Ras GTPase-Activating Protein Regulation of Actin ... · Institut de Ge´ne´tique et Microbiologie, Universite´ de Paris-Sud, Centre Universitaire d’Orsay, 91405 Orsay, France2;

EUKARYOTIC CELL, Jan. 2008, p. 141–153 Vol. 7, No. 11535-9778/08/$08.00�0 doi:10.1128/EC.00346-07Copyright © 2008, American Society for Microbiology. All Rights Reserved.

Ras GTPase-Activating Protein Regulation of Actin Cytoskeleton andHyphal Polarity in Aspergillus nidulans�†Laura Harispe,1,2,4 Cecilia Portela,1 Claudio Scazzocchio,2,3

Miguel A. Penalva,4* and Lisette Gorfinkiel1#Departamento de Bioquımica, Facultad de Ciencias, Universidad de la Republica, Igua 4225, Montevideo CP11400, Uruguay1;

Institut de Genetique et Microbiologie, Universite de Paris-Sud, Centre Universitaire d’Orsay, 91405 Orsay, France2;Department of Microbiology, Imperial College London, Flowers Building, Armstrong Road, London SW7 2AZ,

United Kingdom3; and Departamento de Microbiologıa Molecular, Centro de Investigaciones Biologicas,CSIC, Ramiro de Maeztu 9, Madrid 28040, Spain4

Received 20 September 2007/Accepted 8 November 2007

Aspergillus nidulans gapA1, a mutation leading to compact, fluffy colonies and delayed polarity establishment,maps to a gene encoding a Ras GTPase-activating protein. Domain organization and phylogenetic analysesstrongly indicate that GapA regulates one or more “true” Ras proteins. A gapA� strain is viable. gapA coloniesare more compact than gapA1 colonies and show reduced conidiation. gapA� strains have abnormal conidio-phores, characterized by the absence of one of the two layers of sterigmata seen in the wild type. gapA transcriptlevels are very low in conidia but increase during germination and reach their maximum at a time coincidentwith germ tube emergence. Elevated levels persist in hyphae. In germinating conidiospores, gapA� disrupts thenormal coupling of isotropic growth, polarity establishment, and mitosis, resulting in a highly heterogeneouscell population, including malformed germlings and a class of giant cells with no germ tubes and a multitudeof nuclei. Unlike wild-type conidia, gapA� conidia germinate without a carbon source. Giant multinucleatedspores and carbon source-independent germination have been reported in strains carrying a rasA dominantactive allele, indicating that GapA downregulates RasA. gapA� cells show a polarity maintenance defectcharacterized by apical swelling and subapical branching. The strongly polarized wild-type F-actin distributionis lost in gapA� cells. As GapA-green fluorescent protein shows cortical localization with strong predominanceat the hyphal tips, we propose that GapA-mediated downregulation of Ras signaling at the plasma membraneof these tips is involved in the polarization of the actin cytoskeleton that is required for hyphal growth and,possibly, for asexual morphogenesis.

Guanine nucleotide binding protein (G-protein)-mediatedsignaling is one fundamental mechanism by which eukaryoticcells adapt to changing environments and extracellular signals.Monomeric, typically 21-kDa members of the Ras (rat sar-coma) superfamily (for reviews, see references 5 and 76) rep-resent a major class of G-proteins that includes members of theArf-Sar, Rho, Rab, Ran, and Ras families. Like the structurallyrelated � subunits of heterotrimeric G-proteins, Ras superfam-ily members alternate between GTP- and GDP-bound statescorresponding to different conformations. The GTP-boundconformation represents the active state, leading to productiveinteractions with downstream effector proteins, whereas theGDP-bound conformation represents the inactive state. Gua-nine nucleotide exchange factors (GEFs) bind GDP-boundG-proteins and catalyze replacement of GDP by GTP to acti-vate the molecular switch. Conversely, as G-protein signalingterminates when bound GTP is hydrolyzed to GDP and Pi, therate of GTP hydrolysis determines the lifetime of the activatedstate. Most G-proteins themselves have the ability to hydrolyze

bound GTP, but their intrinsically slow GTPase activity is stim-ulated by GTPase-activating proteins (GAPs), which criticallyregulate G-protein function (68).

Potentially oncogenic mammalian Ras proteins are thefounding members of the “true” Ras protein subfamily (47).Mammalian “true” Ras proteins have been intensively studiedbecause its deregulation results in malignant cell transforma-tion. However, “true” Ras proteins are also present in theproteomes of fungi, where they have been shown to play im-portant roles in cell growth and morphogenesis. In Saccharo-myces cerevisiae haploid cells, “true” Ras proteins respond tonutritional signals and couple cell growth to nutrient availabil-ity. While Saccharomyces cerevisiae Ras2p is involved in glu-cose sensing by signaling through the cyclic AMP-protein ki-nase A pathway, it is well established that yeast Ras proteinshave morphogenetic functions, mediated by the Rho familymember Cdc42 (reviewed in reference 43), which plays a keyrole in cell polarization through its regulation of the actincytoskeleton organization. In diploid cells, Ras2p signalsthrough a mitogen-activated protein kinase (MAPK) moduleto promote pseudohyphal growth under conditions of nutrientstarvation. Signaling to the MAPK module is mediated byCdc42p via its effector kinase, Ste20p (49). In agreement withthis Ras morphogenetic role, Ho and Bretscher demonstratedthe involvement of Ras in regulating the actin cytoskeleton byshowing that ras2� results in loss of Cdc42p and F-actin po-larity at 37°C (32). In turn, actin cytoskeleton dynamics and the

* Corresponding author. Mailing address: Centro de InvestigacionesBiologicas, CSIC, Ramiro de Maeztu 9, Madrid 28040, Spain. Phone:34918373112, ext. 4358. Fax: 34915360432. E-mail: [email protected].

† Supplemental material for this article may be found at http://ec.asm.org/.

# Deceased 9 October 2003.� Published ahead of print on 26 November 2007.

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Ras pathway are mechanistically linked via the actin regulatoryfactor Srv2p/cyclase-associated protein (24). In Schizosaccha-romyces pombe, Ras1 mediates the pheromone-dependentmating response by signaling through a MAPK pathway andindependently controls the elongated shape of fission yeastthrough Cdc42 because the fission yeast Cdc42 exchange factorScd1 is a Ras1 effector (10). ras1� mutants and those specifi-cally deficient in the Ras1/Scd1 pathway are abnormallyround (23).

In view of the morphogenetic role that “true” Ras proteinsplay in these ascomycete yeasts, the involvement of Ras infilamentous, polarized fungal growth and development came asno surprise. NC-ras2 regulates hyphal growth in Neurosporacrassa (37). In Ustilago maydis, expression of dominant activeRas2 or its activating GEF Sql2 promotes filamentation (50).Dominant active Ras2 promotes Magnaporthe grisea appresso-rium formation (58). Dominant active Candida albicans Ras1promotes filamentation and is required for the morphogeneticswitch (and thus, for pathogenicity) (41, 65). Cryptococcus neo-formans Ras1, which additionally activates the pheromone re-sponse pathway, is required for filamentation and for growth at37°C (and thus, for pathogenicity of this basidiomycete) (2).

