應用基因標定突變法研究黏菌基因體功能

行政院國家科學委員會專題研究計畫 成果報告

應用基因標定突變法研究黏菌基因體功能

計畫類別： 個別型計畫

計畫編號： NSC93-2320-B-006-088-

執行期間： 93 年 08 月 01 日至 94 年 07 月 31 日

執行單位： 國立成功大學生物化學暨分子生物學科（所）

計畫主持人： 張文粲

報告類型： 精簡報告

處理方式： 本計畫可公開查詢

中 華 民 國 94 年 10 月 31 日

行政院國家科學委員會補助專題研究計畫 ■ 成 果 報 告

□期中進度報告 應用基因標定突變法研究黏菌基因體功能

計畫類別：■ 個別型計畫 □ 整合型計畫 計畫編號：NSC 93－2320－B－006－088－

執行期間：2004 年 08 月 01 日至 2005 年 07 月 31 日

計畫主持人：張文粲 共同主持人：

計畫參與人員：

成果報告類型(依經費核定清單規定繳交)：■精簡報告 □完整報告

本成果報告包括以下應繳交之附件：

□赴國外出差或研習心得報告一份

□赴大陸地區出差或研習心得報告一份

□出席國際學術會議心得報告及發表之論文各一份

□國際合作研究計畫國外研究報告書一份

處理方式：除產學合作研究計畫、提升產業技術及人才培育研究計畫、

列管計畫及下列情形者外，得立即公開查詢

□涉及專利或其他智慧財產權，□一年□二年後可公開查詢

執行單位：國立成功大學生物化學暨分子生物學科（所）

附件一

Developmental Biology 285, 238-251, 2005.

LrrA, a Novel Leucine Rich Repeat Protein Involved in Cytoskeleton Remodeling, Is Required for Multicellular Morphogenesis in Dictyostelium discoideum

Chia-I Liu

, Tsung-Lin Cheng

, Shu-Zhen Chen

, Ying-Chieh Huang

, and Wen-Tsan Chang

1Department of Biochemistry, and ²Institute of Basic Medical Sciences, National Cheng Kung University Medical College, Tainan 701, Taiwan, ROC

Running title: A novel Dictyostelium morphogenesis gene

*Corresponding author.

Tel: +886-6-2353535 Ext. 5533.

Fax: +886-6-2741694.

E-mail address: [email protected].

ABSTRACT

Cell sorting by differential cell adhesion and movement is a fundamental process in multicellular morphogenesis. We have identified a Dictyostelium discoideum gene encoding a novel protein, LrrA, which composes almost entirely leucine-rich repeats (LRRs) including a putative leucine zipper motif. Transcription of lrrA appeared to be developmentally regulated with robust expression during vegetative growth and early development. lrrA null cells generated by homologous recombination aggregated to form loose mounds, but subsequent morphogenesis was blocked without formation of the apical tip. The cells adhered poorly to a substratum and did not form tight cell-cell agglomerates in suspension;

in addition, they were unable to polarize and exhibit chemotactic movement in the submerged aggregation and Dunn chamber chemotaxis assays. Fluorescence-conjugated phalloidin staining revealed that both vegetative and aggregation competent lrrA

cells contained numerous F-actin enriched microspikes around the periphery of cells.

Quantitative analysis of the fluorescence-stained F-actin showed that lrrA

cells exhibited a dramatically increase in F-actin as compared to the wild-type cells. When developed together with wild-type cells, lrrA

cells were unable to move to the apical tip and sorted preferentially to the rear and lower cup regions. These results indicate that LrrA involves in cytoskeleton remodeling, which is needed for normal chemotactic aggregation and efficient cell sorting during multicellular morphogenesis, particularly in the formation of apical tip.

Key words: Leucine rich repeats (LRRs), Gene knockout, Cell adhesion, Cytoskeleton remodeling, Multicellular morphogenesis, Dictyostelium discoideum

INTRODUCTION

Leucine-rich repeats (LRRs) are short consensus sequence motifs present in many proteins with diverse functions and subcellular localizations. They appear to be generally involved in protein-protein interactions (Kobe and Deisenhofer, 1994; Kobe and Deisenhofer, 1995). LRRs motif-containing proteins not only participate in many biologically important processes, such as signal transduction, cell adhesion, disease resistance and innate immune response (Buchanan and Gay, 1996), but also involve in cell polarization (Bilder and Perrimon, 2000), apoptosis signaling (Inohara et al., 1999; Inohara and Nunez, 2001), mammalian development (Tong et al., 2000a;

Tong et al., 2000b) and cytoskeleton remodeling (Xu et al., 1997; Wu et al., 2000).

LRRs are usually present in tandem with the number of repeats among 1–30 (Buchanan and Gay, 1996). The most common length of a LRR is 24 residues, but repeats containing between 20 and 29 residues have been described (Buchanan and Gay, 1996). There is a distinctive subgroup of LRRs that are 23 amino acids in length instead of 24, and which contain additional consensus residues, especially proline (Fig. 1). These proteins have in common an ability to associate with the Ras family of signalling GTPases (Buchanan and Gay, 1996), such as the middle-repetitive repeat region in Saccharomyces cerevisiae adenylyl cyclase (Cyr1p) mediates interaction of the cyclase with RAS proteins (Field et al., 1990; Suzuki et al., 1990), and the LRR protein SUR-8 in

Caenorhabditis elegans and mammals acts as a scaffold and forms a complex with Ras and Raf

The cellular slime mould Dictyostelium discoideum grows as a unicellular amoeba, but undergoes a relatively simple multicellular development. During early aggregation stage, amoebae move chemotactically towards cAMP, make contact, and join into streams that in turn collect into hemispherical mounds of up to 10⁵ cells (Devreotes, 1989; Firtel, 1995; Chen et al., 1996; Parent and Devreotes, 1996). Within each multicellular assembly amoebae differentiate autonomously and randomly into prespore and prestalk cells (Early et al., 1995; Firtel, 1995;

Firtel, 1996; Shaulsky and Loomis, 1996). The prestalk cells then move differentially to the apex of the mound, where they form a tip that acts as an organizing center (Rubin and Robertson, 1975;

Williams et al., 1989; Clow and McNally, 1999; Kellerman and McNally, 1999; Nicol et al., 1999;

Clow et al., 2000; Firtel and Meili, 2000). This tip elongates and falls over to form a migrating slug, or pseudoplasmodium, within which prestalk and prespore cells organize along an anterior-posterior axis pattern. In response to environmental signals, the slug undergoes culmination, resulting in the formation of a mature fruiting body containing terminally differentiated spores and stalk cells (Gross, 1994; Parent and Devreotes, 1996; Aubry and Firtel, 1999).

