al. 1987; Veron et al. 1988; Firtel and Chapman 1990; Harwood et al. 1992). The levels of both subunits increase during development and peak toward the end of cellular aggregation (Leichtling et al. 1984; de Gunzburg et al. 1986). Recently, AX2 pk-A- cells were constructed by C subunit inactivation (Harwood et al. 1992). These cells were ineffective in cAMP relay and aggregation, but were shown to respond to CAMP and to aggregate partially when mixed with wild-type cells; however, the pk-A - cells in the aggregate did not express the postaggregate genes pspA, ecmA , and ecmB (Harwood et al. 1992). A defective regulatory subunit of pk-A has been detected in one class of rapidly developing mutants (rde)of Dictyostelium and inactivation of this protein resulted in the uncontrolled activity of the catalytic subunit, prematurely triggering disorganized terminal cell differentiation (Simon et al. 1992). Overexpression of the entire Dictyostelium protein kinase (PK2) gene stimulated development and sporulation; however, sporulation competency was lost in those cells expressing PK2 missing most of its catalytic domain (Anjard et al. 1992). Recently, five different members of the protein kinase multigene family as well as type 1 and 2u phosphoprotein phosphatases have been cloned and sequenced (Haribabu and Dottin 1991). The regulation of their expression during the life cycle should prove invaluable in elucidating their roles in signal transduction and cell fate determination. Another class of morphogens include the DIFs. This class of chlorinated hexaphenones is normally present in both the cell mass and the medium during late aggregation and before tip formation (Brookman et al. 1982; Kopachik et al. 1983; Neave et al. 1986; Kay et al. 1988; Kwong and Weeks 1989). DIF-1 is a powerful morphogen stimulating prestalk cell formation and inhibiting prespore formation when added to individual cells at nanomolar concentrations (Kay and Jermyn 1983; Berks and Kay 1990; Berks et al. 1991). In the aggregate, both DIF-1 and DIF-2 promote stalk cell differentiation and inhibit spore cell differentiation (Williams 1988; Xie et al. 1991). The increased rate of DIF-1 induction of prestalk-specific gene expression (sensitive cells transcribe ecmA within 15 min of DIF-1 treatment) suggests that it acts directly at the level of gene expression similar to mammalian steroid hormones (Williams et al. 1987; Insall and Kay 1990). Levels of DIF are two- to three-fold higher in the posterior region of slugs than in the anterior regions and thus, prespore cells have been postulated to produce, but not respond to DIF (Brookman et al. 1987; Inouye 1989; Loomis 1989). There is also conflicting evidence that a subpopulation of prestalk cells, expressing ecmB, produces DIF (Kwong et al. 1990). Raper (1940) has shown that isolated anterior slug fragments do not form spores if forced to terminally differentiate immediately after the excision of

posterior cells. Rather, spores form only after anterior slug fragments are aged 4-8 h before being allowed to terminally differentiate. Therefore, both the spatial distribution of DIF and the regulation of cell proportioning can be explained by a model in which only prespore cells produce and only prestalk cells respond to DIF (see Loomis 1989). The occurrence of DIF-1 is transient since it is sequentially degraded by enzymes belonging to the DIF-1 signalling system; these specific enzymes become more active toward the end of aggregation and reach maximal levels at the tipped mound stage, just as endogenous DIF-1 levels are rising. This localized metabolism may generate DIF-1 gradients in the aggregate (Traynor and Kay 1991). The DIF gradient is also regulated by the recently isolated DIF-1 prestalk binding protein (Insall and Kay 1990). The expression of this protein is developmentally regulated (peaking shortly before the rise in DIF) and is augmented by cAMP (Insall and Kay 1990). Characterization of the prestalk DIF binding protein and its regulation of DIF should provide further insight concerning the mode of action of the DIF series of morphogens. The effects of DIF on isolated cells can be mimicked by weak acids and counteracted by weak bases such as ammonia (Gross et al. 1983, 1988; see Weeks and Gross 1991). The weak base methylamine inhibits DIF-induced expression of the prestalk genes ecmA and ecmB, but does not affect CAMP-induced expression of the prespore gene pspA (Van Lookeren Campagne et al. 1989; Wang et a/. 1990). In contrast, the weak acid DM0 enhances the effects of DIF, but cannot itself induce transcription of prestalk genes (Aerts et al. 1987; Aerts 1988; Wang et al. 1990). Therefore, it appears that DIF sensitivity is dictated by intracellular pH (Kay et al. 1986); in the presence of DIF, DIF synergists (weak acids) and DIF antagonists (ammonia) will inhibit or promote prespore gene expression, respectively (Wang et al. 1990). Invariably, ammonia appears to inhibit DIF accumulation (Neave et al. 1983) and its presence in the aggregate antagonizes the actions of DIF. Wang and Schaap (1989) have observed that DIF-induced stalk cell differentiation occurs in vivo, only after the removal of endogenously produced ammonia. Thus, it appears that this volatile morphogen affects migration, culmination, and cell proportioning in the slug. A class of D. discoideum mutants which are overly sensitive to ammonia (Newel1 and Ross 1982a, 1982b) shows a reduced ratio of prestalk to prespore cells in the slug (MacWilliams and David 1984). Specifically, this volatile morphogen induces prespore gene expression (Oyama and Blumberg 1986c; Oyama et al. 1988) and has been shown to switch cells in the prestalk pathway to that of the prespore pathway (Bradbury and Gross 1989). These morphogens and changes in their levels may differentially regulate the expression of cell-type specific genes, and hence, regulate differentiation. For instance, the expression of ecmA is enhanced by CAMP, while ecmB expression is inhibited by cAMP in the presence of DIF (Jermyn et al. 1987; Berks and Kay 1990; Kwong and Weeks 1990). The early steps during prestalk differentiation are highly cAMP dependent, whereas the later stages become sequentially cAMP independent and then are ultimately inhibited by cAMP (Sobolewski et al. 1983; Sobolewski and Weeks 1988; Kwong et al. 1988a, 1988b). In summary, the interactions and the resulting gradients of the three morphogens (see Fig. 2; source and sink model)


REVIEWS /

important in regulating pattern maintenance in the slug may be as follows. (1) The concentrations of free ammonia are highest in the prestalk region and lowest in the prespore region (I.N. Feit, personal communication). (ill The cAMP concentration is also highest in the anterior prestalk region and decreases down the length of the slug (Pan et al. 1974; Brenner 1977; Weeks and Gross 1991). (iig Lastly DIF-1 and DIF-2 are highest in the prespore region and decrease toward the prestalk region (Brookman et al. 1987). Ammonia, a key morphogen in our model (Fig. 2), is released from the metabolically active prestalk cells. This volatile gas has been shown to disrupt normal cAMP levels (Sussman and Schindler 1978; Schindler and Sussman 1979). Riley and Barclay (1990) specifically suggest that ammonia promotes the accumulation of intracellular cAMP by inhibiting cAMP secretion. In turn, cAMP has been shown to induce a sustained increase in intracellular pH (Van Lookeren Campagne et al. 1989). Since the ammoniainduced high intracellular cAMP (and therefore, high intracellular pH) is inhibitory to DIF-induced terminal stalk differentiation, then it becomes crucial for prestalk cells to effectively remove this morphogen. The high levels of ammonia produced by prestalk and basal disc cells can be reduced by the following mechanisms: (1) simple evaporation during slug migration, (ii) reassimilation into amino acids and proteins, such as the ST430 and ST310 secreted proteins, and (iii) uptake by prespore cells. Prespore differentiation is favoured by both higher intracellular cAMP and pH, and since prespore cells are more resistant to acid loading than prestalk cells (Inouye 1985, 1988; Wang et al. 1990), then they may be highly efficient in assimilating and recycling this ammonia for (0 the formation of free amino acids, (ii) the synthesis of proteins (such as the secreted PsA), and (iii) retention to maintain elevated internal cAMP (high intracellular pH) required for antagonizing DIF effects, promoting prespore gene expression, and terminal spore differentiation. Finally, when ammonia drops below a threshold level, stalk cell differentiation is stimulated (Loomis 1989) by DIF released by prespore cells. DIF antagonizes the effects of ammonia and should override any residual ammonia-induced inhibition of intracellular cAMP secretion and the CAMP-induced inhibition of stalk differentiation. It is possible that DIF decreases the affinity of the cAMP receptor for cAMP and concomitantly inhibits the cAMP response but not the CAMP-inducedcGMP response (Wang et al. 1986). At this stage, cell differentiation appears to be partly dictated by factors such as intracellular pH, ammonia, and CAMPwhich control cell-specific responsiveness to DIF (Wang et al. 1990). It has been shown that the NADH-dependent and the NADPH-dependent glutamate dehydrogenases are present in slugs of D. discoideum (Pamula and Weldrake 1990). It is possible that the NADH-dependent enzyme functions in prestalk cells to degrade glutamate to a-ketoglutarate and NH3, whereas the NADPH-dependent enzyme functions to assimilate the released ammonia with the TCA precursor a-ketoglutarate to form glutamate in prespore cells. Ammonia levels can be regulated in several ways by prestalk and prespore cells to optimize cell-specific CAMP, intracellular pH (which dictates cell-specific response to DIF), the initiation of DIF synthesis or release, and the timing of both slug migration and terminal cell differentiation. In our model of ammonia recycling, prestalk and prespore