However, our understanding of the mechanisms by whichRas regulates morphogenesis in “true” filamentous fungi isrelatively limited. Filamentous fungi usually contain a pair of“true” Ras paralogues. Most filamentous ascomycetes and ba-sidiomycetes undergo morphological switches and develop-mental processes involving relatively complex specialized re-productive structures, and Ras paralogues may show partiallyoverlapping contributions to these processes. Moreover, therole of Ras proteins is not restricted to the regulation of mor-phogenesis. In N. crassa, the characterization of the Ras1bd

dominant allele has revealed the involvement of ras1 in circa-dian regulation of regulated conidial formation (4). Last, po-larized filamentous growth involves specialized apical growthregulatory factors that are absent from yeasts (26). One exam-ple is the presence in “true” filamentous fungi of Rac homo-logues. Rac, like their related monomeric G-proteins Cdc42and Rho, is a key regulator of the actin cytoskeleton in mam-malian cells (36), acting downstream of Ras (71). Filamentousfungal Rac proteins reportedly regulate polarized hyphalgrowth (11, 45). Penicillium marneffei CflACdc42 and CflBRac

act downstream of Ras to coordinately regulate polarization inhyphae (7), and in the dimorphic yeast Yarrowia lipolytica, Racis required for hyphal growth, but not for pseudohyphal oryeast growth (34). Notably, in C. neoformans, Cdc42 seems tomediate the function of Ras in polarity establishment andmaintenance in yeast cells, while Rac1 is required for Ras-dependent filamentation, which illustrates how different mor-phogenetic processes are regulated by Ras acting through dif-ferent pathways (51).

One additional level of complexity stems from the fact thatdifferent thresholds of Ras activity may be required at differentdevelopmental/morphogenetic processes within the same or-ganism. In their pioneer study of “true” Ras function in fila-mentous fungi, Som and Kolaparti (74) demonstrated that highRasA activity is required for Aspergillus nidulans conidiosporegermination but prevents the morphogenetic switch from iso-tropic to polarized growth. Their data additionally indicatethat levels of active RasA critically contribute to the fate of

aerial hyphae, such that high levels of RasA determine vege-tative hyphal growth, whereas lower levels promote conidio-phore development (74). Thus, understanding the role of Rasand the relative contributions of downstream Cdc42, Rac, andperhaps other Rho GTPases to filamentous fungal polarity anddevelopment clearly awaits further genetic and biochemicalanalyses.

Recent work in S. pombe emphasized another major aspectof Ras regulation by showing that not only the lifetime of theGTP-bound state (determined by the GAPs [see above]) butalso the site at which Ras is activated determines the actualsignaling. Spatially restricted Ras signaling from endomem-branes determines cell morphology through Scd1-Cdc42 effec-tors, whereas Ras signaling at the plasma membrane throughthe Byr2 MAPK cascade determines mating (53).

Here we identify an A. nidulans gene involved in polarityestablishment and maintenance which encodes a Ras-GAPthat contributes to the normal organization of the polarizedactin cytoskeleton in hyphae. Remarkably, this Ras-GAP lo-calizes to the plasma membrane but strongly predominates inthe apexes, strongly suggesting that spatially restricted down-regulation of Ras at the hyphal tip is an important determinantof hyphal growth.

MATERIALS AND METHODS

Aspergillus nidulans and growth media. Aspergillus nidulans strains are shownin Table 1. Gene symbols are defined at the http://www.gla.ac.uk/ibls/molgen/aspergillus/loci.html website. Standard complete and minimal media for A. nidu-lans were used (14). In minimal medium, the sole nitrogen and carbon sourceswere 5 mM ammonium L-(�)-tartrate and 1% (wt/vol) glucose, respectively,unless indicated otherwise.

Conidiation. To measure the levels of conidiation, spores were inoculated ontominimal medium and incubated at 37°C for 8 days. Three 9.6-cm2 circles wereimprinted onto the plate surface, and conidia were scraped from these areas,resuspended in 10 ml of 0.1% Tween, and counted in triplicate. The number ofspores per ml was divided by 9.6 cm2 to give a value of spores per cm2, and meanvalues and standard errors were calculated.

Genetic methods. Standard genetic methods were used throughout. Thestrains used to map gapA1 are shown in Table 1. Mutagenesis of A. nidulansstrain CS1717 with N-methyl-N�nitro-N-nitrosoguanidine (13) resulted in 95%kill. Mutagenized conidiospores were plated onto medium containing uric acid(0.1 mg/ml) as the sole nitrogen source and grown at 25°C for 5 days. As strainCS1717 carries a null uapA mutation (15), utilization of uric acid depends on theUapC transporter, which is not active at 25°C (16). We attempted to obtain amutant where the UapC transporter was active at 25°C. One colony growingabove the background of the mycelia not utilizing uric acid was identified andisolated. On further testing, the isolated colony (carrying the mutation that wefurther characterized as gapA1) is, due to its compact morphology, an apparentand nonspecific suppressor of every mutation tested, which results in leaky,straggly growth on a given nitrogen source.

DNA and RNA manipulations. Oligonucleotides used in this study are listed inTable S1 in the supplemental material. Total RNA was isolated from restingconidiospores, germlings, or mycelia of A. nidulans as described previously (44),treated with glyoxal, and separated on a 1% agarose gel. A PCR-amplifiedfragment of gapA (see Table S1 in the supplemental material) was used as aprobe in Northern blots. The 3� terminus of the gapA transcript was determinedusing the 5�/3�-RACE kit (Roche Diagnostic, Indianapolis, IN) using the specificoligonucleotide 3RACEgap.

Cloning the A. nidulans gapA gene. A. nidulans LH11 was transformed with thegene library constructed in the autonomously replicating plasmid Prg3-AMA1-Not1 (54), obtained from A. Apostolaki. This plasmid contains the N. crassa pyr4selective marker complementing the A. nidulans pyrG89 mutation. Transformedprotoplasts were plated on selective (minus pyrimidine) minimal medium andincubated at 37°C for 3 days. Transformants with wild-type morphology wereselected. From one of these, a plasmid able to complement the morphologicalphenotype caused by gapA1 was isolated (T1pl11). Primers pRG3F and pRG3R,were used to amplify the boundaries of the insert. These sequences were used to

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identify by BLAST analysis the cognate contig in the A. nidulans genomic data-base. New primers (F2, R2, F3, and R3; see Table S1 in the supplementalmaterial) were designed and used to extend the sequence, and by reiterating thisprocedure, the whole insert of plasmid T1pl11 was obtained. The putative openreading frames of AN4998 and AN4997 were amplified from genomic DNA withprimer pairs pRG3F/fGap and pSec/pRG3R, respectively; the complete insertwas amplified with primers pRG3F/pRG3R. The three PCR products werecloned in pGEM-T Easy (Promega). Strain LH11 was cotransformed with a mixof each of the newly constructed plasmids and the integrative plasmid pPyr4,which contains the pyr4 gene of N. crassa, to determine which of the two genescomplements the gapA1 phenotype.