During tip formation and subsequent morphogenesis, cell sorting is mediated by differential chemotaxis to cAMP via similar chemotaxis pathways and differential cell adhesion systems (Traynor et al., 1992; Reymond et al., 1995; Firtel and Meili, 2000; Dormann et al., 2000;

Verkerke-van Wijk et al., 2001). Therefore, interfering with cAMP signaling by disrupting a prestalk cell specific cAMP receptor subtype 2 (CAR2) (Saxe III et al., 1993), overproducing the secreted form of cAMP phosphodiesterase (Traynor et al., 1992), or disrupting genes encoding

components involved in cytoskeleton remodeling, such as F-actin cross-linking proteins (Witke et

al., 1992), myosin heavy chain (Traynor et al., 1994), myosin II regulatory light chain (Clow et al., 2000) and limB (Chien et al., 2000), blocks development at the mound stage. Similarly,

lagC (Dynes et al., 1994; Wang et al., 2000; Kibler et al., 2003) and dtfA (Ginger et al., 1998),

To identify novel genes involved in pattern formation of multicellular development in D.

discoideum, we searched genomic and cDNA databases (All Dictyostelium BLAST Search

MATERIALS AND METHODS Cell and molecular biology methods

Cell culture, development and transformation of Dictyostelium discoideum cells, Southern, Northern and Western blotting, and lacZ reporter analysis have all been described previously (Chang et al., 1995; Chang et al., 1996; Huang et al., 2004). Molecular cloning and analysis approaches are standard as described by Sambrook et al. (1989) and Ausubel et al. (1998).

Cellular location of LrrA protein

The complete lrrA coding sequence (encoding amino acids 1-510) and a mutant version with the leucine zipper motif deleted (deleting amino acids 197-321) were created by PCR from the full-length lrrA cDNA with the appropriate primers and were cloned into the pAct15-GFP plasmid. The resulting constructs contained the wild-type or mutated lrrA coding sequence fused in-frame with the DNA sequence for the green fluorescent protein (GFP-C3) (Crameri et al., 1996) under the control of the actin 15 promoter and actin 8 terminator. These constructs were used for transformation of wild-type Ax2 and lrrA^- cells by electroporation, selecting for resistance to G418.

Generation of lrrA null mutants

To effectively target the lrrA locus in the D. discoideum genome, we created two lrrA knockout constructs, BglII-KO and SnaBI-KO (see Supplementary data, Suppl. Fig. 1A) as described previously (De Lozanne, 1987). Constructs were made containing 1.2 kb (BglII-KO) and 1.6 kb (SnaBI-KO) of lrrA genomic DNA with a blasticidin S-resistance cassette (Bsr^r) inserted into the lrrA-coding sequence in the same orientation in each construct, so that its strong actin 8 termination sequence blocked any transcriptional read through. Transformants were then plated on SM-agar with K. aerogenes as a food source and developed. Colonies that displayed

aberrant developmental morphology were picked from the SM-agar plates and recloned. The insertional mutants obtained were analyzed by Southern and Northern blotting to verify disruption of the lrrA gene.

Complementation by transformation

The complete lrrA coding sequence (encoding amino acids 1-510) was generated by PCR from the full-length lrrA cDNA with the appropriate primers and cloned into the pAct15-GST plasmid (Huang et al., 2004). The resulting construct contained the wild-type lrrA cDNA fused in-frame with DNA for glutathione S-transferase (GST) under the control of the actin 15 promoter and actin 8 terminator. The constructs, act15-lrrA-gst and act15-gst, were used for complementation of lrrA null mutant SnaBI-KO using electroporation and selection for resistance to G418.

Cell-substratum adhesion assay

The cell-substratum adhesion assay was adopted from Vogel (1987) and Chen and Katz (2000). Exponentially grown cells were harvested and washed three times with Sorensen buffer, and resuspended to 1×10⁶ cells/ml in the same buffer. Four ml of this suspension was incubated in 50 ml glass cell culture flasks on a gyratory shaker at 120 rpm for 10 minutes at room temperature. The cells were incubated for a further 40 minutes without shaking to allow them to adhere. After that the flasks were agitated gently for 3 minutes at 60 rpm and the supernatants were transferred to a test tube. Non-adherent cells in each supernatant were determined using a hemacytometer.

Cell agglutination assay

The cell agglutination assay was performed as described by Varney et al. (2002).

Exponentially grown cells were harvested and washed three times with KK2 buffer, and resuspended to 3×10⁶ cells/ml in the same buffer. Two ml of the suspension was transferred to a plastic Falcon 2059 15-ml test tube (BD Biosciences, San Jose, CA), and left for 6 hours at 22^oC on a rotary wheel set to 45 rpm. Samples were then removed from the tubes with a Pasteur pipette and overlaid onto a borosilicate cover slip for photography.

F-actin staining by Alexa Fluor 488-conjugated phalloidin

For F-actin staining, vegetative cells from axenic culture or agglutination-competent cells were deposited onto glass cover slips for 10 minutes, fixed in 3.7% formaldehyde solution for 10 minutes, permeabilized in 0.2% Triton X-100 solution for 5 minutes, and stained with Alexa Fluor 488-conjugated phalloidin solution (Molecular Probes, Eugene, OR) for 20 minutes. The amount of F-actin was analyzed by flow cytometry. The flow cytometry measurements were performed at a FACSCalibur^TM flow cytometer (BD Biosciences, San Jose, CA).

Dunn chamber chemotaxis assay

The chemotactic migration assay of D. discoideum cells was assessed by directly

observation and recording of cell behavior in stable concentration gradients of chemoattractant, cAMP or folic acid, using the Dunn chemotaxis chamber (Weber Scientific International Ltd., Teddington, UK) (Zicha et al., 1991; Webb et al., 1996; Allen et al., 1998). Chemoattractants added to the outer well of the apparatus diffuse across the annular bridge to the inner blind well of the chamber and form a gradient. In these experiments, the outer well of the chamber was filled with developing buffer containing 10 μM of cAMP or 100 μM of folic acid and the concentric inner well with only developing buffer. Coverslips with cells were inverted onto the Dunn chamber and cell migration was observed through the annular bridge between the concentric inner and outer wells.

Aggregation in submerged monolayers

Exponentially grown cells were harvested and washed three times with aggregation buffer (pH6.2). Four ml of the cell suspension was plated in 5-cm tissue culture dishes (Nalge Nunc International, Rochester, NY) to a final density of 2 x 10⁵ cells/cm². Cells were incubated at 22^oC in a humid box.

Accession number

The DDBJ/EMBL/GenBank accession number for the lrrA gene is AF200466 and for the

lrrA promoter is AF388909.