cells reciprocally serve as both sources and sinks of the key morphogens. This pattern of ammonia recycling sets the stage for cell-specific responses induced by the prime gene regulating morphogens cAMP and DIF. The accumulated amino acids in the prespore cells include Q and N which are lysosomotropic (Klein et al. 1990); i.e., they may specifically accumulate along with ammonia in lysosomes and raise the pH of the organelle to alkaline shock resident enzymes such as the cysteine proteases. As a consequence, proteolytic activity and perhaps lysosomal enzyme activity in general would gradually diminish in cells committed to the prespore pathway. Such a conclusion is supported by the neutral red staining pattern of slugs. The anterior (prestalk region) of slugs stains positively with neutral red, whereas the posterior (prespore region) remains primarily unstained (Yamamoto and Takeuchi 1983; MacWilliams and Bonner 1979; see Raper 1984). Accordingly, the prestalk region (approximately 20% of the slug) contains cells with acidic vesicles which are either absent or are clearly not as acidic in the prespore cells. In yeast and fungi, lysosomotropic amino acids may have a lysosomal accumulation ratio of 5- to 20-fold over the concentration in the cytoplasm (see Holtzman 1989). It has also been observed that autophagic vacuoles in mammalian cells are prevented from fusing with primary lysosomes in the presence of N (Holtzman 1989). Thus, it is not surprising that the autophagic/lysosomal vacuolar system in prespore cells of slugs does not show the neutral red staining characteristics of the prestalk cells in which protein catabolism is expected to be high. Intuitively, prestalk cells can maintain acidic vesicles (as shown by neutral red staining), if they possess a mechanism for the secretion of ammonia. Studies have shown that the transcription of the genes cprA and cprB, encoding cysteine proteases CPl and CP2, respectively, occurs in both prestalk and prespore cells (Williams et al. 1985; Pears et al. 1985; Presse et al. 1986a, 1986b); however, the total protease activity actually decreases during development when examined by many techniques including gelatin-SDS-PAGE and protease assays using chromogenic synthetic peptide substrates (Fong and Rutherford 1978; Gustafson and Thon 1979; North and Cotter 1991a). If prespore cell extracts are electrophoresed on gelatin-SDS-PAGE gels and the gels are acid shocked in 10% acetic acid for 30-60 s, then two bands of proteolytic activities (M, 48 000 and 43 000) are observed (Cotter et al. 1992). This same acid shocking technique has also revealed the presence of the two cysteine proteases in dormant spore extracts (North et al. 1991b). Using conventional SDSPAGE techniques, these two protease activities normally have been shown to appear during spore germination (North et al. 1990a). The results suggest packaging of the two proteases in early prespore cells and their gradual deactivation during slug migration and culmination; the acid shock treatment revealing cryptic protease activity may mimic lysosomal acidification by acidosomes (see Padh et al. 1989, 1991) during spore germination. A number of CAMP-inducible genes such as cprA and cprB (pst-cath) contain CA-rich sequences in their promoter regions (Hjorth et al. 1990). A nuclear factor (CBF) binds cAMP and interacts with the CA-rich sequences activating transcription. In the case of the prespore cotC gene encoding a spore coat protein (SP60), there are three CAE with the consensus sequence CACACAYYYCACACA which lie


902

BIOCHEM. CELL BIOL.

upstream of the transcription start site. Deletion of the 5' most distal CAE of cotC reduces expression primarily in the prespore cells located in the posterior of the slug; deletion of the two most distal CAE restricts transcription to prespore cells located at the prestalk-prespore boundary of the slug. It is suggested that the spatial patterning of cotC expression could result from gradients within the prespore region of the slug (Haberstroh and Firtel 1990; Haberstroh et al. 1991). Possibly, the gradients in question may be those of cAMP and ammonia. Williams (1991) points out that this effect is similar to the blanching of neutral red stained nascent slugs in which clearing first becomes visible at the prestalk-prespore boundary and then proceeds posteriorly.

Culmination The presence cf overhead light or dry conditions can interrupt slug migration and trigger culmination (Schindler and Sussman 1977; Raper 1984). During culmination, the decline in ammonia concentration (Wang and Schaap 1989) is concurrent with the construction of a cellulose tube through which prestalk cells enter by a reverse fountain effect (Bonner 1967). As prestalk cells swell and die, they lift the prespore mass off the substrate. During this developmental stage, the continued transcription of a cAMP receptor (CAR4) suggests that adenyl cyclase and cAMP generation are required for sporulation (Saxe et al. 1991; Kimmel and Firtel 1991). Kay (1989) further hypothesizes that the decline in ammonia, triggering culmination, facilitates the two- to three-fold intracellular increase in cAMP during spore maturation (Brenner 1978; Pahlic and Rutherford 1979; Abe and Yanagisawa 1983; Merkle et al. 1984). Ammonia gas is also involved in negative chemotaxis or orientation of culminating fruiting bodies away from each other and the substratum (Feit and Sollitto 1987). Carbohydrate metabolism is regulated by the CAMPdirected signalling pathway. Specifically, glycogen phosphorylase activity is affected by cAMP during culmination (Brickey et al. 1990). In addition, a biphasic synthesis of trehalose appears to involve stalk and spore cells during culmination (Garrod and Ashworth 1973); in stalk cells this sugar is also rapidly hydrolyzed presumably to provide additional carbohydrate for cellulose synthesis (Jefferson and Rutherford 1976). The net transient accumulation of trehalose in stalk cells suggests that this sugar has a more significant role than simply one of storage. In fact, trehalose accumulates in heat, cold, and heavy metal stressed vegetative cells up to a level of 2% of the total dry weight in less than 2 h (Temesvari and Cotter 1991). Perhaps trehalose provides a protective effect against stressors; such a mechanism may have been selected for during evolution to protect membranes of spores, microcysts, and macrocysts (Emyanitoff and Wright 1979; Klein et al. 1990, 1992; Temesvari and Cotter 1991). In dormant spores, trehalose levels are controlled by the amount of glycogen initially present in starving vegetative cells, and the level of glycogen in vegetative amoebae is controlled by glucose concentrations (Hames and Ashworth 1974a, 1974b). Therefore, in addition to the accumulated amino acids, the carbohydrate trehalose also serves as a significant dormant spore energy reserve (Ceccarini and Filosa 1965; Emyanitoff and Wright 1979; Klein et al. 1990). Ultimately, the prespore mass reaches the top of the mature stalk and the spores encapsulate when prespore

VOL. 70,

1992

vesicles discharge their contents containing the spore coat proteins SP60, SP70, and SP96; cellulose and mucopolysaccharide are also deposited in the spore wall (Hohl and Hamamoto 1969; Maeda and Takeuchi 1969; Newel1 et al. 1969; Gregg and Badman 1970; Muller and Hohl1973; West and Erdos 1990).