Deletion of gapA. The gapA� strain was constructed by transforming a pyrG89pabA1 pantoB100 strain (LH29 [Table 1]) with a gene disruption cassette built bydouble-joint PCR (80), using Long Expand polymerase (Roche). The upstream(3,011-bp) and downstream (2,914-bp) flanking regions of gapA gene were am-plified from genomic DNA using oligonucleotides DEPC1 and DEPC2 andoligonucleotides DEPC5 and DEPC6, respectively (see Table S1 in the supple-mental material). The Aspergillus fumigatus pyrG gene, used to replace thecomplete gapA coding region, was amplified using oligonucleotides DEPC3 andDEPC4 (see Table S1 in the supplemental material). The PCR fusion productwas amplified with the nested primer pair DEPC7 and DEPC8, purified with theQiagen PCR purification kit and used in transformation. Protoplasts were platedon selective (minus pyrimidine) minimal medium and incubated at 37°C for 3days. Nine transformants that showed the compact morphology typical of gapA1strains were purified and analyzed by Southern blotting. In all of them, the gapAgene had been replaced by pyrG. Strains with a single integration event wereselected for further analysis.

Construction of a gapA::gfp fusion strain. To generate the gapA::gfp transfor-mation cassette, we used a three-way, PCR-based protocol for C-terminal tag-ging of proteins with green fluorescent protein (GFP) (79). Primers are listed inTable S1 in the supplemental material. The GFP-pyrG cassette was amplified byPCR, using primers GapGFP3 and GapGFP4 and a plasmid containing (Gly-Ala)5-GFP plus A. fumigatus pyrG (kindly provided by S. Osmani) as a template.The second fragment, corresponding to the upstream targeting region (the 3� endin the gapA coding region), was PCR amplified using primers GapGFP1 andGapGFP2 with genomic DNA as a template. The third amplified fragmentcontains the 3� untranslated region and was amplified using primers GapGFP5and GapGFP6. The fusion product was amplified with primers GapGFP1 andGapGFP6 and used to transform A. nidulans strain MAD1425 (see Table 1),kindly provided by B. Oakley.

Microscopic techniques. For staining, coverslips with adhered germlings wereincubated for 5 min at room temperature in a solution containing 60 ng/ml4�,6-diamidino-2-phenylindole (DAPI) from Sigma (diluted in 50% glycerol) or10 �g/ml Calcofluor (fluorescent brightener 28; Sigma). The coverslips were thenrinsed in distilled water and mounted. When indicated, phosphate-buffered sa-line with 4% (vol/vol) paraformaldehyde was used to fix cells prior to staining. Allexperiments were repeated at least three times with essentially identical results.

Immunofluorescence detection of actin was done essentially as described previ-ously (62), using anti-actin C4 monoclonal (ICN Biomedicals Inc.) (1/500) andAlexa Fluor 568-labeled goat anti-mouse immunoglobulin G (Molecular Probes)(1/1,000) as primary and secondary antibodies, respectively. Germlings of strainsexpressing GapA-GFP protein were cultured on the surface of glass coverslipsimmersed in 2.5 ml of appropriately supplemented “watch” minimal medium(WMM) (61) containing 1% glucose (wt/vol) as the sole carbon source. Fluo-rescence was detected with an ORCA-ER digital camera (Hamamatsu) coupledto a Nikon E-600 microscope equipped with 60� and 100� objectives. G2-A,B-2A, and UV-2A Nikon filters were used for red, green, and blue fluorescence,respectively. The subcellular distribution of fluorescence was photographed withan ORCA-ER digital camera (Hamamatsu) driven by Metamorph (MolecularDynamics Inc.). Conidiophore imaging was carried out essentially as describedpreviously (56).

Phylogenetic analysis. A multiple-sequence alignment including residues cor-responding to H-Ras positions 1 through 164 in the 29 protein sequences in-cluded in the tree was constructed using Clustal W. Pairwise amino acid se-quence comparisons were computed using a pairwise-distance (p-distance)model. The phylogenetic tree was constructed with Mega 4.0 (http://www.megasoftware.net) using the neighbor-joining method. All positions containingalignment gaps and missing data were eliminated only in pairwise sequencecomparisons (pairwise deletion option). There were a total of 224 positions in thefinal data set. Bootstrap values represent data from 1,000 replicates.

RESULTS

Isolation and genetic characterization of the gapA1 muta-tion. In a mutagenic screen designed to obtain mutations al-tered in the folding or in the expression at the plasma mem-brane of the UapC purine permease (see Materials andMethods), we serendipitously isolated a mutation leading tocompact colony morphology, which results in nonspecific ap-parent suppression of mutations that lead to straggly, diffusegrowth on a number of nitrogen sources (data not shown). Thismutation was denoted gapA1 (see below for gene designation).The growth rate of gapA1 colonies was considerably reducedcompared to the wild type. Radial colony rates that we deter-mined on complete medium at 37°C were 0.38 � 0.007 mm/hand 0.65 � 0.007 mm/h in gapA1 and gapA� strains, respec-tively (Fig. 1A). Notably, microscopic observation of strainscarrying the gapA1 mutation showed that a proportion of ger-minating conidia do not give rise to a germ tube but continuegrowing isotropically while nuclei undergo mitosis, yieldinggiant, multinucleate spherical cells (see below), suggesting thatpolarity establishment is affected by the mutation. gapA1 seg-regates 1:1 in crosses, is recessive in diploids, and was mappedby mitotic haploidization to chromosome III. Meiotic crossesestablished linkage with cnxH (29 cM), pantoC (25 cM), andgalE (8 cM) in the left arm of chromosome III. Three pointcrosses established that gapA is centromere distal relative togalE, in agreement with its position in contig 1.84 of chromo-some III (see below).