RESULTS

Identification of the leucine rich repeat protein LrrA

By searching the expressed sequence tag (EST) database of Dicty_cDB (http://www.csm.biol.tsukuba.ac.jp/cDNA/database.html) with the Saccharomyces cerevisiae adenylyl cyclase (Cyr1p) amino acid sequence (Kataoka et al., 1985), we identified a cDNA clone FCL-AC05, which has homology to the Cyr1p LRR motif. The cDNA contains one long open reading frame of 1530 bp that encodes a predicted protein of 510 amino acids with a calculated molecular mass of 57 kDa. It contains 19 tandem repeats of the LRR motif (amino acids 31-470), including a putative leucine zipper motif (amino acids 198-219) (Fig. 1). The full-length cDNA includes 284 bp of 5’-untranslated sequence and 71 bp of 3’-untranslated sequence. Sequencing the genomic DNA revealed a single intron interrupting the coding sequence of lrrA (data not shown).

Alignment of the high scoring segments obtained from a BLAST search of the non-redundant NCBI database (http://www.ncbi.nlm.nih.gov/BLAST/) using the deduced amino acid sequence of the isolated cDNA coding region revealed that the putative product displayed weak similarity to SUR-8, which is a positive regulator of Ras signaling that enhances MAP kinase activation and forms a complex with Ras and Raf (Sieburth et al., 1998; Li et al., 2000).

Since the predicted protein was composed almost entirely of LRR motifs, the gene was designated as lrrA. Searching the non-redundant NCBI database, as well as the D. discoideum genomic and cDNA databases (All Dictyostelium BLAST Search; http://dicty.sdsc.edu/) with the

lrrA nucleotide sequence failed to reveal any other related genomic matches, indicating that lrrA

Analysis of the lrrA gene

The temporal expression pattern of lrrA during development was determined by extracting RNA at various times of development and hybridizing with full-length lrrA cDNA. lrrA encodes a single transcript of 1.6 kb that is substantially expressed during vegetative growth. We observed that lrrA expression increased upon starvation, peaked at 6-9 hours of the aggregation stage, and declined abruptly from 16 hours of development (Fig. 2A).

To identify the spatial pattern of lrrA expression during development, we isolated and sequenced the lrrA promoter region including the partial coding sequence of the upstream clcA gene (AF414428), and made a reporter construct with the Escherichia coli lacZ gene under its control. A strain harboring this construct was developed on KK2 buffered-agar, fixed, and stained with X-gal. All cells stained strongly during early development. However, the pattern of expression in first fingers and early culminates became progressively decreased in the staining cells. In fruiting bodies, both the spore and stalk cells exhibited weak staining with X-gal (data not shown).

Amino acid sequence analysis indicated that LrrA did not contain any putative targeting sequences or transmembrane domains, suggesting that LrrA is a cytoplasmic protein. To localize LrrA within the cell, constructs driven by the actin 15 promoter were made with either full-length

lrrA or using a mutant with deleted leucine zipper motif fused in-frame with the highly sensitive

Δ -gfp transformants was uniform

Disruption of the lrrA gene

To investigate the function of LrrA protein during D. discoideum growth and development, two independent lrrA knockout strains were generated specifically (see Materials and Methods).

Disruption of the lrrA gene was confirmed by Southern blot (see Supplementary data, Suppl. Fig.

1B) and Northern blot analyses using an upstream lrrA DNA fragment as a probe. No transcripts were found in both the BglII-KO and SnaBI-KO mutants (data not shown). Efficiency of the homologous recombination with both the BglII-KO and SnaBI-KO targeting constructs was approximately 30 percent. All of the isolated mutants displayed identical phenotypes. The lrrA^- cells grew normally in association with K. aerogenes on SM-agar plate and in liquid axenic medium (data not shown). During development, the cells aggregated and formed mounds with the similar timing as that of the wild-type cells, but the mounds failed to form apical tips both on bacterial lawns (data not shown) and on KK2 agar (Fig. 3). However, the loose mounds formed by

lrrA

To confirm that the phenotype was caused by disruption of lrrA, we transformed lrrA^- cells with an lrrA expression construct fused in-frame with the glutathione S-transferase (GST), under

the control of the actin 15 promoter. The results revealed that only the expressed LrrA-GST fusion construct restored the ability of lrrA null cells to develop normally and produce mature fruiting bodies of essentially wild-type appearance (Fig. 3), consistent with a fully functional LrrA-GST fusion. Expression of the LrrA-GST fusion protein was verified by Western blot analysis using anti-GST antibody (see Supplementary data, Suppl. Fig. 2).

Effects on cell adhesion and cytoskeletal F-actin

The earliest detectable phenotypic trait in the mutant cells was their weak adhesion to the surfaces of plastic petri dishes, as revealed by their detachment upon gentle pipetting or even suspension in the axenic medium without any attachment to the surface (data not shown). To investigate this phenotypic trait directly, we measured their adhesion to the substratum and to other cells. The cell substratum adhesion assay quantifies the ability of amoebae to attach to a glass surface (Vogel, 1987; Chen and Katz, 2000). Wild-type and lrrA^- cells were allowed to attach to the bottom of glass cell culture flasks prior to gentle and brief agitation. The non-adherent cells were then counted. Wild-type cells adhered more than did the lrrA^- cells (Fig.

4A).

Microscopic examination of vegetative cells growing in plastic petri dishes for 24 hours revealed that wild-type cells displayed an elongated and polarized appearance, while lrrA^- cells exhibited a rounded and nonpolar morphology (Fig. 4B). To examine this difference directly, we stained the actin cytoskeleton in wild-type and lrrA^- cells from axenic culture with Alexa Fluor 488-conjugated phalloidin. In wild-type cells, the majority of F-actin was recruited into the macropinocytic crowns whereas in lrrA^- cells the F-actin was enriched in cell surface projections as microspikes (Fig. 4C). To determine more precisely the levels of F-actin, the amount of fluorescence-stained F-actin was analyzed using a fluorescence-activated cell sorter. The levels of F-actin in the lrrA^- cells were markedly elevated with respect to the wild-type cells (Fig. 4D).

These results indicated that vegetative lrrA^- amoebae were defective in organization of the actin cytoskeleton.

When D. discoideum cells were washed free of growth medium, resuspended in non-nutrient buffer, and shaken in suspension, large cell agglomerates formed (Varney et al., 2002). The agglomerates formed by lrrA^- cells starved in KK2 buffer for 6 hours were significantly smaller and looser than those formed by wild-type cells (Fig. 4E). Microscopic examination of the lrrA^- cells showed that they were rounded and had many microspikes around their periphery, and appeared not to be polarized, suggesting of an aberrant organization of the actin cytoskeleton. To examine this idea directly, we stained the actin cytoskeleton in wild-type and lrrA^- cells with Alexa Fluor 488-conjugated phalloidin. F-actin was observed to be highly localized to the anterior leading edge and to a lesser degree in the posterior region of polarized agglutination-competent wild-type cells (Fig. 4F). In contrast, lrrA^- cells possessed many F-actin-enriched microspikes around the periphery of the cells (Fig. 4F). To determine more precisely the levels of F-actin, the amount of fluorescence-stained F-actin was analyzed using a fluorescence-activated cell sorter. The levels of F-actin in the lrrA^- cells were dramatically elevated with respect to the wild-type cells (Fig. 4G), indicating that aggregation competent lrrA^-

cells were markedly defective in the regulation of F-actin organization.