Control of spore maturation, dormancy, and germination Spore formation in Dictyostelium appears to be driven by elevated internal cAMP (Kay et al. 1988; Kay 1989; Riley and Barclay 1986, 1989, 1990; Maeda 1992). Spore differentiation has been induced in wild-type amoebae incubated in submerged monolayers containing permeant cAMP analogues, Br-CAMP and 8-chlorophenylthio-CAMP, but not CAMPitself; however, such spores were unstable and a fraction germinated, but to a lesser extent when discadenine was present in the medium (Kay 1989). Maeda (1992) has also found Br-CAMPto be effective in inducing in vitro sporulation of NC4 prespore cells; in contrast, differentiation was inhibited by the protein kinase inhibitors K252a and staurosporine. Br-CAMP was shown to increase the phosphorylation of SP96 spore coat protein, whereas K252a blocked it; therefore, this protein may be a target of a protein kinase(s) that is activated by Br-CAMP (Maeda 1992). Both serinekhreonine- and tyrosine-specific kinases have been recently identified (Mann and Firtel 1991; Tan and Spudich 1990). Riley and Barclay (1990) suggest that ammonia favours spore formation by promoting the accumulation and maintenance of intracellular CAMP, and in turn, high intracellular cAMP may negatively control both the expression of the cellular cAMP phosphodiesterase gene and the accumulation of its enzyme activity. Both high pH and high ammonia levels stimulate the precocious accumulation of intracellular cAMP in both sporogenous and wildtype cells (Riley and Barclay 1990). This treatment stimulated the expression of the prespore pspA gene and inhibited the accumulation of prestalk enriched genes in the sporogenous mutants HM29 and HM18 (Town et al. 1987; Bradbury and Gross 1989). The cAMP signalling pathway also effects a rise in intracellular c a 2 + ,and studies using c a 2 + and calmodulin antagonists all suggest a role for this ion in gene expression (Coukell and Cameron 1988; Blumberg et al. 1989). Specifically, calcium antagonists have been shown to regulate the CAMP-mediated prespore and prestalk gene expression with particularly stronger effects on prespore genes (Schaap et al. 1986; Blumberg et al. 1988; Haribabu et al. 1989; Peters et al. 1991). Fosnaugh and Loomis (1991) implicated a ca2+-dependentmechanism in the regulation of the three major spore transcripts encoding SP60, SP70, and SP96 spore coat proteins. These transcripts were induced in wildtype cells by the addition of 0.1 mM c a 2 + or 20 pM cAMP (Fosnaugh and Loomis 1991). Studies using sporogenous mutants and permeant cAMP analogues suggest that these mutants possess a defective cAMP signal transduction pathway; target mutated genes might include those coding for a Gi protein affecting adenyl cyclase, the regulatory subunit of CAMP-dependent protein kinase, and (or) intracellular cAMP phosphodiesterase (Kay 1989). Wild-type dormant spores of D. discoideum and other slime molds are inhibited from germinating in the sorocarp by the autoinhibitor discadenine and the high osmotic pressure exerted by the presence of components in the


extracellular matrix (Russell and Bonner 1960; Snyder and Ceccarini 1966; Cotter and Raper 1966; Cotter 1975). This spore germination autoinhibitor is synthesized during culmination (Abe et al. 1976; Loomis 1975; Ihara et al. 1986), but it remains unclear whether discadenine is directly involved in initiating dormancy, as well as in maintaining the dormant spore state. Since autoinhibitor can deactivate heat-activated spores, it may participate both in sporulation and the reacquisition of dormancy (Cotter 1981). High osmotic pressure and ammonia are known to contribute to microcyst formation and dormancy maintenance in P. pallidurn (Choi and O'Day 1982; Klein et al. 1992). Therefore, complete requirements for the formation of stable dormant spores in D. discoideum may include high intraspore CAMP,high osmotic pressure, discadenine, and ammonia. Asexual development in D. discoideum is generally considered complete upon the formation of dormant spores (Loomis 1975; Raper 1984). This may not be precise because mechanisms regulating dormancy, spore aging, and spore germination competency may also be functioning in the maturing or developing dormant spore. In cellular slime molds, the germination of dormant spores, microcysts, and macrocysts is a developmental process which may occur in the absence of nutrients (Cotter 1981; Raper 1984). In the case of spores, the fundamental germination stages include spore activation, lag phase, spore swelling, and ultimately the emergence of amoebae (see Fig. 3 and Cotter 1981). Spore activation requires both the removal of the spore matrix containing the autoinhibitor (Abe et al. 1976) and the imposition of an external activating stimulus such as bacterial growth factors, nutrients, or other activating treatments (e.g., heat or osmotic shock) (Cotter 1981). Activated D. discoideum spores do not require exogenous nutrients, and hence, possess all endogenous carbon and energy reserves required for germination (Cotter and Raper 1966, 1968a). For instance, Cotter and Raper (1966) reported that glucose and vitamins (i.e., nicotinamide, riboflavin, biotin, thiamine, and pyridoxine) did not affect germination. Interestingly, hydrophobic amino acids were effective in triggering germination, and tryptophan (W), phenylalanine (F), and methionine (M) were the most effective (Cotter and Raper 1966). Early germination events are particularly sensitive to the external environment and completion of the germination sequence is contingent upon appropriate environmental cues such as oxygen tension, pH, temperature, and osmotic pressure (Cotter et al. 1979; Glaves and Cotter 1989). Spore activation is a critical reversible stage that mediates the transition from dormancy to the irreversible spore swelling stage (Cotter 1981). Activated spores are readily capable of responding to an adverse environment by returning to the dormant state, i.e., deactivating (Cotter et al. 1979). In contrast, the spore swelling stage represents the irreversible loss of spore resistance and deactivation properties, and germination will continue until emergence (Cotter and Raper 1968b). Wild-type spores aged 10-14 days in fruiting bodies acquire the ability to germinate spontaneously (autoactivate) when released from the spore matrix. This germination process is spore directed, density dependent, and bypasses the requirement for external activation (Dahlberg and Cotter 1978; Cotter and Glaves 1989). The spore-swelling stage of

0

1

2

3 4 HOURS

5

6

FIG. 3. The kinetics of spore gemination in Dictyostelium discoideum. The mean and standard deviation are shown for each 30-min period. 0,swollen spores; , nascent amoebae. The

semidiagrarnatic sequence of germination includes unswollen spores in lag phase at 30 min after activation, early spore swelling at 1 h, middle spore swelling at 1.5 h, late spore swelling at 2 h, amoeba emergence at 2.5 h, and postemergence after 2.5 h (after Cotter and Raper 19686). autoactivation is characterized by the secretion of an extracellular low molecular weight, phosphorylated adenine derivative, the autoactivator (Glaves and Cotter 1989), which is not CAMP nor cGMP (Cotter et al. 1990). This germination-specific factor is a potent germination stimulator which results in rapid and synchronous germination of responsive spore populations (Cotter and Glaves 1989). Harsh environmental conditions prevent autoactivation, but only marginally interfere with the response of spores to exogenous autoactivator (Glaves and Cotter 1989). Specifically, the addition of exogenous autoactivator extends germination parameters, allowing germination in otherwise unfavourable autoactivation conditions (Cotter and Glaves 1989). Autoactivation is also inhibited in the presence of autoinhibitor; however, the addition of autoactivator overrides this inhibition and promotes autoactivation (Cotter and Glaves 1989). The antagonistic actions demonstrated by autoinhibitor and autoactivator competition studies are similar to those of extracellular hormones which compete for binding to cell surface receptors (Cotter and Glaves 1989). Our working hypothesis is that autoinhibitor and adverse environmental conditions normally prevent initiation of the germination cascade by blocking the synthesis and (or) secretion of the spore autoactivator. The early metabolic events during autoactivation are remarkably sensitive to various pharmacological agents (Cotter et al. 1990). For instance, the rate of autoactivation is enhanced by ~ g and ~ delayed ' by c a 2 + (Virdy et al. 1992). In addition, adenine, adenosine, and caffeine have been found to greatly accelerate autoactivation (Cotter and Raper 1968b; Virdy et al. 1992). This germination program is entirely blocked by DTT and EDTA, and significantly delayed in the presence of CAMP and dibutyryl-CAMP (Virdy et al. 1992). Spores activated by mechanical or physical treatments do not synthesize, secrete, or respond to autoactivator (Cotter and Glaves 1989) and they are also insensitive to many of the pharmacological agents which affect autoactivation (Cotter et al. 1990; Virdy et al. 1992).