Molecular characterization of gapA. The phenotype of thegapA1 mutation suggested that its main defect was in the estab-lishment and/or maintenance of polarity (see below). The gapAgene was cloned by complementation of the morphologicalphenotype of a pyrG89 gapA1 strain with a genomic library con-structed in the replicative plasmid pRG3-AMA1-Not1 (see Ma-terials and Methods). Four clones among the 20,000 pyrimidine-independent transformants screened showed wild-type colonymorphology at 37°C. Plasmids recovered from these transfor-mants complemented the gapA1 compact colony growth pheno-type and showed overlapping restriction patterns (data notshown). Nucleotide sequencing of one such plasmid revealed that

TABLE 1. Aspergillus nidulans strains used in this work

Strain Relevant genotype Purpose

CS2498 pabaA1 Wild-type controlCS1717 uapA::amdS niiA4 biA1 Mutagenized to obtain

gapA1LG10 gapA1 prnB117 pabaA1 Genetic mapping of gapA1FGSC A622 pantoC3 cnxH3 sC12 Genetic mapping of gapA1LG12 gapA1 pantoC3 pabaA1

yA2Genetic mapping of gapA1

LG13 galE9 sC12 Genetic mapping of gapA1LH11 gapA1 uapA� pyrG89

azgA4 pabaA1pyroA4

Cloning of gapA

LH29 pabaA1 pantoB100pyrG89

Recipient strain fordeleting gapA

LH102 gapA�::pyrG pabaA1yA2

gapA� strain forimmunofluorescence

MAD1425 pyrG89 pyroA4 argB2nkuA�::argB�

Recipient strain for gapA-GFP

MAD1768 gapA::gfp::pyrG pyroA4nkuA�::argB�

Strain with gapA::gfp genereplaced

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the insert corresponds to A. nidulans supercontig 1.84 that, likegapA1, maps to the left arm of chromosome III according to theAspergillus Comparative Database (http://www.broad.mit.edu/annotation/fungi/aspergillus/). The contig 1.84 region containedin the rescued plasmid comprises nucleotides 4311570 through437570 and includes autocalled genes AN4998 and AN4997, re-spectively, encoding a GTPase-activating protein and an A. nidu-lans homologue of S. cerevisiae SEC14. Subcloning and transfor-mation experiments showed that the AN4998 sequence, but notthe AN4997 sequence, is able to complement the gapA1 mutation.

gapA1 maps �118 kb centromere distal to galE (autocalledgene AN4957), which lies within contig 1.84, and �481 kbcentromere distal from cnxH. These physical distances corre-spond to �15 kb per cM, which is larger than the values of 2 to9 kb per cM that have been reported for right arm chromo-some III markers which do not lie in the proximity of thecentromere (18). Physical linkage to galE represents prelimi-

nary evidence that reversion of the compact colony phenotypedid not result from extragenic multicopy suppression.

Domain organization of GapA. The 780-residue GapApolypeptide contains N-terminal RasGAP (PF00616) and C-ter-minal RasGAP_C (PF03836) PFAM domains characterizing reg-ulatory proteins that accelerate the intrinsically slow GTPase ac-tivity of Ras protein family members (Fig. 1C). GapA sharesits domain organization with Schizophyllum commune Gap1,Schizosaccharomyces pombe Sar1, and Dictyostelium discoideumDdRasGAP1p, which indeed represent its closest relatives(64%, 43%, and 29% amino acid sequence identity, respec-tively) among functionally characterized GAP proteins (42, 70,78). The structural bases of human H-Ras GTPase activitystimulation by a cognate GAP are well understood (69). Thepresence in GapA of the four conserved motifs that character-ize catalytically active Ras GAPs (Fig. 2), including the “Argfinger” residue which is critical to promote GTP hydrolysis byH-Ras (69), strongly suggests that the physiological role ofGapA is stimulating the GTPase activity of Ras protein(s).

gapA1 is a partial loss-of-function mutation truncating theprotein upstream of the conserved RasGAP_C domain. Toconfirm that we had cloned the gapA gene and not a multicopysuppressor of the gapA1 phenotype, we sequenced the gapA1allele from three different genetic backgrounds obtained bymeiotic crossing. In each case, we found a frameshifting dele-tion of nucleotides 1718 through 1721 in AN4998, demonstrat-ing that the cloned gene is not an extragenic suppressor. Thisframeshift mutation truncates GapA after residue 536, addinga 36-residue out-of-frame amino acid sequence at the C ter-minus of the mutant protein (Fig. 1C). This mutation does notaffect transcript stability as determined by Northern blotting(see below). As the truncating mutation removes the entireRasGAP_C domain, these data suggest that this domain playsa functionally important role in GapA function (we note thatwe cannot rule out the possibility that the out-of-frame C-terminal tail leads to protein instability).

A complete gapA deletion results in a more extreme pheno-type than the gapA1 deletion does. To address whether gapA1represents a complete loss-of-function mutation, we con-structed a null gapA allele after substituting its complete cod-ing region by the A. fumigatus pyrG gene, using a transforminglinear DNA fragment assembled by double-joint PCR as de-scribed in Materials and Methods. Nine pyrG� strains showingthe characteristic gapA1 compact colony phenotype were iso-lated. All of them carried the expected gene replacementevent, as demonstrated by Southern blot analysis (data notshown). One was selected for further study. The cognate mu-tation will be referred to as gapA�. Both gapA1 and gapA�strains show fluffy growth on some media, noticeably on thosecontaining ammonium as the sole nitrogen source, but thecompact colony phenotype is markedly more conspicuous ingapA� strains than in gapA1 strains, in agreement with theirrelative colony growth rates (0.32 � 0.006 mm/h in gapA�compared to 0.38 � 0.007 mm/h in gapA1; this gapA� growthrate is less than half that of the wild type [see above]). Fluffygrowth denotes an abnormally high number of aerial non-conidiating hyphae. However, this phenotype differs from thatpreviously described for fluffy mutations (1) in that it is notinvasive and the aerial nonconidiating hyphae are confined toa compact colony (not shown). Unlike the boundaries of

FIG. 1. Genetic and molecular characterization of gapA. (A) Wild-type, gapA1, and gapA� strains inoculated on Aspergillus completemedium and incubated for 5 days at 37°C. (B) Genetic and physicalmapping of gapA. Genetic distance (in centimorgans) and physicaldistance (in kilobases) are shown above and below the schematicallydepicted chromosome III (Chr III), respectively. gapA is AN4998, galEis AN4957, and cnxH is AN4841 in the A. nidulans genomic database.pantoC has not been annotated. The pantoC-cnxH distance wasindependently determined to be 18 cM (3). The circle represents thecentromere, and the filled bar on the left indicates the telomere.(C) Domain structure of GapA and the truncated gapA1 product.The RasGAP (hatched box) and RasGAP_C (gray box) domains cor-respond to SM00323 and PF03836, respectively. Deletion of four basesin the gapA1 allele (from positions 1718 to 1721 of the nucleotidesequence) results in a GapA protein truncated after residue 536 (in-dicated by an arrow), containing a 35-residue C-terminal peptide trans-lated in one incorrect reading frame (filled box in the diagram andlowercase letters for amino acid single-letter codes). The asterisk de-notes a stop codon. aa, amino acids.



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gapA� and gapA1 colonies, those of gapA� strains are irregu-lar. gapA� additionally results in a �50% reduction in conid-iospore production compared to isogenic gapA� and gapA1strains (7.4 � 107 � 0.1.1 � 107 in the gapA� strain comparedto 1.5 � 108 � 0.22 � 108 conidiospores per cm2 in the wildtype and in the gapA1 mutant). We concluded that gapA1 is apartial loss-of-function mutation, and therefore, all subsequentphenotypic analyses were carried out with gapA� strains.