Effects on chemotactic response

It is known that cAMP is especially active as a chemotactic agent during the aggregation stage and shows relatively little activity during the vegetative growth. To analyze the effect of

lrrA mutation on chemotactic migration at the aggregation stage, we examined the chemotactic

In D. discoideum cells, reorganization of the actin cytoskeleton is essential for cellular motility in response to chemotactic agents. During vegetative growth, amoebae are chemotactic to folic acid, which is specifically secreted by bacteria and is used in food seeking (Pan et al., 1972; Pan et al., 1975). To assess the effect of lrrA mutation on cellular motility at the vegetative phase, we examined the chemotactic responsiveness of wild-type and lrrA^- cells from axenic culture and starved for 6 hours in shaken suspension to gradients of folic acid. Wild-type vegetative amoebae exhibited strongly chemotactic responses to gradients of folic acid at concentration 100 μM whereas lrrA^- cells were totally unresponsive to gradients of folic acid (Fig.

6A). Similar results were also observed when chemotaxis assays were conducted on cells that had been starved for 6 hours in shaken suspension (Fig. 6B). The results were consistent with the chemotactic properties to gradients of cAMP observed during aggregation stage.

Effects on multicellular morphogenesis

To analyze in detail the effect of lrrA null mutation on multicellular development, we characterized several aspects of the development of lrrA^- cells. Firstly, we examined aggregation of the cells at different cell densities on KK2 agar. The lrrA^- cells aggregated efficiently even at very low densities whereas wild-type cells were delayed in their aggregation, and produced larger loose mounds. The mutant cells never formed streams at any cell density (Fig. 7A). In addition, in the submerged monolayer aggregation assay, lrrA^- cells did not polarize properly, and exhibited severe impairment in chemotaxis (Fig. 7B). In contrast, the wild-type cells formed streams of migrating cells joined head to tail as they moved in towards the aggregation center, which was emitting pulses of the chemoattractant cAMP (Fig. 7B).

These results suggested that lrrA^- cells were impaired in cAMP signaling during early aggregation. To examine this possibility, wild-type and lrrA^- cells were pulsed with 30 nM cAMP for 6 hours to mimic the normal oscillatory pulses of cAMP that occur during early aggregation, and then were plated on non-nutrient agars. Unlike the response seen in wild-type cells, the lrrA^- cells formed aggregates, but still arrested at the mound stage without the formation of apical tip (data not shown).

Next, we examined the developmental phenotype of Ax2 and lrrA^- cells in mixtures with various ratios. The chimeric aggregates were able to form apical tips and mature fruiting bodies but the size of the fruiting bodies decreased dramatically when half of the cells in the mixtures were mutant cells (Fig. 8A). In contrast, formation of the tipless aggregates increased significantly in the numbers and sizes with respect to the ratios of mutant cells increases, indicating that disruption of the lrrA gene results in sorting out of mutant from wild-type cells during aggregation. These results suggested that lrrA^- cells either failed to co-aggregate with wild-type cells or, if co-aggregating, were unable to be rescued by wild-type cells.

To examine the ability of lrrA^- cells to sort and form specific structures in these chimeric organisms, we tagged mutant cells with the constitutively expressed reporter construct act15-gfp.

Most of the gfp⁺ mutant cells were found at the periphery of the mound. Few were found inside the mound, and no GFP-expressed cells were observed in the apical tip domain (Fig. 8B), indicating that most of the lrrA^- cells were excluded from cell sorting in the mound. During the post-aggregative stage, the mutant cells sorted preferentially to the rear of the slug and lower cup region of spore head, and tended to be lost from the rear of standing and migrating slugs (Fig. 8B).

We confirmed these results using the lacZ reporter tagged lrrA^- cells. The spatial pattern of mutant cells in chimeric organisms with wild-type cells was similar to that observed using GFP labeled lrrA^- cells (see Supplementary data, Suppl. Fig. 3). In addition, when mixed with an excess of wild-type cells, very few of the lrrA^- detergent- and heat-resistant spores were formed when spore viability was assayed (<3%; data not shown).

Effects on developmental gene expression

To investigate why lrrA^- cells could aggregate to form loose mounds but failed to form streams, we examined the expression of the aggregation-stage genes aca (aggregation adenylyl cyclase; Pitt et al., 1992; Pitt et al., 1993; Kriebel et al., 2003), car1 (cAMP receptor 1; Klein et

al., 1988; Sun and Devreotes, 1991), pde (cAMP phosphodiesterase; Lacombe et al., 1986; Faure et al., 1988; Faure et al., 1990; Hall et al., 1993), cadA (calcium-dependent cell adhesion

Desbarats et al., 1992). Northern blots of RNAs extracted from the wild-type and lrrA^- cells at various times during development showed that the time of appearance of transcripts of the early genes aca, car1 and pde was delayed by roughly 6 hours in the mutant and their expression level was markedly reduced (Fig. 9A), indicating that lrrA^- cells were impaired in cAMP signaling during early aggregation.

The initial expression of cadA was similar in wild-type and mutant cells, but expression thereafter differed, with maximal levels being attained at nine hours in wild-type cells and at 12 hours in mutant cells. In wild-type cells, cadA expression was down regulated after 9 hours, whereas expression remained elevated in lrrA^- cells (Fig. 9A), possibly because of the arrest of the lrrA^- cells at the loose mound stage. In lrrA^- cells, however, the level of csA expression was slightly reduced compared with that of wild-type cells, and the kinetics of induction was delayed by roughly 6 hours. In wild-type cells, csA expression was down regulated after 6 hours but remained elevated in mutant cells, likely due to the arrest of the mutant cells at the mound stage

(Fig. 9A). These results indicated that the defect of streaming in lrrA^- cells could not be ascribed to the failure to express cell adhesion molecules, such as DdCAD-1 and csA.

To examine post-aggregative cell-type differentiation, we analyzed the expression of the cell-specific prestalk and prespore genes, ecmA and psA (Ceccarelli et al., 1987; Jermyn et al., 1987; Early et al., 1988a; Early et al., 1988b). In lrrA^- cells, ecmA and psA expression was delayed by roughly 4 hours and their level of expression was greatly decreased (Fig. 9B), again pointing to very limited post-aggregative development.