804


Interestingly, the activity of the lysosomal enzymes 0-glucosidase and trehalase is detected during early spore swelling in autoactivating spores, but in heat-activated spores it is delayed until emergence (Chan and Cotter 1982). In addition, amoebae from autoactivated spores, in contrast to nascent amoebae from heat-activated spores, are aggregation competent within 30 min of emergence (Cotter et al. 1990), and hence, appear primed for multicellular development. The autoactivator signal (autoactivation) cascade may be temporally coupled to the developmental CAMP-signalling pathway, to allow the precocious expression of the full complement of developmental genes in nascent amoebae, before or during emergence. In contrast, the protein denaturing spore activation treatments, such as heat or DMSO shock, may impair interactions between the former cascades and delay the initiation of multicellular development. The soluble autoactivator accumulates transiently in the media of autoactivating spore populations (Cotter et al. 1990) and is promptly inactivated by nascent amoebae as they emerge from swollen spores (Virdy and Cotter 1991). Our current research indicates that a cell surface protein, with an electrophoretic profile resembling that of alkaline phosphatase/5 ' -nucleotidase (Armant and Rutherford 1981, 1982; Bhanot and Weeks 1985), inactivates this germination signal compound (Virdy et al. 1992). Autoactivator accumulation also appears to be initially regulated by a sporespecific inactivating factor which is present before spore swelling (Virdy and Cotter 1991). We are currently attempting to elucidate the early germination signalling cascade (autoactivation) using the SG1 and SG2 D. discoideum premature maturation mutants. The germination phenotype of these two strains and their germination requirements have been characterized, and most notably, their autoactivation competency bypasses the extensive aging and maturation times required by wild-type strains (Cotter and Dahlberg 1977; Dahlberg and Cotter 1978; Cotter and Glaves 1989). Young wild-type NC4 spores (1-3 days old) do not respond to or produce autoactivator (Cotter and Dahlberg 1977). Although they do not autoactivate, external activating treatments, such as heat or DMSO shock, will initiate germination (Cotter and Dahlberg 1977; Dahlberg and Cotter 1978). In contrast, aging wild-type spores gradually acquire the ability to respond to autoactivator, and after 10-14 days of aging in the sorocarp, strongly respond to and autonomously produce this factor (Dahlberg and Cotter 1978). The concurrence of autoactivation competency and spore development implies the existence of late spore maturation genes (Glaves and Cotter 1989). Two adenyl cyclase genes of D. discoideum have been isolated, sequenced, and disrupted (Pitt et al. 1992). This research confirms the expected requirement of aca (adenyl cyclase - aggregation), coding for the transmembrane protein ACA, for cellular aggregation and the initiation of multicellular development; acg (adenyl cyclase germination) appears to code for a transmembrane protein (ACG) and the message is produced after fruiting body formation and during early spore germination. The acg gene has been disrupted by the insertion of a G418 antibiotic resistance gene (Pitt et al. 1992). The disruption of this late spore maturation gene results in spores which, in contrast to those of SG1, SG2 and aged NC4, do not autoactivate in a concentration-dependent manner and are insensitive to both

autoactivator and autoinhibitor (K. Virdy, T. Sands, D. Cotter, G. Pitt, and P. Devreotes, unpublished). A number of studies have already shown the dependency of prespore development and spore formation on internal CAMP(Kay et al. 1988; Kay 1989; Riley and Barclay 1986, 1990; Riley et al. 1989; Maeda 1992). We propose that the subsequent developmental stage of spore dormancy is also dependent on intraspore cAMP signalling and continued cAMP production. We further postulate that microcyst and macrocyst dormancy is also maintained by a similar mechanism. During terminal sporulation, the functioning CAR4/ACA pathway (Saxe et al. 1991) can be superseded by one involving ACG for the maintenance of long-term dormancy. This spore cAMP cascade may regulate protein kinases (e.g., pk-A and (or) PK2), phosphorylation of spore proteins (Anjard et al. 1992; Harwood et al. 1992), and hence, spore dormancy. Other cascade enzymes (e.g., phosphoprotein phosphatases), which may include a vanadate-sensitive class (Cotter et al. 1990), could regulate this pathway by opposing CAMP-directedkinase reactions. A net increase in phosphatase activity may contribute to the termination of dormancy and the initiation of the germination cascade. Mutant D. discoideum strains may contain lesions in this putative internal cAMP signal transduction pathway involving ACG, protein kinases, phosphoprotein phosphatases, or phosphodiesterases and may form supersensitive or superdormant spore strains (Cotter and Raper 1968c; Kessin and Newel1 1974; Ennis and Sussman 1975; Dahlberg and Cotter 1978; Ennis 1983). The N-terminal putative extracellular domain of ACG is sufficiently large to serve as its own receptor for specific ligands (Pitt et al. 1992). Perhaps the autoinhibitor and (or) autoactivator bind or compete for binding to this extracellular domain. In this scenario, bound autoinhibitor would maintain ACG activity generating internal CAMP,while the binding of autoactivator could inactivate ACG, terminating cAMP production. In the case of competition, the autoactivator may have a higher binding affinity for this binding domain than autoinhibitor. Macko (1981), in his review of germination inhibitors and stimulants in fungi, has found that many of the natural inhibitors and stimulators are structural analogues. This appears to be true for D. discoideum, because both the autoinhibitor and autoactivator seem to be adenine derivatives. Therefore, competition between these analogues for a putative common binding site is not surprising. Our model suggests that spore dormancy requires energy and perhaps the decline of a key reserve during spore aging results in a reduction of spore cAMP below a critical threshold which may allow germination. In our pharmacological studies (described above), we have employed some agents known to interfere with the classical aggregation cAMP cascade. For instance, caffeine and adenosine which accelerated the germination program are known to impair normal cAMP relay, by inhibiting CAMP-inducedactivation of adenyl cyclase activity (Brenner and Thoms 1984), and cAMP receptor binding (Newel1 1982; Newel1 and Ross 1982b), respectively. More strikingly, the presence of CAMP, dibutyryl-CAMP,or c a 2 + blocked or delayed autoactivation. These results suggest that agents which antagonize the cAMP signalling pathway (maintaining dormancy) temporally accelerate autoactivation; in contrast,


CAMP-signallingagonists which are normally required for spore formation, delayed or inhibited this germination process. A previous study suggesting that cAMP enhances germination under certain conditions (Katilus and Ceccarini 1975) may have resulted from the hydrolysis of cAMP to adenosine, the latter of which promotes germination (Cotter and Raper 1968~; Virdy et al. 1992). Our hypothesis is consistent with results showing the relationship of spore dormancy to continued intraspore CAMPsignalling and of spore autoactivation to a decline in spore CAMP.The autoactivation cascade is initiated by a small number of earlygermination competent spores; these spores secrete the initial pulse of autoactivator to initiate this autocatalytic germination cascade (Cotter and Glaves 1989). In these precocious autoactivation competent spores, the ACG protein may oscillate between active and inactive conformations and could inactivate in the absence of the autoactivator signal. The disruption of acg, which does not block sporulation, could affect spore dormancy by the following mechanisms: (0 altered spore cAMP production, (ii) impairment of long term dormancy and spore viability, and (or) (iii) initiation of germination simply by a reduction of osmotic pressure in the sorus. Wild-type spores aged for longer than 14 days in the fruiting body begin to lose viability. Perhaps autoactivation in wild-type spores is a final germination attempt to search for a more nutrient-rich environment before the aging process results in loss of spore viability (Glaves and Cotter 1989). Sustained ATP levels have been observed by 3 1 ~ analysis ~ in both ~ ~dormant spores and microcysts incubated under aerobic conditions (Klein, G., et al. 1988, 1992). The 3 1 ~ peak - of~ ATP ~ in ~dormant spores was maintained metabolically, since it disappeared completely upon anaerobiosis (Klein, G., et al. 1988). Interestingly, only purine nucleoside triphosphates (ATP, GTP) are represented in significant amounts in spores, and the amount of pyrimidine nucleoside triphosphates (CTP, UTP) were found to be at least 15-20 times lower (Klein, G., et al. 1988). Similar results were obtained using HPLC extracts from dormant spores (Hamer and Cotter 1983). This is not the case in vegetative amoebae where the purine/pyrimidine ratio was found to be 2.7 (Satre et al. 1986) or in nascent amoebae arising from germinated spores (Klein, G., et al. 1988). The spore ATP and GTP can easily support the proposed basal metabolic activities in dormant spores and interference with spore metabolism under aerobic conditions may shorten or terminate dormancy. We have found that when wild-type spores were freed of autoinhibitor and allowed to age 5-6 days in an anaerobic environment, they rapidly germinated after reaeration; however, during germination, exogenous autoactivator was not detected (D. Cotter and K. Virdy, unpublished). This is consistent with the findings of Glaves and Cotter (1989), which indicated that the production and secretion of autoactivator was very sensitive to oxygen tension. It is probable that the key enzymatic step(s) in the synthesis and secretion of autoactivator is exquisitely aerobic (Glaves and Cotter 1989). Heat or osmotic shock activation of young wild-type spores might temporarily interfere with ATP synthesis, cAMP production, and (or) protein kinase activity, stimulating such spores