Bioinformatic analysis of A. nidulans Ras and Ras GAPs. Inview of the fact that gapA encodes a Ras GAP, we reexaminedthe Ras content of A. nidulans in the context of the completegenome sequence, using TBLASTN searches to identify puta-tive Ras family members. We confirmed the presence of two“true” Ras proteins corresponding to the previously describedA. nidulans/A. fumigatus RasA and RasB (21, 22, 74) andadditionally identified two as-yet-unreported proteins with sig-nificant similarity to Ras. To determine their closest fungalrelatives, we carried out a phylogeny analysis, using Ras-re-lated Rheb orthologues as an outgroup to construct the phy-logenetic tree shown in Fig. 3, as Rheb proteins are proteinsrelatively distant to “true” Ras and the A. fumigatus Rheborthologue has been characterized (55). RasA and RasB clus-ter with “true” Ras proteins, including human H-Ras (Fig. 3,branches 1 and 2). One of the two as-yet-undescribed “Ras-

like” proteins is clearly grouped (branch 3) with S. cerevisiaeRsr1p (Bud1p), which is involved in bud site selection (39, 66)and apparently corresponds to the A. nidulans orthologue of N.crassa Krev-1, a Ras-like protein that does not appear to playa role in hyphal growth (35). The second protein (the AN3434product [branch 4]) is more distantly related to RasA/B and,notably, appears to be specific of aspergilli (data not shown).

As reported previously (70), we detected only three RasGAP-like proteins in A. nidulans. Domain organization andphylogenetic analysis clearly separate the GapA/Gap1/Sar1fungal subfamily from the AN9463 and AN3735 products rep-resenting the two other Ras GAPs in the A. nidulans proteome(70). AN3735 contains lipid-binding C2 and RasGAP domainsand is the almost certain homologue of Bud2p, the S. cerevisiaeGAP for Rsr1p. Thus, in all likelihood, AN3735 is the GAPregulating the sole A. nidulans Rsr1p homologue. AN9463contains calponin homology (CH) and IQ calmodulin bindingdomains, in addition to RasGAP and RasC-ter domains, mak-ing this protein a clear relative of human IQGAP1 and S.pombe Rng2, a potential effector of the Rho family of GTPasesand a component of the actomyosin ring involved in cytokinesis(17). IQGAP proteins are Ras GTPase-related proteins with-out actual Ras GTPase activity that function as Cdc42 effectors(8, 31).

FIG. 2. Multiple-sequence alignment including representative fungal and metazoan GAP proteins. The four amino acid sequence blocks character-istic of Ras GAPs (see Fig. 3 in reference 69) are boxed. The fully conserved “Arg finger” residue that plays a critical role in catalysis is indicated by anarrow. Numbers on the right indicate amino acid residue numbering. Abbreviations and GenBank accession numbers (in parentheses) are as follows: A.nidulans GapA (AAO38800); Sp_GAP1, S. pombe Gap1 (NP_595370); Sc IRA1 and Sc IRA2, S. cerevisiae Ira1p (P18963) and Ira2p (CAA99093),respectively; dmGAP1, Drosophila melanogaster GAP1 (M86655); rrGAP1m, Rattus rattus GAP1m (BAA06398); ceGAP1, Caenorhabditis elegans GAP1(NP_509594); HsNF1, Homo sapiens NF1 (P21359); Hs p120GAP, H. sapiens p120 GAP (P20936). Gaps introduced to maximize alignment are indicatedby dashes.



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The third A. nidulans Ras GAP, GapA, is clearly the ortho-logue of S. pombe Sar1p (78). Sar1p is the GAP for Ras1p (31),the single fission yeast “true” Ras, whose closest homologue inA. nidulans is RasA. Thus, although we have not made anyattempt to determine the specificity of GapA, it is very likelythat this protein is a negative regulator of RasA and, possibly,of RasB, as GAPs show little specificity with regard to “true”Ras proteins (5).

GapA is involved in conidiophore development. One pre-dicted consequence of gapA� is Ras hyperactivation. In A.

nidulans, a threshold of RasA activity is required for the initi-ation of conidiophore development. This activity must subse-quently decrease for progression of aerial hyphae through thedevelopmental pathway (74). In A. fumigatus, where RasA andRasB play different but overlapping developmental roles, dom-inant active RasA expression reduces conidiation and leads tomalformed conidiophores, while expression of a dominant neg-ative rasB allele promotes ectopic conidiophore development(21). These reports led us to examine the morphology of con-idiophores, which are indeed abnormal (Fig. 4A and B). In thewild type, primary sterigmata, named metullae, bud directlyfrom the conidiophore vesicle (Fig. 4A). These metullae giverise to a layer of secondary sterigmata, named phialides, fromwhich conidia are originated (Fig. 4A). These two layers ofsterigmata seen in the wild type are clearly absent in the mu-tant, where a single layer of abnormal sterigmata from whichconidia bud directly was clearly observed (Fig. 4C). In somecases, it seems that cells that morphologically resemble conidiawould appear to arise directly from the vesicle without theformation of metullae or phialides (Fig. 4C). However, weclearly observed in the mutant examples of abnormal, sphericalsterigmata whose diameter was similar or slightly larger thanthat of conidiospores (Fig. 4D, mutant conidiospore showingseveral abnormal, spherical sterigmata; an ellipsoid sterigmataarising from the same vesicle is also indicated). The presence of

FIG. 3. Phylogenetic analysis of Ras proteins in filamentous fungi andyeasts. “True” Ras and “Ras-like” proteins are included in clades 1through 4 (numbers within black circles). Ras-related Rheb proteins(clade 5) were used as an outgroup. Mega 4.0 software was used forconstructing the phylogenetic tree (details in Materials and Methods).The numbers beside the nodes are bootstrap values corresponding to1,000 replicates. Hypothetical proteins are denoted by Entrez databaseaccession numbers. In those cases where the gene/protein has been de-scribed, protein designations have been given (for all A. nidulans proteins,the systematic gene number is also included). Entrez database accessionnumbers for the proteins are as follows: H. sapiens p21 H-ras, P01112; S.cerevisiae Ras1, P01119; S. cerevisiae Ras2, P01120; S. cerevisiae Rhb1,P25378; S. cerevisiae Rsr1, P13856; N. crassa Ras1, CAA37612; N. crassaRas-2, BAA03708; N. crassa NCU01444, XP_955966; N. crassa Krev-1,BAA32410; U. maydis Ras1, AAO19640; U. maydis Ras2, AAO19639; U.maydis UM02047, XP_758194; U. maydis UM05654, EAK86057; S.pombe Ras1, P08647; S. pombe Rhb1, NP_595194; Ashbya gossypiiAAS51658, AAS51658; A. gossypii Rsr1p, AAS53835; M. grisea Ras2,XP_369310; M. grisea MGG06914, XP370417; M. grisea Ras1, XP_364654; M. grisea MGG02727, XP_366651; A. nidulans RasA, XP_657786(AN0182); A. nidulans RasB, XP_663436 (AN5832); A. nidulans AN3434,XP_661038; A. nidulans AN4685, XP_662289; A. nidulans RhbA,XP_682137 (AN8868); P. marneffei RasA, AAO64439; C. neoformansRas1, AAD55937; and C. neoformans Ras2, AAG10598. Sequence simi-larities were scored using the Blosum62 matrix. H. sapiens, Homo sapiens.