DISCUSSION

In this study, we identified and analyzed the biological function of novel protein, LrrA, in D.

discoideum. LrrA consists almost entirely of a series of nineteen 23-amino acid LRRs that

D. discoideum expresses a large number of Ras proteins with a wide range of functions

Lim et al., 2002). For instance, the small GTPase RasG protein may control the actin cytoskeleton and play a role in adhesion (Tuxworth et al., 1997). Cells expressing constitutively active RasG[G12T] show increased cell-substrate adhesion (Chen and Katz, 2000). In addition, the developmental expression of constitutively active RasD[G12T] or RasG[G12T] results in multi-tipped aggregates, which do not undergo further morphogenesis (Reymond et al., 1986;

Jaffer et al., 2001). Furthermore, recent studies have identified a novel RasG-interacting protein (RIP3) whose function is required for both chemotaxis and cAMP signaling (Lee et al., 1999). In view of the well-established relation between LRRs and Ras proteins, we tested whether LrrA can function as a scaffold protein in Ras signaling pathways. Using the yeast two-hybrid system, we examined the possible interaction between LrrA and RasD[G12T] or RasG[G12T]. We found no indications of such an interaction (S.-Z. Chen, unpublished). While this suggests that LrrA functions independently of interaction with Ras proteins, we cannot exclude the possibility that it interacts with some other Ras protein.

As a novel protein, LrrA clearly plays a decisive role in multicellular morphogenesis, especially in the formation of apical tip. The defect in tip formation in the lrrA^- cells is significant as this structure serves as the signaling center in the mound, and is essential for differentiation and morphogenesis to proceed. Tip formation is a process of differential cell movement in which prestalk cells sort and accumulate to form the anterior domain of the multicellular structure (Durston and Vork, 1979; Williams et al., 1989; Jermyn et al., 1996; Vasiev and Weijer, 1999).

How the tip is formed is still a mystery. Only a handful of genes are defective in tip formation,

such as null mutations in cAR2 (Saxe III et al., 1993), limB (Chien et al., 2000), myosin II (De Lozanne and Spudich, 1987; Knecht and Loomis, 1987; Clow et al., 2000), and dtfA (Ginger et

al., 1998). The lrrA

However, there are differences in the mound morphology and the timing of aggregation; the limB^- mounds are of a normal size but there is a delay in their formation. Thus, LrrA has a distinct role in mound and tip formation.

Formation of a multicellular structure in D. discoideum requires cell-cell adhesion. During early development, there are EDTA-sensitive and EDTA-resistant cell-cell adhesions. The former is mediated by DdCAD-1 (Brar and Siu, 1993; Wong et al., 1996) and the latter by csA (Siu et al., 1985). The earliest detectable phenotypic trait in lrrA^- cells is a defect in cell-substratum adhesion.

Presently, cell adhesion analyses revealed that lrrA^- cells are adhesion defective. Northern blot analysis of cell adhesion molecules showed that the expression level of cadA and csA in lrrA^- cells is roughly similar to that of the wild-type, but the expression kinetics of both genes is markedly delayed. These results indicate that lrrA^- cells are capable of aggregating to form larger loose mounds, but are incapable of aggregating tightly to form an apical tip to organize the subsequent morphogenesis, rather than to the inability to express cell adhesion molecules such as DdCAD-1 and csA. In addition, microscopic examination of single, aggregation stage cells demonstrated that lrrA^- cells do not become polarized and have defects in actin organization by extending more F-actin enriched microspikes. This abnormal cell shape may reduce the ability to make physical contacts and again contribute to its developmental phenotype. These results indicate that LrrA plays an important role in both adhesion and cytoskeleton remodeling during early development in D. discoideum.

In conclusion, our results clearly indicate that LrrA is not needed for cell growth but is required for effective adhesion to the substratum and to other cells, which is absolutely essential for the formation of apical tip and subsequent developmental morphogenesis. This is entirely consistent with the primary function of the LRR motifs to provide a versatile structural framework for the protein-protein interactions (Kajava, 1998; Kobe and Kajava, 2001). Our preliminary analyses from a comparative protein structural modeling analysis (SWISS-MODEL,

the GlaxoSmithKline, Geneva, Switzerland;

http://www.expasy.ch/swissmod/SWISS-MODEL.html) and a fully automated protein structure meta prediction system (3D-Jury, http://BioInfo.PL/Meta; Ginalski et al., 2003; Ginalski and Rychlewski, 2003) indicate that the LRR motif in the LrrA protein resembles the structure of LRRs in Listeria monocytogenes internalin (InlA), which mediates bacterial adhesion and invasion of epithelial cells in the human intestine through specific interaction with its host cell receptor E-cadherin (Schubert et al., 2001; Schubert et al., 2002), suggesting that LrrA might function as a scaffold in protein-protein interaction (T.-L. Cheng, unpublished). To elucidate the biochemical function of the LrrA protein in the cytoskeleton remodeling during multicellular morphogenesis, it will be useful to identify protein(s) that interact with it.

ACKNOWLEDGMENTS

We thank the Dictyostelium cDNA project in Japan for kindly providing the original cDNA clone FCL-AC05. This work was supported by grants from the National Science Council of Taiwan, ROC (to Wen-Tsan Chang).

REFERENCES

Allen, W. E., Zicha, D., Ridley, A. J., and Jones, G. E. (1998). A role for Cdc42 in macrophage chemotaxis. J. Cell Biol. 141, 1147-1157.

Aubry, L., and Firtel, R. A. (1999). Integration of signaling networks that regulate Dictyostelium differentiation. Annu. Rev. Cell Dev. Biol. 15, 469-517.

Ausubel, F. M., Brent, R., Kingston, R. E., Moore, D. D., Seidman, J. G., Smith, J. A., and Struhl, K. (1998). Current Protocols in Molecular Biology. John Wiley and Sons, New York.

Bilder, D., and Perrimon, N. (2000). Localization of apical epithelial determinants by the basolateral PDZ protein Scribble. Nature 403, 676-680.

Brar, S. K., and Siu, C. H. (1993). Characterization of the cell adhesion molecule gp24 in

Dictyostelium discoideum. Mediation of cell-cell adhesion via a Ca

J. Biol. Chem. 268, 24902-24909.

Buchanan, S. G. ST. C., and Gay, N. J. (1996). Structural and functional diversity in the leucine-rich repeat family of proteins. Prog. Biophys. Molec. Biol. 65, 1-44.

Ceccarelli, A., McRobbie, S. J., Jermyn, K. A., Duffy, K., Early, A., and Williams, J. G. (1987).

Structural and functional characterization of a Dictyostelium gene encoding a DIF inducible, prestalk-enriched mRNA sequence. Nucleic Acids Res. 15, 7463-7476.