to initiate germination (Cotter 1981). Deactivation (dormancy reacquisition) during adverse environmental conditions may be accomplished by the reestablishment of the spore cAMP generating pathway, rephosphorylation of spore proteins, and the concurrent inhibition of phosphoprotein phosphatase reactions. It is interesting that unlike many lysosomal enzymes in dormant spores which are inactivated during heat-induced activation (Tisa and Cotter 1980), acid phosphatase is highly heat resistant and alkaline phosphatase activity is reduced only by one third (Tisa and Cotter 1979). These heat-resistant phosphatases are possible candidates for the class of phosphoprotein phosphatases capable of opposing CAMP-directed kinase reactions and thus regulating the germination cascade. Clearly, the deactivation (dormancy sustaining) cascade must become fully active during the brief lag phase, after spore activation but long before the irreversible step in the germination cascade. In addition to the fruiting body stage, the transcription of acg has also been observed directly after DMSO-induced spore activation, during the lag phase of germination (Pitt et al. 1992). This may represent an attempt to regenerate spore cAMP levels and reestablish dormancy after such a harsh activating treatment. Spore deactivation occurs most rapidly at a transition point (fail-safe point) between the end of the lag phase and the initiation of spore swelling. The longer spores remain at the end of the lag stage (subjected to autoinhibitor or harsh environmental conditions such as low 0 2 , high osmotic pressure, high or low temperature, or high or low pH), the greater the percentage of spore deactivation (Cotter et al. 1979). Superdormant spore mutants, with the ability to resist heat- or DMSO-induced activation, have been isolated (Cotter and Raper 1968~;Ennis 1983); other mutants include those with the ability to rapidly deactivate at the fail-safe point (Ennis 1983). In contrast, SG1 and SG2 mutants are supersensitive to all activation treatments and do not deactivate rapidly enough to prevent their swelling and death when subjected to harsh environmental conditions (Cotter and Dahlberg 1977). These mutants also have larger territory sizes and recruit more cells into their fruiting bodies than their parental strains NC4 and V12, respectively (Cotter and Dahlberg 1977). A partial defect in the regulation of cAMP phosphodiesterase activity throughout the life cycle might explain most SG1 and SG2 germination and developmental features. Many of the common D. discoideum strains in use today have undergone genetic divergence (Kuspa et al. 1992) and harbour a sizable proportion of spore germination mutants (Cotter and Dahlberg 1977). To minimize the effects of genetic drift, it is important to repeatedly examine postsporulation development among the various strains. We also suspect that the germination cascade leads to the activation of acidosomes (see Padh et al. 1991), which then fuse with the lysosomal/endosomal vacuolar system and activate resident lysosomal enzymes such as trehalase-1 and the developmental CP48 and CP43 proteases (Jackson et al. 1982; Klein et al. 1990; Gupta and Cotter 1990; North et al. 1990a; North and Cotter 1991~).Presumably, this event precedes the spore-swelling stage, the hydrolysis of trehalose, and the utilization of the excess amino acid reserves including the large pools of Q, E, N, etc. (Ennis 1981; Klein et al. 1990). Microcyst germination similarly utilizes trehalose reserves and these same pools of amino acids


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BIOCHEM. CELL BIOL. VOL. 70. 1992

(Ennis 1981; Klein et al. 1992). Protein turnover and the utilization of amino acid pools may be used for de novo protein synthesis for germination-specific proteins (i.e., cellulases). Cellulase(s) are involved in the degradation of the spore wall after the initiation of spore swelling (Rosness 1968; Cotter et al. 1969; Jones et al. 1979; Hemmes et al. 1972; Klein et al. 1990; Ennis et al. 1991). Cellulose hydrolysis is marked by the transient accumulation of cellobiose in both spore and microcyst germination culture broths (Klein et al. 1990, 1992). Concurrently, lipid precursors are incorporated during the synthesis and turnover of the expanding membrane surfaces (Cotter et al. 1969; Maeda and Takeuchi 1969; Gregg and Badman 1970; Klein, G., et al. 1988, 1990). The emergence of an amoeba with a de novo vegetative complement of actin, ribosomal proteins, and other enzymes (Macleod et al. 1980; Ramagopal and Ennis 1982; Singleton et al. 1991) allows the nascent amoeba to initiate growth in the presence of nutrients or a microcycle in the absence of nutrients, and to participate in the formation of a second fruiting body (Cotter and Raper 1968c; Cotter et al. 1990). In summary, the contiguous developmental stages of D. discoideum sporulation, spore dormancy, and spore maturation may be based on a common CAMP-generating intraspore metabolic cascade. If this proves to be the case, then multicellular spore development and differentiation, beginning with cell aggregation and culminating with spore maturation, would be primarily cAMP directed. Intuitively, the disruption of this cAMP intraspore cascade could trigger the germination program, thereby reinitiating the life cycle. If microcyst and macrocyst formation and dormancy are also driven by a similar pathway, then this would be consistent with the frugality with which cellular slime molds utilize and recycle key intra- and extra-cellular signal molecules in their signalling and metabolic pathways. Other implications of ammonia assimilation in developing cells In mammals, ammonia is particularly toxic to the central nervous system and glutamine synthetase appears to serve a detoxification role by using free ammonia in the formation of glutamine. This scavenger system temporarily removes free ammonia from the cellular pool, but as needs arise, the resulting glutamine reserve can be used as a source for amino groups (Moat 1979). Glutamine, a key compound in nitrogen metabolism, is the most abundant free amino acid in the human body and is involved in the transport of carbon and nitrogen between tissues (Smith 1986). The glutamine cycle links catabolism and anabolism, and thus, buffers variations in the nutrient supply, drives energy generation and carbon flow for optimal cell function (Mora 1990; Pisoni and Thoene 1991), and may also regulate protein degradation (see Watford 1989; MacLennan et al. 1988). In Saccharomyces cerevisiae, only 10 amino acids accumulate in the central vacuole/lysosome; Q and N are among the basic and neutral amino acids primarily stored (Klionsky et al. 1990). Mutants of S. cerevisiae with defective vacuolar function do not accumulate these amino acids and do not sporulate (Kitamoto et al. 1988). In P. pallidum and D. discoideum, ammonia is produced in high amounts as protein laden vegetative cells are induced to turnover their proteins for the formation of either microcysts or fruiting bodies. Ammonia may also be impor-

tant during the sexual life cycle, since extensive cell degradation and protein turnover parallels the appearance of new cysteine proteases during aggregation and the maturation of macrocysts (Lydan and O'Day 1990; North et al. 1990b; North and Cotter 1991a, 1991b). In dormant spores and microcysts, 1 3 ~ analysis ~ has~revealed ~ that, of all the amino acids, free glutamine (Q) was present in the highest concentration (Klein et al. 1990, 1992). Possibly, this reserve results from a cellular slime mold version of the classical ammonia detoxification system. As mentioned previously, the high concentration of lysosomotropic Q may also contribute to the deactivation of prespore and spore lysosomal proteins such as the developmental CP48 and CP43 enzymes. As lysosomal proteases become less active in prespore protein turnover, such cells may become more dependent on the assimilation of NH3 with glycolytic and TCA precursors to form amino acids for de novo protein synthesis. In addition to random mutation and transposition events, this ammonia assimilation pathway may have effected the selection of a codon bias favouring the nine amino acids which accumulate to the highest levels in dormant spores. We describe these amino acids as SAAR. In our analysis of D. discoideum polypeptide compositions (deduced primarily from published gene sequence data), we have selected P as a SAAR. Since only P and R analogues have been shown to be effective in blocking spore germination (Cotter and Raper 1970), one could rationalize that both P and R could also be included in the SAAR collection; however, R at concentrations of 3 x l o p 2M resulted in abnormal fruiting bodies in which the proportion of spores to stalk cells was greatly increased and the spores were more rounded than normal (Loomis 1975). Because spores never accumulate this level of R and are normally capsule shaped, we omitted the inclusion of R, but not P. Therefore, the SAAR collection includes Q, K, G, E, T, P, A, N, D, and S (see Table 1). Interestingly, the amino acids which are essential for axenic growth are also the same amino acids required to block aggregation (Marin 1976). All 10 non-SAAR fit this category; however, the SAAR G, SAAR T, and SAAR K may partially belong in this category. It should be noted that G increased less than fivefold from vegetative growth to sporulation when measured by ' 3 ~ (Klein et ~ al.~ 1990) ~ and not at all in another study (Ennis 1981). Thus, G may be a marginal member of the SAAR collection of amino acids. Marin (1976) has reported that D. discoideum is capable of both synthesizing and degrading D, E, A, N, S, Q, and P; we believe that his original data indicate partial abilities with respect to T and K as well. In conclusion, it appears that dormant spores only accumulate free amino acids which they are capable of synthesizing, but not those which are essential for axenic growth. The 10 SAAR are generally, but not exclusively, hydrophilic according to the scale of Kyte and Doolittle (1982). However, according to a scale by Rose et al. (1985) all the SAAR are, without exception, classified as hydrophilic or neutral. It is interesting to note that during the life cycle of D. discoideum hydrophobic amino acids stimulate spore germination (Cotter and Raper 1966), are essential for axenic growth, and block aggregation (Marin 1976). In contrast, hydrophilic and neutral amino acids (SAAR) are synthesized by vegetative cells, accumulate as