FIG. 4. Deletion of GapA affects conidiophore development.(A) Wild-type conidiophore. v, vesicle; m, metullae; p, phialides; c,conidia. (B) Mutant conidiophore. (C) Mutant conidiospore whereconidia arise from primary sterigmata (white arrowheads). A normalprimary sterigmata is not discernible in the spore chain indicated witha white star. c, conidia. (D) Mutant conidiospore showing severalabnormally round-shaped metullae (white stars) and an ellipsoid one(white arrow). Bars, 10 �m.



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these class of spherical sterigmata precluded us from reaching afirm conclusion on the possible absence of sterigmata in somemutant spore chains.

Despite the developmental effects of the gapA� mutation,we observed no difference in conidiospore viability betweenthe wild type and the mutant. In the wild type, a single nucleusmigrates into each bud during budding of primary sterigmatafrom the vesicle, and thus, daughter phialides and conidio-spores are always uninucleate. However, about 6% of thegapA� conidiospores have two nuclei and spores with threenuclei were occasionally seen. We found no significant differ-ence in conidiospore diameter between the wild type and themutant when the whole population of gapA� conidiosporeswas considered (wild type, 3.5 � 0.2 �m; gapA�, 3.5 � 0.5 �m).However, gapA� binucleate spores were clearly larger (4.6 �0.6 �m in diameter; n 15).

gapA transcript levels reach their maximum during polarityestablishment. A. nidulans conidiospores contain a single nu-cleus arrested in G1. During the initial stages of germination,conidiospores undergo a period of isotropic growth and reen-ter the nuclear division cycle. In response to poorly understoodsignals, germinating conidiospores establish a polarity axis andswitch from isotropic to polarized growth, which leads to theemergence of a germ tube (26, 28, 48, 77). Forced, high-levelexpression of mutant hyperactive RasAG17V in germinatingconidiospores leads to large, swollen multinucleated cells thatdo not proceed any further (20, 74), indicating that high Rasactivity impairs polarity establishment. Thus, we monitoredgapA transcription in germinating conidiospores, using a syn-thetic minimal medium containing 10 mM ammonia and 1%glucose as the sole N and C sources, respectively. Under theseconditions, germ tube emergence roughly coincides with theonset of the first mitosis (data not shown) and has essentiallytaken place in all germinating conidiospores after 7 h at 37°C(see also below). Figure 5 shows that gapA transcript levels,which are very low in conidiospores, increase during the iso-tropic growth phase (overlapping the 3-h time point) to reachtheir maximum at the 6-h time point (roughly corresponding topolarity establishment/germ tube emergence). Transcript lev-els remained essentially unchanged and unaffected by thegapA1 mutation after 17 h of incubation (Fig. 5), when germ-lings had already given rise to hyphae. These data, in conjunc-tion with previous work (74), are consistent with involvementof GapA in polarity establishment and suggested that GapAmight play an additional role in hyphal cells.

Isotropic growth is uncoupled from spore polarization in agapA� strain. After a 7-h incubation under the above condi-

tions, all wild-type germinating conidiospores (n 130) hadundergone the morphogenetic switch. These germlings con-tained two or four nuclei (Fig. 6A and E, wild-type strain). Theeven number of nuclei is in agreement with the report that thefirst three nuclear divisions occur simultaneously (29). In starkcontrast, two classes were seen in the mutant population at thesame time point. Nearly half of gapA� conidiospores (43%;n 130) had germinated (“swelled”) but had not given rise toa germ tube and contained a variable number of nuclei, rang-ing from one to eight (Fig. 6D) (note that approximately 1/10of these mutant swelled conidiospores have not undergone thefirst nuclear division). gapA� spores in the second class (57%)had led to a germ tube but showed delayed polarity establish-ment with apparent uncoupling of isotropic growth from po-larity establishment and from nuclear division (Fig. 6E, gapA�strain). Delayed polarity establishment after an abnormallylong isotropic growth was evident because the size of the mu-tant swelled conidiospores (6.5 � 1.5 �m) was significantlylarger than in the wild type (4.5 � 0.3 �m), as determinedusing two-sample t and Wilcoxon-Mann-Whitney tests (P val-ues of 6.1 � 1021 and 1 � 1012, respectively). In contrast, themutant germ tubes were noticeably shorter (Fig. 6A and B).Finally, some mutant conidiospores did not give rise to a germtube even after a 16-h incubation at 37°C, thus forming giantmultinucleated spores resembling those described by Som andKolaparthi (74). The diameter of these giant spores in somecases approaches 15 �m (Fig. 6C), and these cells may containas many as 11 nuclei. Uncoupling of polarity establishment andnuclear division/migration was indicated by the broad distribu-tion in the number of nuclei that we found in gapA� germlings,with a significant proportion of them (37%) having eight nu-clei, many of which had not yet migrated into the germ tube(Fig. 6B and E). This was in striking contrast to the predom-inating single class in the wild type, containing two or fournuclei evenly spaced along the complete length of the germling(Fig. 6A and E).

The absence of gapA allows spore germination in the ab-sence of carbon source. In S. cerevisiae, Ras is involved incarbon source sensing and couples cell growth to nutrientavailability (for a review, see reference 67). Osherov and May(54) demonstrated that RasAG17V induces conidial germina-tion in the absence of a carbon source. In their experiments,mutant conidia underwent swelling and arrested growth beforethe morphogenetic switch. We repeated these experiments us-ing our gapA� allele. Both wild-type and gapA� conidiosporesgerminated in the presence of 10 mM glucose. However, whilegapA� conidiospores underwent swelling in the absence of acarbon source, no isotropic growth was detected for the wild-type conidiospores after a 14-h incubation at 37°C under thesame conditions (average diameters of gapA� and gapA� con-idiospores were 4.9 � 0.9 �m and 3.5 � 0.3 �m, respectively[Fig. 7]). In addition to conidiospore germination, gapA� ac-tivates the nuclear division cycle in the absence of glucose, as30% of the swelled gapA� conidiospores contained two or fournuclei (Fig. 7; note that only 6% of mutant conidiospores werebinucleated before incubation [see above]). These data closelyresemble those obtained with RasAG17Vconidia (54) and thusstrongly support our contention that RasA is a GapA sub-strate. We additionally observed that 5% of wild-type sporesthat do not swell in the absence of glucose gave rise to an

FIG. 5. Transcript analysis of gapA mRNA during conidial germi-nation. Transcript levels were determined by Northern blot analysisusing an specific gapA probe and 18S rRNA as loading control. Theexperiment was carried out at 37°C. con, control.