Chang, W.-T., Gross, J. D., and Newell, P. C. (1995). Trapping developmental promoters in

Dictyostelium. Plasmid 34, 175-183.

Chang, W.-T., Newell, P. C., and Gross, J. D. (1996). Identification of the cell fate gene stalky in

Dictyostelium. Cell 87, 471-481.

Chen, C. F., and Katz, E. R. (2000). Mediation of cell-substratum adhesion by RasG in

Dictyostelium. J. Cell Biochem. 79, 139-149.

Chen, M. Y., Insall, R. H., and Devreotes, P. N. (1996). Signaling through chemoattractant receptors in Dictyostelium. Trends Genet. 12, 52-57.

Chien, S., Chung, C. Y., Sukumaran, S., Osborne, N., Lee, S., Ellsworth, C., McNally, J. G., and Firtel, R. A. (2000). The Dictyostelium LIM domain-containing protein LIM2 is essential for proper chemotaxis and morphogenesis. Mol. Biol. Cell 11, 1275-1291.

Chubb, J. R., and Insall, R. H. (2001). Dictyostelium: an ideal organism for genetic dissection of Ras signalling networks. Biochim. Biophys. Acta. 1525, 262-271.

Clow, P. A., Chen, T., Chisholm, R. L., and McNally, J. G. (2000). Three-dimensional in vivo analysis of Dictyostelium mounds reveals directional sorting of prestalk cells and defines a role for the myosin II regulatory light chain in prestalk cell sorting and tip protrusion.

Development 127, 2715-2728.

Clow, P. A., and McNally, J. G. (1999). In vivo observations of myosin II dynamics support a role in rear retraction. Mol. Biol. Cell 10, 1309-1323.

Crameri, A., Whitehorn, E., Tate, E., and Stemmer, W. P. C. (1996). Improved green fluorescent protein by molecular evolution using DNA shuffling. Nature Biotechnol. 14, 315-319.

De Lozanne, A. (1987). Homologous recombination in Dictyostelium as a tool for the study of developmental genes. Methods Cell Biol. 28, 489-495.

De Lozanne, A., and Spudich, J. A. (1987). Disruption of the Dictyostelium myosin heavy chain

gene by homologous recombination. Science 236, 1086-1091.

Desbarats, L., Lam, T. Y., Wong, L. M., and Siu, C. H. (1992). Identification of a unique cAMP-response element in the gene encoding the cell adhesion molecule gp80 in

Dictyostelium discoideum. J. Biol. Chem. 267, 19655-19664.

Devreotes, P. N. (1989). Dictyostelium discoideum: a model system for cell-cell interactions in development. Science 245, 1054-1058.

Dormann, D., Vasiev, B., and Weijer, C. J. (2000). The control of chemotactic cell movement during Dictyostelium morphogenesis. Philos. Trans. R. Soc. Lond. B. Biol. Sci. 355, 983-991.

Durston, A. J., and Vork, F. (1979). A cinematographical study of the development of vitally stained Dictyostelium discoideum. J. Cell Sci. 36, 261-279.

Dynes, J. L., Clark, A. M., Shaulsky, G., Kuspa, A., Loomis, W. F., and Firtel, R. A. (1994). LagC is required for cell-cell interactions that are essential for cell-type differentiation in

Dictyostelium. Genes Dev. 8, 948-958.

Early, A. E., Abe, T., and Williams, J. G. (1995). Evidence for positional differentiation of prestalk cells and for a morphogenetic gradient in Dictyostelium. Cell 83, 91-99.

Early, A. E., McRobbie, S. J., Duffy, K. T., Jermyn, K. A., Tilly, R., Ceccarelli, A., and Williams, J. G. (1988a). Structural and functional characterization of genes encoding Dictyostelium prestalk and prespore cell-specific proteins. Dev. Genet. 9, 383-402.

Early, A. E., Williams, J. G., Meyer, H. E., Por, S. B., Smith, E., Williams, K. L., and Gooley, A.

A. (1988b). Structural characterization of Dictyostelium discoideum prespore-specific gene D19 and of its product, cell surface glycoprotein PsA. Mol. Cell Biol. 8, 3458-3466.

Faure, M., Franke, J., Hall, A. L., Podgorski, G. J., and Kessin, R. H. (1990). The cyclic nucleotide phosphodiesterase gene of Dictyostelium discoideum contains three promoters specific for growth, aggregation, and late development. Mol. Cell Biol. 10, 1921-1930.

Faure, M., Podgorski, G. J., Franke, J., and Kessin, R. H. (1988). Disruption of Dictyostelium

discoideum morphogenesis by overproduction of cAMP phosphodiesterase. Proc. Natl. Acad.

Sci. USA 85, 8076-8080.

Field, J., Xu, H.-P., Michaeli, T., Ballester, R., Sass, P., Wigler, M., and Colicelli, J. (1990).

Mutations of the adenylyl cyclase gene that block RAS function in Saccharomyces cerevisiae.

Science 247, 464-467.

Firtel, R. A. (1995). Integration of signaling information in controlling cell-fate decisions in

Dictyostelium. Genes Dev. 9, 1427-1444.

Firtel, R. A. (1996). Interacting signaling pathways controlling multicellular development in

Dictyostelium. Curr. Opin. Genet. Dev. 6, 545-554.

Firtel, R. A., and Meili, R. (2000). Dictyostelium: a model for regulated cell movement during morphogenesis. Curr. Opin. Genet. Dev. 10, 421-427.

Ginalski, K., Elofsson, A., Fischer, D., and Rychlewski, L. (2003). 3D-Jury: a simple approach to improve protein structure predictions. Bioinformatics 19, 1015-1018.

Ginalski, K., and Rychlewski, L. (2003). Detection of reliable and unexpected protein fold predictions using 3D-Jury. Nucleic Acids Res. 31, 3291-3292.

Ginger, R. S., Drury, L., Baader, C., Zhukovskaya, N. V., and Williams, J. G. (1998). A novel

Dictyostelium cell surface protein important for both cell adhesion and cell sorting.

Development 125, 3343-3352.

Gross, J. D. (1994). Developmental decisions in Dictyostelium discoideum. Microbiol. Rev. 58, 330-351.

Hall, A. L., Franke, J., Faure, M., and Kessin, R. H. (1993). The role of the cyclic nucleotide phosphodiesterase of Dictyostelium discoideum during growth, aggregation, and morphogenesis: overexpression and localization studies with the separate promoters of the pde. Dev. Biol. 157, 73-84.

Huang, Y.-C., Chen Y.-H., Lo, S.-R., Liu, C.-I, Wang, C.-W., and Chang, W.-T. (2004). Disruption of the peroxisomal citrate synthase CshA affects cell growth and multicellular development in

Dictyostelium discoideum. Mol. Microbiol. 53, 81-91.