free amino acids in spores (Klein et al. 1990), and therefore should occur in a higher frequency in developmental polypeptides (e.g., sporulation and germination polypeptides). In our analysis of D. discoideum polypeptides, the increase in SAAR can be represented by any of the following: (I] an overall compositional increase, (ill a compositional increase unique to a domain, (iil] as homopolymeric repeats in specific domains, (iv) as heteropolymeric repeats in specific domains, and (v) combinations of the above. Because these SAAR include most of the classically defined hydrophilic amino acids and no protein is entirely hydrophilic, there are constraints lowering the overall SAAR composition which are most apparent in (I] membrane-bound proteins because they contain hydrophobic membrane spanning segments, (ill highly conserved proteins with sequences required for functional and structural integrity, (iio extracellular proteins with a high C content for disulfide cross-linking, and (iv) signal sequences in polypeptides which do not contribute to the final functional polypeptide. Sharp and Devine (1989) by using a multivariate statistical analysis demonstrated that the greatest variation among genes in D. discoideum involved a relative usage of a particular subset of codons, many ending with cytosine. We have used Sharp and Devine's data of the total codon frequencies for 56 D. discoideum genes to calculate an average percentage of SAAR per polypeptide of 59.6%. This average was then used comparatively in our analyses of specific groups of D. discoideum polypeptides. Theoretically, if a peptide contains 10 amino acids in which five are SAAR (50% SAAR), then two extreme cases for the distribution of SAAR within the peptide are (0 a SAAR always followed by a non-SAAR or a non-SAAR always followed by a SAAR, and (ill a block of five SAAR either at the N-terminus or C-terminus of the peptide. If we calculate the SAAR which occur contiguously, then in the first extreme case (I] the SAAR length is 1, while in the second case (ii) the SAAR length is 5. In some of our comparisons, both the percentage of SAAR per polypeptide and the SAAR length were considered. The analysis included both vegetative proteins and those synthesized throughout the life cycle. For instance, the SAAR composition of the vegetative V1 ribosomal protein is 54% (see Table 1) and the SAAR positioning within the sequence is shown in Fig. 4. The longest SAAR length is only 6. Similarly, the vegetative V14 polypeptide also has a SAAR composition below the average 59.6%, with the longest SAAR length being 8. In D. discoideum, 14 of the 17 actin genes are transcribed throughout the life cycle; however, a single predominant amino acid sequence is obtained from the purified protein (Vanderchove and Weber 1980). Again, the SAAR percentage is lower than the average 59.6% (see Table 1). Highly conserved actin-binding proteins such as the capping proteins (Hartmann et al. 1990) also have average SAAR percentages. Cyclophilin is identical to a peptidyl-prolyl cis-trans isomerase which catalyzes the cis-trans isomerization of proline imidic peptide bonds and can accelerate the refolding of proteins in vitro (Barisic et al. 1991). This gene shares 64% homology with the human gene (Barisic et al. 1991) and its SAAR composition is slightly higher than the average. The preaggregation genes for the secreted cAMP phosphodiesterase (pdsA) and phosphodiesterase inhibitor (pdiA) code for proteins with an approximate average SAAR percentage (62.5 and 6 1.5% ,

20

PsA

S A A R Length FIG. 4. A SAAR compositional analysis of three proteins. The percentage of SAAR per polypeptide is plotted against the SAAR length. Note that the total percentage of SAAR per polypeptide is calculated by summing the individual SAAR percentage for each SAAR length. The upper plot represents the ribosomal protein VI, the middle plot represents the mature form of the prespore protein PsA, and the lower plot represents the prespore protein 3F.

respectively) without the N-terminal signal sequences included in the analysis. Cells containing the surface protein gp80 demonstrate EDTA-resistant binding during cellular aggregation (Loomis 1982). The mature protein contains three distinct segments: a terminal domain involved in cell-to-cell contact; a stalk domain high in S, T, and P repeats; and a C-terminal hydrophobic membrane spanning anchor (Siu and Kamboj 1990). Without the N-terminal signal sequence, gp80 contains almost 66% SAAR (see Table 1). In contrast, the cAMP cell-surface receptor CAR1 (Saxe et al. 1988) has a SAAR composition of only 51.9%; it is a polypeptide with seven transmembrane domains and is largely divided into SAAR-positive and SAAR-negative regions. Similarly the membrane proteins ACA and ACG have average SAAR values, but are enriched with repeats of N (Pitt et al. 1992). Proteins such as those of the ras family are highly conserved


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BIOCHEM. CELL BIOL. VOL. 70. 1992

from organism to organism and both Ddras and DdrasG have average SAAR percentages (see Table 1). A compositional analysis of CP1 and CP2 suggests that they are proteins typical of the late aggregation stage of development; they have no SAAR length greater than 9. If the signal sequence is not included in the calculations, the SAAR percentage of CP1 increases to 60.8 and the removal of another 99 N-terminal residues increases the SAAR percentage only to 60.9. Interestingly, the CP2 polypeptide without the signal sequence has a higher SAAR percentage (65.1). Cell surface proteins such as PsA are produced by prespore cells (Early et al. 1988a)and show molecular weight polymorphisms among different strains of D. discoideum (Grant et al. 1985; Early et al. 19886). Length polymorphisms of approximately 2 kDa result from the presence of C-terminal variable length repeats of a glycosylated tetrapeptide sequence, rich in P and T (21% total T in the protein) (Early et al. 1988a, 1988b). The overall SAAR percentage (64.3) is higher than average, and when the signal peptide is removed, the percentage increases to 66.4. The mature protein without the C-terminus results in a further increase to 69.6%. Since the length polymorphism of this surface protein does not seem to affect its function, we agree with Early et al. (1988a) in that PsA is an example of a prespore protein with genetic variation resulting from unequal crossing-over, and we further suggest that the PTVT repeats may reflect consumption and detoxification of excess ammonia (see Fig. 4). This length polymorphism may be a common theme in D. discoideum since the monoclonal antibody MUDI, as well as MUDSO, recognizes PsA, but also cross-reacts with other developmentally regulated polymorphic glycoproteins (Grant et al. 1985; see Smith et al. 1989). The genes ecmA and ecmB expressed in pstA and pstB cells after tip formation contain 41 and 70 repeats of a 24-residue sequence, all containing high C (Ceccarelli et al. 1987; Williams et al. 1987). For every three C in the repeat, there are four SAAR A, four SAAR T, two and a half SAAR S, two SAAR N, nearly two SAAR K, and two SAAR P, giving the proteins an overall SAAR percentage near 70. The secretion of these proteins by prestalk and stalk cells, while contributing to the construction of the developing slug and sorocarp might also serve to lessen the ammonia load of the cell mass. In addition, the fact that the two genes code for such similar proteins suggests that the duplication of genes coding for extracellular proteins may have a limited evolutionary significance. Annexin VII (synexin) is presumably a c a 2 + channel protein which is highly conserved in eucaryotes including man; it is found throughout the D. discoideum life cycle (Doring et al. 1991). The N-terminus of the protein is longer than that of other annexins and contains 12 perfect repeats of a GYPPQQ motif, as well as a few imperfect repeats (Doring et al. 1991). These repeats are not involved in c a 2 + binding. While the N-terminal repeats involving 85 amino acids have a total SAAR percentage of 83.0, the rest of the protein has a SAAR percentage of 58.9, with an overall SAAR percentage of 65.1% (Table 1). The sequence GYPPQQ is also found repeated in the CAMP-bindingprotein CABPl (Grant and Tsang 1990). The transcription levels of capA increase three- to fourfold during development (Grant and Tsang 1990). The encoded CABPl protein is composed of two subunits, with the smaller (CABPlB) missing 37 amino acids near the