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abnormally narrow, abortive germ-tube primordium (Fig. 7).This observation appears to suggest that isotropic growthmight not be an absolute requirement for polarity establish-ment/selection of a polarity axis.

GapA is required for polarity maintenance and signals tothe actin cytoskeleton. The most conspicuous phenotype ofgapA� strains is a marked defect in polarity maintenance.Sixty-two percent of the mutant germlings (n 117) showedmorphological abnormalities (Fig. 8C to F), including an ab-normal high frequency of apical branching, randomly localizedincreases in hyphal diameter, and loss of polarity which insome cases led to “budding yeast-like” germlings that resemblea class of abnormal cells seen in A. nidulans swoF mutantsaffected in conidiospore swelling (Fig. 8D) (73).

Germinating A. nidulans conidia typically give rise to a sec-ond germ tube after the primary polarity axis has been estab-lished. The normal pattern of germ tube emergence is depen-dent on the actin cytoskeleton (27). In most cases (95% for thewild type in our conditions; n 150), the second germ tubeemerges opposite or nearly opposite (�180°) the first. In aminor proportion of wild-type germlings (5%), the secondgerm tube emerges at �90° relative to the first (quaterpolarpattern [27]). In marked contrast, 23% of the gapA� germlings(n 150) showed a quaterpolar or nearly quaterpolar pattern

(Fig. 8C). We additionally noticed that a proportion of gapA�germlings showed apical swelling, contrasting both with thewild type and with other relatively normal mutant germlings(Fig. 8A, B, and I). Both apical branching and isotropic swell-ing of the tip are indicative of defective organization of theactin cytoskeleton (46, 64, 75).

Ras involvement in the regulation of polarity and the actincytoskeleton in fungi has been reported. In S. cerevisiae, Rasproteins function in the regulation of actin polarity (32) andRas2p signals function through the key actin cytoskeleton reg-ulator Cdc42p to induce diploid filamentous growth (49),which resembles hyphal growth in that it requires a highlypolarized actin cytoskeleton (9). Thus, we examined the actincytoskeleton of A. nidulans gapA� germlings using indirectimmunofluorescence (Fig. 8G to L). In wild-type cells, thisprocedure reveals faintly labeled actin cables and strongly la-beled, highly polarized actin patches (29) (L. Araujo, M. A.Penalva, and E. Espeso, unpublished data) (Fig. 8G and H). In“normal” gapA� germlings, actin cables and patches were stillvisible (Fig. 8I and J). However, a marked loss of actin patchpolarization was clearly seen in these germlings (Fig. 8I and J;compare to wild-type controls in panels G and H). To providea semiquantitative estimation of actin polarization, we mea-sured the relative fluorescence of a region containing the hy-

FIG. 6. Delayed polarity establishment and uncoupling of nuclear division from the morphogenetic switch. Germlings of the wild-type andgapA� strains were cultured in minimal medium for 7.5 h at 37°C, fixed, and stained with DAPI. (A) Wild-type germlings. The merged image ofNomarski (left) and DAPI (center) channel images is shown on the right. (B) gapA� strain (channels as in panel A). (C) An example of giantmultinucleated spores seen with the gapA� strain. Bars, 10 �m. (D) Germinating gapA� conidiospores that had not undergone the morphogeneticswitch (i.e., “isotropically growing cells” without a noticeable germ tube) at the time of the analysis were classified according to their number ofnuclei. While this population involves 43% of the germinated gapA� spores, essentially all wild-type conidiospores have given rise to a germ tubeat the sampled time point. (E) As in panel D, but including only germinated conidiospores that had given rise to a germ tube (“polarized cells”).Note the highly regular pattern observed in wild-type germlings (see panel A) and the relatively increased number of nuclei in the mutant(representing “isotropic” and “polarized” cells, denoted as total germinated conidia). gapA1 strains showed a similar phenotype.



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phal tip compared to an equivalent region at the base of thegerm tube in 30 germlings from different experiments. Weobtained values of 4.13 � 1.6 and 1.8 � 0.4 for wild-type andgapA� germlings, respectively. We conclude that an abnormalpolarization of the actin cytoskeleton, predictably resultingfrom an abnormally high Ras activity, underlies the polaritymaintenance defects shown by mutant germlings.

Branching was markedly more frequent in the gapA� mutantthan in the wild type (4.8% of the mutant germlings showedbranching compared to 0.7% in the wild type [n 700]). Inmalformed gapA� germlings, abnormal apical branching asso-ciated with tip swelling was frequently observed (Fig. 8C andL). Branching in the mutant involves a complete loss of actinpolarization in the tip of the main polarity axis and strongpolarization of actin in the new branch tip (Fig. 8K and L),strongly suggesting that, in the gapA� mutant, activation ofpolarized growth in the branch occurs at the expense of theinactivation of the main polarity axis.

GapA-GFP localizes to the plasma membrane and is polar-ized in hyphae. As the polarity establishment and maintenanceroles of GapA in swelled conidiospores and hyphal tips, re-spectively, appear to involve regulation of the cortical actincytoskeleton, we used a GapA-GFP fusion to determine thesubcellular localization of GapA, using the gene replacementprocedure reported by Yang et al. (79) to express the fusionprotein at physiological levels. GapA-GFP localizes to the cy-tosol and to the cell periphery in swelled conidiospores, germ-

lings, and hyphae (Fig. 9A to C). The cortical distributionshowed some polarization in germlings (Fig. 9B) but was mark-edly polarized in hyphae (Fig. 9C), in agreement with therequirement of GapA in polarity maintenance describedabove. GapA-GFP additionally localized to septa (Fig. 9D).These and the above data strongly indicate that GapA is re-quired to downregulate Ras activity at or near the plasmamembrane and that this downregulation predominates at thecortical regions of hyphal tips and at septa, where the actincytoskeleton plays major roles.

DISCUSSION

This work shows that an A. nidulans Ras GAP plays animportant role in polarity establishment and maintenance andis required for conidiophore development. Polarized A nidu-lans growth is crucially dependent on the actin and microtu-bule cytoskeletons (30, 33, 38, 64, 75). The actin cytoskeletonis highly polarized (29) (Araujo, Penalva, and Espeso, unpub-lished). The absence of GapA results in actin cytoskeletondisorganization and depolarization defects.

In relatively young germlings, these defects correlate with anabnormally high frequency of apical branching in the proximityof an abnormally swelled tip where the actin cytoskeleton or-ganization has been fully lost, strongly indicating that apicalbranching occurs as a result of loss of actin organization in themain growth axis tip. In longer hyphae, GapA deficiency re-sults in a marked loss of F-actin polarization. In contrast, thegapA deletion has no obvious effect in microtubule organiza-tion as assessed with wild-type and mutant strains expressing a�-tubulin–GFP fusion (data not shown).