Inohara, N., Koseki, T., del Peso, L., Hu, Y., Yee, C., Chen, S., Carrio, R., Merino, J., Liu, D., Ni, J., and Nunez, G. (1999). Nod1, an Apaf-1-like activator of caspase-9 and nuclear factor-kappaB. J. Biol. Chem. 274, 14560-14567.

Inohara, N., and Nunez, G. (2001). The NOD: a signaling module that regulates apoptosis and host defense against pathogens. Oncogene 20, 6473-6481.

Jaffer, Z. M., Khosla, M., Spiegelman, G. B., and Weeks, G. (2001). Expression of activated Ras during Dictyostelium development alters cell localization and changes cell fate. Development

128, 907-916.

Jermyn, K. A., Berks, M., Kay, R. R., and Williams, J. G. (1987). Two distinct classes of prestalk-enriched mRNA sequences in Dictyostelium discoideum. Development 100, 745-755.

Jermyn, K., Traynor, D., and Williams, J. (1996). The initiation of basal disc formation in

Dictyostelium discoideum is an early event in culmination. Development 122, 753-760.

Kajava, A. V. (1998). Structural diversity of leucine-rich repeat proteins. J. Mol. Biol. 277, 519-527.

Kataoka, T., Broek, D., and Wigler, M. (1985). DNA sequence and characterization of the S.

cerevisiae gene encoding adenylate cyclase. Cell 43, 493-505.

Kellerman, K. A., and McNally, J. G. (1999). Mound-cell movement and morphogenesis in

Dictyostelium. Dev. Biol. 208, 416-429.

Kibler, K., Svetz, J., Nguyen, T. L., Shaw, C., and Shaulsky, G. (2003). A cell-adhesion pathway regulates intercellular communication during Dictyostelium development. Dev. Biol. 264, 506-521.

Klein, P. S., Sun, T. J., Saxe, C. L. 3^rd., Kimmel, A. R., Johnson, R. L., and Devreotes, P. N.

(1988). A chemoattractant receptor controls development in Dictyostelium discoideum.

Science 241, 1467-1472.

Knecht, D. A., and Loomis, W. F. (1987). Antisense RNA inactivation of myosin heavy chain gene expression in Dictyostelium discoideum. Science 236, 1081-1086.

Kobe, B., and Deisenhofer, J. (1994). The leucine-rich repeat: a versatile binding motif. Trends

Biochem. Sci. 19, 415-421.

Kobe, B., and Deisenhofer, J. (1995). Proteins with leucine-rich repeats. Curr. Opin. Struct. Biol.

5, 409-416.

Kobe, B., and Kajava, A. V. (2001). The leucine-rich repeat as a protein recognition motif. Curr.

Opin. Struct. Biol. 11, 725-732.

Kriebel, P. W., Barr, V. A., and Parent, C. A. (2003). Adenylyl cyclase localization regulates streaming during chemotaxis. Cell 112, 549-560.

Lacombe, M. L., Podgorski, G. J., Franke, J., and Kessin, R. H. (1986). Molecular cloning and developmental expression of the cyclic nucleotide phosphodiesterase gene of Dictyostelium

discoideum. J. Biol. Chem. 261, 16811-16817.

Lee, S., Parent, C. A., Insall, R., and Firtel, R. A. (1999). A novel Ras-interacting protein required for chemotaxis and cyclic adenosine monophosphate signal relay in Dictyostelium. Mol. Biol.

Cell 10, 2829-2845.

Li, W., Han, M., and Guan, K.-L. (2000). The leucine-rich repeat protein SUR-8 enhances MAP kinase activation and forms a complex with Ras and Raf. Genes Dev. 14, 895-900.

Lim, C. J., Spiegelman, G. B., and Weeks, G. (2002). Cytoskeletal regulation by Dictyostelium Ras subfamily proteins. J. Muscle Res. Cell Motil. 23, 729-736.

Nicol, A., Rappel, W., Levine, H., and Loomis, W. F. (1999). Cell-sorting in aggregates of

Dictyostelium discoideum. J. Cell Sci. 112, 3923-3929.

Noegel, A., Gerisch, G., Stadler, J., and Westphal, M. (1986). Complete sequence and transcript regulation of a cell adhesion protein from aggregating Dictyostelium cells. EMBO J. 5, 1473-1476.

Pan, P., Hall, E. M., and Bonner, J. T. (1972). Folic acid as second chemotactic substance in the cellular slime moulds. Nat. New Biol. 237, 181-182.

Pan, P., Hall, E. M., and Bonner, J. T. (1975). Determination of the active portion of the folic acid molecule in cellular slime mold chemotaxis. J. Bacteriol. 122, 185-191.

Parent, C. A., and Devreotes, P. N. (1996). Molecular genetics of signal transduction in

Dictyostelium. Annu. Rev. Biochem. 65, 411-40.

Pitt, G. S., Brandt, R., Lin, K. C., Devreotes, P. N., and Schaap, P. (1993). Extracellular cAMP is sufficient to restore developmental gene expression and morphogenesis in Dictyostelium cells lacking the aggregation adenylyl cyclase (ACA). Genes Dev. 7, 2172-2180.

Pitt, G. S., Milona, N., Borleis, J., Lin, K. C., Reed, R. R., and Devreotes, P. N. (1992).

Structurally distinct and stage-specific adenylyl cyclase genes play different roles in

Dictyostelium development. Cell 69, 305-315.

Reymond, C. D., Gomer, R. H., Nellen, W., Theibert, A., Devreotes, P., and Firtel, R. A. (1986).

Phenotypic changes induced by a mutated ras gene during the development of Dictyostelium transformants. Nature 323, 340-343.

Reymond, C. D., Schaap, P., Veron, M., and Williams, J. G. (1995). Dual role of cAMP during

Dictyostelium development. Experientia 51, 1166-1174.

Rubin, J., and Robertson, A. (1975). The tip of the Dictyostelium discoideum pseudoplasmodium as an organizer. J. Embryol. Exp. Morphol. 33, 227-241.

Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989). Molecular Cloning: A Laboratory Manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York.

Saxe, C. L. III, Ginsburg, G. T., Louis, J. M., Johnson, R., Devreotes, P. N., and Kimmel, A. R.

(1993). CAR2, a prestalk cAMP receptor required for normal tip formation and late development of Dictyostelium discoideum. Genes Dev. 7, 262-272.

Schubert, W. D., Gobel, G., Diepholz, M., Darji, A., Kloer, D., Hain, T., Chakraborty, T., Wehland, J., Domann, E., and Heinz, D. W. (2001). Internalins from the human pathogen Listeria

monocytogenes combine three distinct folds into a contiguous internalin domain. J. Mol. Biol.