N-terminus (Grant and Tsang 1990). Interestingly, the C-terminus has 47% identity (86% functionally conserved substitutions) with polypeptides encoded by a bacterial plasmid for tellurium anion resistance. The N-termini of CABPlA and CABPlB are rich in SAAR P, SAAR Q, and SAAR G, and contain a large block of GYPPQQ motif repeats. Our analysis shows that almost three quarters of the amino acids are SAAR and the excess SAAR P and SAAR G are frequently in separate blocks containing two to five contiguous residues which are then part of larger repeated sequences. Antisera against CABPl cross-react with the regulatory subunit of CAMP-dependent protein kinase, suggesting that they are part of a gene family of CAMP-binding proteins (Tsang and Tasaka 1986). Is it possible that annexin VII, CABPI, and CAMP-dependent protein kinases all bind c a 2 + ? When strains of V12 are compared with strains of NC4, the molecular masses of CABPlA (43 kDa) and CABPlB (38 kDa) appear reduced by approximately 2 kDa (Tsang et al. 1988). This may be another example of length polymorphism involving repeated blocks of SAAR; there are many repeated regions in the capA gene capable of resulting in unequal crossing-over leading to deletions and helping to explain the observed strain variations. The possibility of SAAR length variations mediated by transposable elements should also be considered. The SAAR percentage of Dd3 1 (51.1) occurs because of the SAAR coding differences by two exons of spiA: the first half of exon 1 codes for high SAAR content with SAAR lengths of 13 and 11 and the second half codes for low SAAR content. Exon 2 also codes for very low SAAR, resulting in an overall average similar to vegetative genes (see Table 1). The spore-specific gene for this membrane protein is transcribed during sporulation and transcripts may persist in dormant spores (Richardson et al. 1991). Spore coat proteins account for 5-9070 of the total protein synthesis during culmination (Orlowski and Loomis 1979; Wilkinson and Hames 1983). Following the signal sequence, the N-terminus of the spore coat protein SP60 contains four hexapeptide repeats of the sequence GDWNN. The remaining sequence is exceptionally rich in C and D, constituting about 11 and 18%, respectively, of the mature protein (Fosnaugh and Loomis 1989a; Widdowson et al. 1990). The rest of the protein is also largely composed of amino acid repeats with almost 65% of the last 85 residues being D, N, E, or Q. Together, the hydrophobic amino acids along with the high C levels result in an overall SAAR percentage of 60.1. Widdowson et al. (1990) found that the 5' upstream noncoding region has both a C-rich and several AC-rich sequences with a core region similar to those observed in the promoter region of the pspA gene (Early and Williams 1989). Widdowson et al. (1990) speculated that much of the gene has been derived by intragenic duplication. A second spore coat protein, SP70, has a high C content similar to SP60 and contains 17% T and S, with five copies of the sequence GSHTTTGGSTT in the center of the mature protein (Fosnaugh and Loomis 1989a). A third spore coat protein, SP96, also has a high C and extremely high T and S levels; the last 190 amino acids are 65% S and T, with the remaining being primarily A and P repeats. The SP96 protein shares repeats with both SP60 and SP70, and it appears that the three genes encoding for these proteins diverged from duplications of a common precursor (Fosnaugh and Loomis 1989b). The three spore coat genes


REVIEWS / S Y N T H ~ S E S

may share regulatory mechanisms since they are expressed coordinately under a variety of conditions (Fosnaugh and Loomis 1989b; Tasaka et al. 1992). The high C content of the three proteins may be involved in extensive cross-linking of the proteins by disulfide bonds during or after secretion from prespore vesicles (Fosnaugh and Loomis 1989a, 1989b; Widdowson et al. 1990). Another spore gene of unknown function, EB4, contains three exons coding for high C content which is characteristic of spore coat proteins. The sequence of the gene is incomplete, but there are C repeats and 17 hexameric repeats with the consensus sequence AQH--AKP (Hildebrandt et al. 1991). There are also short repeats of 3 to 6 Q or A in the third exon. Disruption of the EB4 gene has no apparent effect on the organism (Hildebrandt et al. 1991); perhaps the spore coat genes are largely redundant and their individual roles as ammonia sinks can be accommodated by other members of the family. The late developmental spore gene 3F does not encode high C levels (Widdowson et al. 1989). The last 95 C-terminal amino acids are all SAAR giving the protein a SAAR percentage of 79.1 (Fig. 4). A family of spore germination proteins containing the repeat TETP are unique to sporulation and spore germination (Giorda et al. 1990; Ennis et al. 1991). The N-terminus of pRK270-6 cellulase is similar to that of avocado cellulase; the C-terminus, containing high T, P, and S as well as repeats of the above tetrapeptide, gives the entire protein a SAAR percentage of 68.9. Also, pRK270-11 contains a high frequency of these same tetrapeptide repeats, resulting in a SAAR percentage of 79.1, or 87.5 without the putative signal sequence. We also note sequences such as PTTT, PSSS, and PTPTPS in the above two polypeptides. Other members of the 270 family defined by the tetrapeptide repeat are not transcribed during spore germination or during the rest of the life cycle (Giorda et al. 1990; Ennis et al. 1991). Ennis et al. (1991) have noted that the concentration of the amino acids making up the TETP repeats in this family of proteins is twofold that of the average D. discoideum protein, and that proteins such as gp80 (Noegel et al. 1986; Siu and Kamboj 1990) and PsA (Early et al. 1988a, 19886) contain similar (i.e., TPTP and PTVT) but not identical repeats (Ennis et al. 1991). Other proteins synthesized during germination may either aid the transition from dormant spores to vegetative cells or be primarily vegetative proteins; these latter germination proteins may have SAAR percentages more similar to vegetative proteins than to sporulation proteins (see pLK109, V1, and V14 in Table 1). Similarly, ubiquitin synthesized during spore germination does not differ from ubiquitin in other stages of the life cycle (Ennis et al. 1988). It is interesting to note however, that the V4 gene is transcribed upon late emergence and also during the growth phase; this transcript undergoes a rapid 1000-fold decrease in abundance during the preaggregation phase of development (Singleton et al. 1987, 1991). The putative 160-residue polypeptide (McPherson and Singleton 1992) has a SAAR percentage of 63.8, but the last 21 C-terminal residues are pure SAAR with high A and Q (see Table 1). Protein domains that are destined for 0-glycosylation are rich in S, T, and P. The consequences of 0-glycosylation are resistance to protease degradation and the formation of random coils (Jentoft 1990). The random coils frequently separate functional globular domains in proteins (see Gilkes et al. 1991). In cellulases, the globular catalytic domain may

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be separated from the globular cellulose binding domain by an 0-glycosylated random coil domain. This barbell arrangement is found in the D. discoideum cellulase (pRK270-6), while the membrane protein gp80 has a lollipop on a stick motif. The 0-glycosylated stick portion of polypeptides must have a minimal length for function. We contend that there may be less constraint on the maximum length of the 0-glycosylated repeats in secreted proteins, such as the sheath and the spore coat proteins, which must bind to cellulose. During spore germination, the binding of pRK270-6 cellulase to the spore wall cellulose may aid in the hydrolysis of the polymer. The most active cellulases are produced by organisms such as Trichoderma reesei; the cellulases bind to the insoluble cellulose substrate with either a protruding N-terminal or C-terminal domain rich in 0-glycosylated T and S (Stahlberg et al. 1988). Sequences such as PTTTT and TPPTS occur in the domains of the T. reesei cellulases. We find PSSS and PTTT sequences in the spore coat proteins SP70 and SP96, 3F, and the D. discoideum cellulase pRK270-6. Interestingly, antibodies recognizing high T and P cross-react with many of the prespore and spore coat proteins (West and Erdos 1990; Champion et al. 1991). SAAR repeats may have originated from repeated regions of conserved proteins, such as annexin VII or p24 (Doring et al. 1991; Greenwood and Tsang 1991, 1992). The addition or deletion of these repeats may have little consequence on the activity of such proteins. In fact, the disruption of annexin VII itself has no apparent effect on the growth or development of D. discoideum (Doring et al. 1991). We recognize that the length polymorphisms of PsA and PsB have a relationship to developmental time (Smith et al. 1989). Hence, those strains with higher numbers of SAAR repeats may develop more rapidly. Since PsA and ST310/ST430 are secreted and bind to the cellulose in the migrating slug, the longer the migration period, the more the slug commits its amino acid reserves to these three proteins; i.e., there may be no maximum or minimum amount of these proteins required for development of dormant spores. If slug migration time is partially dictated by ammonia gradients, then these three proteins may serve as suitable ammonia sinks. Therefore, the greater the number of SAAR repeats in these proteins, the more rapidly the NH3 gradient is dissipated with their secretion and the more rapid the development. There are other interesting homopolymer SAAR repeats in D. dkcoideum. A gene encoding a putative serine/threonine specific protein kinase, DdPK2, seems related to mammalian and yeast CAMP-dependentprotein kinases; the C-terminal catalytic domains show over 50% sequence homology. However, the N-terminus of the PK2 protein appears unrelated to any known protein kinase and contains poly(QTN) encoded by (AAC), repeats (Anjard et al. 1992). A prespore gene, D7, shares no homology with other prespore genes, but contains CAA repeats, making the predicted protein product very Q-rich (Blumberg et al. 1991). Another Q-rich protein is a calmodulin-dependent protein phosphatase (Kincaid et al. 1991). In D. discoideum, Q-rich proteins may have evolved from bacterial regulatory and sensory transduction proteins which are rich in Q, R, D, S, and P (Wooton and Drummond 1989) and may be related to the CREB family of Q-rich eucaryotic transcription activators (Brindle and Montminy 1992). Generally, it appears that there is an increase in the frequency of SAAR