312, 783-794.

Schubert, W. D., Urbanke, C., Ziehm, T., Beier, V., Machner, M. P., Domann, E., Wehland, J., Chakraborty, T., and Heinz, D. W. (2002). Structure of internalin, a major invasion protein of

Listeria monocytogenes, in complex with its human receptor E-cadherin. Cell 111, 825-836.

Shaulsky, G. and Loomis, W. F. (1996). Initial cell type divergence in Dictyostelium is independent of DIF-1. Dev. Biol. 174, 214-220.

Sieburth, D. S., Sun, Q., and Han, M. (1998). SUR-8, a conserved Ras-binding protein with leucine-rich repeats, positively regulates Ras-mediated signaling in C. elegans. Cell 94, 119-130.

Siu, C. H., Lam, T. Y., and Choi, A. H. (1985). Inhibition of cell-cell binding at the aggregation stage of Dictyostelium discoideum development by monoclonal antibodies directed against an 80,000-dalton surface glycoprotein. J. Biol. Chem. 260, 16030-16036.

Stege, J. T., Shaulsky, G., and Loomis, W. F. (1997). Sorting of the initial cell types in

Dictyostelium is dependent on the tipA gene. Dev. Biol. 185, 34-41.

Sun, T. J., and Devreotes, P. N. (1991). Gene targeting of the aggregation stage cAMP receptor cAR1 in Dictyostelium. Genes Dev. 5, 572-582.

Suzuki, N., Choe, H.-R., Nishida, Y., Yamawaki-Kataoka, Y., Ohnishi, S., Tamaoki, T., and Kataoka, T. (1990). Leucine-rich repeats and carboxyl terminus are required for interaction of yeast adenylate cyclase with RAS proteins. Proc. Natl. Acad. Sci. USA 87, 8711-8715.

Tong, Z. B., Gold, L., Pfeifer, K. E., Dorward, H., Lee, E., Bondy, C. A., Dean, J., and Nelson, L.

M. (2000a). Mater, a maternal effect gene required for early embryonic development in mice.

Nat. Genet. 26, 267-268.

Tong, Z. B., Nelson, L. M., and Dean, J. (2000b). Mater encodes a maternal protein in mice with a leucine-rich repeat domain homologous to porcine ribonuclease inhibitor. Mamm. Genome

11, 281-287.

Traynor, D., Kessin, R. H., and Williams, J. G. (1992). Chemotactic sorting to cAMP in the multicellular stages of Dictyostelium development. Proc. Natl. Acad. Sci. USA 89, 8303-8307.

Traynor, D., Tasaka, M., Takeuchi, I., and Williams, J. G. (1994). Aberrant pattern formation in myosin heavy chain mutants of Dictyostelium. Development 120, 591-601.

Tuxworth, R. I., Cheetham, J. L., Machesky, L. M., Spiegelmann, G. B., Weeks, G., and Insall, R.

H. (1997). Dictyostelium RasG is required for normal motility and cytokinesis, but not growth.

J. Cell Biol. 138, 605-614.

Varney, T. R., Casademunt, E., Ho, H. N., Petty, C., Dolman, J., and Blumberg, D. D. (2002). A novel Dictyostelium gene encoding multiple repeats of adhesion inhibitor-like domains has effects on cell-cell and cell-substrate adhesion. Dev. Biol. 243, 226-248.

Vasiev, B., and Weijer, C. J. (1999). Modeling chemotactic cell sorting during Dictyostelium

discoideum mound formation. Biophys. J. 76, 595-605.

Verkerke-van Wijk, I., Fukuzawa, M., Devreotes, P. N., and Schaap, P. (2001). Adenylyl cyclase A expression is tip-specific in Dictyostelium slugs and directs StatA nuclear translocation and

CudA gene expression. Dev. Biol. 234,151-160.

Vogel, G. (1987). Endocytosis and recognition mechanisms in Dictyostelium discoideum.

Methods Cell Biol. 28, 129-137.

Wang, J., Hou, L., Awrey, D., Loomis, W. F., Firtel, R. A., and Siu, C. H. (2000). The membrane glycoprotein gp150 is encoded by the lagC gene and mediates cell-cell adhesion by heterophilic binding during Dictyostelium development. Dev. Biol. 227, 734-745.

Webb, S. E., Pollard, J. W., and Jones R. E. (1996). Direct observation and quantification of macrophage chemoattraction to the growth factor CSF-1. J. Cell Sci. 109, 793-803.

Wilkins, A., and Insall, R. H. (2001). Small GTPase in Dictyostelium: lessons from a social amoeba. Trends Genet. 17, 41-48.

Williams, J. G., Duffy, K. T., Lane, D. P., McRobbie, S. J., Harwood, A. J., Traynor, D., Kay, R.

R., and Jermyn, K. A. (1989). Origins of the prestalk-prespore pattern in Dictyostelium development. Cell 59, 1157-1163.

Witke, W., Schleicher, M., and Noegel, A. A. (1992). Redundancy in the microfilament system:

abnormal development of Dictyostelium cells lacking two F-actin cross-linking proteins. Cell

68, 53-62.

Wong, E. F., Brar, S. K., Sesaki, H., Yang, C., and Siu, C. H. (1996). Molecular cloning and characterization of DdCAD-1, a Ca²⁺-dependent cell-cell adhesion molecule, in Dictyostelium

discoideum. J. Biol. Chem. 271, 16399-16408.

Wu, H., Maciejewski, M. W., Marintchev, A., Benashski, S. E., Mullen, G. P., and King, S. M.

(2000). Solution structure of a dynein motor domain associated light chain. Nat. Struct. Biol.

7, 575-579.

Xu, P., Mitchelhill, K. I., Kobe, B., Kemp, B. E., and Zot, H. G. (1997). The myosin-I-binding protein Acan125 binds the SH3 domain and belongs to the superfamily of leucine-rich repeat proteins. Proc. Natl. Acad. Sci. USA 94, 3685-3690.

Yang, C., Brar, S. K., Desbarats, L., and Siu, C. H. (1997). Synthesis of the Ca²⁺-dependent cell adhesion molecule DdCAD-1 is regulated by multiple factors during Dictyostelium development. Differentiation 61, 275-284.

Zicha, D., Dunn, G., and Brown, A. F. (1991). A new direct-viewing chemotaxis chamber. J. Cell

Sci. 99, 769-775.

FIGURE LEGENDS

FIG. 1. Amino acid sequence and LRR motif structure of the LrrA protein. The full-length

FIG. 2. Developmental expression of the lrrA gene and subcellular localization of the LrrA

FIG. 3. Phenotype and complementation of the lrrA

FIG. 4. Effect of the lrrA null mutation on cell adhesion and actin cytoskeleton. (A)

lrrA