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in polypeptides involved in development i.e., during the transition from vegetative growth to sporulation and early germination (see Table 1 and Fig. 4). Conclusions The cellular slime molds are particularly amenable to studies of the regulatory mechanism(s) governing growth and development in eucaryotic cells. In D. discoideum, controlling mechanisms at each stage in the life cycle(s) are exquisitely regulated by a number of key factors including CAMP, ammonia, DIF, discadenine, and autoactivator. In this review, we have detailed the established as well as the potential roles of these selected factors during development, cell differentiation and patterning, spore dormancy, and germination. In D. discoideum, the role of ammonia in the life cycle has not been well defined; however, evidence to date suggests that it functions in the timing of cellular development. The pattern of ammonia production and utilization may be a hallmark of cell fate determination effected by the position of a cell in the cell cycle. The gradient(s) established between ammonia-producing and ammonia-consuming cells may drive the transition from preaggregation to terminal differentiation. Ammonia triggers both microcyst and fruiting body formation in P. pallidurn and D. discoideum, respectively. Our source and sink model of ammonia assimilation by prespore cells predicts an accumulation of free hydrophilic and neutral amino acids (i.e., SAAR) in these cells, particularly that of glutamine (see Fig. 2). This increase in lysosomotropic glutamine may affect lysosomal protease activity so that protein degradation is minimized and the synthesis of spore-specific proteins is facilitated. In contrast, the decline of free glutamine during germination is probably involved in reestablishing lysosomal proteolytic mechanisms required for germination. Therefore, ammonia recycling appears to be coupled to glutamine cycling and the regulation of nitrogedprotein turnover during sporulation. We have shown that there is an increase in the frequency of SAAR in developmental proteins. Free hydrophilic amino acids accumulate during development; alternatively, hydrophobic amino acids stimulate germination, are required for axenic growth, and block aggregation. Thus, the increase in SAAR both as free amino acids and in developmentally significant polypeptides may be a distinguishing feature of differentiation. In effect, we suggest that the developmental role(s) of ammonia have been truly underestimated, since it regulates aggregation, slug migration, timing of differentiation (cell sensitivity to DIF and CAMP), cell proportioning, sorocarp orientation, lysosomal enzyme activity, dormant spore amino acid reserves, and the frequency of SAAR in developmental (sporulation and germination) proteins. The amino acids (free and in proteins) appear to be significant sinks for this volatile morphogen, which must be effectively removed for cell-specific sensitivity to other morphogens. The nucleotide cAMP also has multiple roles in the life cycle, but the primary functions may be to serve as the chemotactic signal, to promote prespore gene expression, to induce sporulation, and to maintain dormancy. Both the CAMP- and ammonia-directed pathways may serve to regulate D I F production and cell-specific response. The pro-

duction of cAMP in spores may be regulated by discadenine and (or) autoactivator; temporary interruption of its production may trigger spore germination. Ultimately, further research on the molecular mechanism(s) of action of these developmental morphogens will improve our understanding of the regulation of growth and development in eucaryotic cells. Acknowledgements The work was supported by a NATO Collaborative Research Grant and by grants from the Natural Sciences and Engineering Research Council of Canada. We thank Dr. Ira N. Feit for stimulating discussions and Mrs. J. Durocher for her invaluable help in the preparation of the manuscript. Abe, T., and Maeda, Y. 1986. Induction of macrocyst germination in the cellular slime mould Dictyostelium mucoroides. J. Gen. Microbiol. 132: 2787-2791. Abe, H . , Uchiyama, M., Tanaka, Y., and Saito, H. 1976. Structure of discadenine, a spore germination inhibitor from the cellular slime mold, Dictyostelium discoideum. Tetrahedron Lett. 42: 3807-3810. Abe, K., and Yanagisawa, K. 1983. A new class of rapidly developing mutants in Dictyostelium discoideum: implications for cyclic AMP metabolism and cell differentiation. Dev. Biol. 95: 200-210. Abe, T., Maeda, Y., and Iijima, T. 1988. Transient increase of the intracellular c a 2 + concentration during chemotactic signal transduction in Dictyostelium discoideum cells. Differentiation (Berlin), 39: 90-96. Aerts, R. 1988. Changes in cytoplasmic pH are involved in the cell type regulation of Dictyostelium. Cell Differ. 23: 125-132. Aerts, R., Durston, A., and Konijn, T. 1987. Cytoplasmic pH at the onset of development in Dictyostelium. J. Cell Sci. 87: 423-430. Albe, K.R., and Wright, B.E. 1992. Systems analysis of the tricarboxylic acid cycle in Dictyostelium discoideum. 11. Control analysis. J. Biol. Chem. 267: 3106-3114. Amagai, A., and Filosa, M.F. 1984. The possible involvement of cyclic AMP and volatile substance(s) in the development of a macrocyst-forming strain of Dictyostelium mucoroides. Dev. Growth Differ. 26: 583-589. Anjard, C., Pinaud, S., Kay, R.R., and Reymond, C.D. 1992. Overexpression of the Dd PK2 protein kinase causes rapid development and affects the intracellular cAMP pathway of Dictyostelium discoideum. Development (Cambridge, U.K.), 115: 785-790. Armant, D.R., and Rutherford, C.L. 1981. Copurification of alkaline phosphatase and 5'AMP specific nucleotidase in Dictyostelium discoideum. J. Biol. Chem. 256: 12 710-12 718. Armant, D.R., and Rutherford, C.L. 1982. Properties of a 5'AMP specific nucleotidase which accumulates in one cell type during development of Dictyostelium discoideum. Arch. Biochem. Biophys. 216: 485-494. Barisic, K., Mollner, S., Noegel, A.A., Gerisch, G., and Segall, J.E. 1991. cDNA sequence of cyclophilin from Dictyostelium discoideum. Dev. Genet. 12: 50-53. Barklis, E., and Lodish, H.F. 1983. Regulation of Dictyostelium discoideum mRNAs specific for prespore or prestalk cells. Cell, 32: 1139-1 148. Berks, M., and Kay, R.R. 1988. Cyclic AMP is an inhibitor of stalk cell differentiation in Dictyostelium discoideum. Dev. Biol. 126: 108-1 14. Berks, M., and Kay, R.R. 1990. Combinational control of cell differentiation by cAMP and DIF-1 during development of Dictyostelium discoideum. Development (Cambridge, U.K.), 110: 977-984.


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Mutants of Dictyostelium discoideum defective in spore germination.

Protein and RNA synthesis during spore germination in the cellular slime mold Dictyostelium discoideum.

The physiological effects of restrictive environmental conditions on Dictyostelium discoideum spore germination.

The Dictyostelium discoideum spore germination-specific cellulase is organized into functional domains.

Coordinate regulation of the spore coat genes in Dictyostelium discoideum.

Mathematical modeling of differentiation in Dictyostelium discoideum.

Requirement for cell differentiation in Dictyostelium discoideum.

Cell differentiation and patterning in Dictyostelium.

Differentiation without mitosis in Dictyostelium discoideum.

Cellulases released during the germination of Dictyostelium discoideum spores.

Rates of degradation and synthesis of glycosidases de novo during growth and differentiation of Dictyostelium discoideum.

Effect of cerulenin on the growth and differentiation of Dictyostelium discoideum.

Chemotactic signals induce cell differentiation in Dictyostelium discoideum.

Germination and mitochondrial damage in spores of Dictyostelium discoideum following supraoptimal heating.

Products of endogenous N-acetylglucosaminyltransferase activity of Dictyostelium discoideum during growth and early development.

A chymotrypsin-sensitive step in the development of Dictyostelium discoideum.

The Dictyostelium discoideum RACK1 orthologue has roles in growth and development.

Spore coat is altered in modB glycosylation mutants of Dictyostelium discoideum.

Patterning in Dictyostelium discoideum: the proportions of the three differentiated cell types (spore, stalk, and basal disk) in the fruiting body.

Inhibition of development in Dictyostelium discoideum by sugars.

V4, a gene required for the transition from growth to development in Dictyostelium discoideum.

Synthesis of plasma membrane proteins during development of Dictyostelium discoideum.

Inhibition of 5'-nucleotidase activity after growth of Dictyostelium discoideum.

Fruiting bodies of the social amoeba Dictyostelium discoideum increase spore transport by Drosophila.