Predominance of tip swelling/apical branching in relativelyyoung germlings suggests that accurate regulation of the actincytoskeleton plays a more prominent role in these cells than inhyphae. Notably, Horio and Oakley have reported significantdifferences in growth rate and mitotic behavior of cytoplasmicmicrotubules between these two types of cells (33). Microtu-bules are not required for polarity establishment and germtube emergence (52) but appear to be involved in focusingactin at the tips of growing hyphae (33). Thus, it is likely thatapical actin organization is more sensitive to the absence ofGapA function in germlings than in hyphae, where the contri-bution of other factors like microtubules reaching the hyphaltip might to some extent compensate for the consequences ofthe GapA deficiency, somehow preventing the complete dis-organization of apical actin.

Bioinformatic analyses strongly indicate that GapA is a“true” Ras-specific GAP, and thus, gapA� would be expectedto increase the activated lifetime of its Ras substrate(s), result-ing in abnormally persistent signaling. Aspergilli contain twoRas genes, rasA and rasB, encoding the only “true” Ras pro-teins in the A. nidulans genome and the major candidates tomediate the physiological role of GapA. A. nidulans rasB hasnot yet been characterized. However, the findings that A. nidu-lans rasA is essential (74) whereas A. fumigatus rasB is not (22)and that, in A. fumigatus, rasA is expressed at relatively highlevels whereas expression of rasB is very low (21) would suggestthat RasA plays a more important physiological role.

Several arguments strongly suggest that GapA plays its phys-iological roles by regulating at least RasA. (i) gapA� promotes

FIG. 7. Mutant gapA� conidiospores germinate in the absence ofglucose. Conidia were fixed and stained with DAPI after 14 h ofincubation at 37°C in minimal medium without a carbon source, acondition which prevents germination in the wild type (gapA�) but notin the gapA� mutant (gapA�). Nomarski and DAPI-stained images areshown. While the wild-type spores do not germinate in the absence ofa carbon source, mutant spores undergo isotropic growth and activatenuclear division. We noticed that a minor proportion of wild-typespores gave rise to an abnormally slender and abortive germ tube(indicated by a white arrow in the Nomarski image). DIC, differentialinterference contrast. Bar, 5 �m.



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vegetative growth of aerial hyphae and reduces the productionof conidiospores, resembling the consequences of expressingA. nidulans dominant active RasA at moderate levels (74). (ii)In A. fumigatus, expression of dominant active RasA at phys-iological levels leads to reduced growth and conidiation, pro-motes the formation of aerial hyphae, and results in mal-

formed conidiophores, affecting the morphology of sterigmata.These phenotypes are strikingly similar to those of A. nidulansgapA�. (iii) In P. marneffei, RasA regulates polarized hyphalgrowth (7). (iv) During A. nidulans germination, morphogen-esis is coordinated with nuclear division (25). One phenotypicconsequence of gapA� is the uncoupling of germ tube emer-

FIG. 8. The actin cytoskeleton phenotype resulting from the gapA� mutation. (A) Nomarski image of a wild-type germling. (B) Example of agapA� germling showing abnormal apical swelling. (C) Abnormal germling showing two germ tubes, one of which shows apical branching.(D) Abnormal, budding yeast-like germling. (E and F) Calcofluor staining of gapA� germlings showing the positions of the septa and abnormallyswelled regions (white arrowheads in panel E), often associated with apical branching. (G and H) The wild-type actin cytoskeleton, as observedby indirect immunofluorescence using a mouse anti-actin monoclonal antibody. In panel G, the positions of two actin cables are indicated withblack arrows. Actin patches predominate in the apical regions (black stars), where they lead to a much stronger signal than that of actin cables.(Note that these apical pixels become saturated when images are contrasted to observe actin cables). (I) The strong actin signal at the tip and themarked polarization of actin patches seen in the wild type are lost in mutant germlings. The black arrow indicates an actin cable. (J) An exampleof apical actin depolarization in a germling with an abnormally swelled tip. (K) Activation of a subapical branch leads to loss of actin polarizationat the main tip. (L) Predominance of actin patches at the tip of an abnormal subapical branch; note that the main tip is clearly swelled and showslittle actin polarization. Bars, 5 �m.



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gence from nuclear division, such that mutant germlings con-tain, on average, more nuclei than the wild type. A similarphenotype has been observed in A. fumigatus after expressionof a dominant active RasA protein at physiological levels (21).(v) We show that gapA� conidia are able to germinate in theabsence of a carbon source. According to Osherov and May(54), this is a distinctive phenotype of A. nidulans conidiaexpressing high levels of dominant active RasA. Notably, thisphenotype is also shown by A. fumigatus conidia expressingphysiological levels of dominant active RasA (21). (vi) Finally,in both A. nidulans (20, 54, 74) and A. fumigatus (21), theformation of giant multinucleated spores resulting from de-layed polarity establishment that we observed in gapA� conidiais a typical phenotype of conidia expressing dominant activeRasA. However, despite the above evidence that GapA acts, atleast in part, by regulating Ras activity, we cannot exclude thepossibility that GapA has Ras-independent functions, likethose played by the mammalian p120 Ras GAP in the regula-tion of cell motility (40).

GapA-GFP shows cytosolic and cortical localization, at orvery near the plasma membrane. However, while GapA doesnot appear to be polarized in very young germlings, it becomespolarized in longer germlings and polarization is very conspic-uous in hyphal cells, in agreement with its involvement inpolarity maintenance and with the depolarization of the actincytoskeleton seen in gapA� strains. GapA has no evident mem-brane-targeting domains, suggesting that it may be recruited toits cortical localization indirectly, perhaps by its cognate Ras.Ras is known to be targeted to the endoplasmic reticulum,Golgi bodies, and plasma membranes trough posttranslational

modification (12), but the subcellular localization of A. nidu-lans RasA and RasB remains to be investigated.

A. nidulans GapA localization to a crescent at the hyphal tipand to septa resembles the localization of the single A. nidulansformin and tropomyosin proteins, respectively, denoted SepAand TmpA (60, 72). In S. cerevisiae, null alleles of RAS2 andTPM1 (encoding tropomyosin) are synthetically lethal, whichstrongly suggests that these genes are involved in a commoncellular pathway.

A well understood example illustrating how a Ras familymember can act directly upstream of the key actin cytoskeletonregulator Cdc42 takes place during S. cerevisiae vegetativegrowth, where Cdc42p recruitment/activation to the nascentbud site is directed by the bud site selection Ras-like G-proteinRsr1p/Bud1p (a Rap1A subfamily member and thus, not a“true” Ras). Cdc42p and its GEF (Cdc24p) are effectors ofRsr1-GTP (39; for reviews, see references 59 and 63). Recruit-ment of Cdc42p to sites of polarized growth underlies symme-try breaking/polarity establishment (81), as Cdc42p subse-quently recruits the Bni1p formin to these sites to locallynucleate actin cables mediating targeted delivery of secretoryvesicles (19).

In mammalian cells, “true” Ras proteins are upstream reg-ulators of Rho family proteins, regulating the activation statesof RhoA, Rac1, and Cdc42. “True” Ras effectors acting infungal morphogenesis have been thoroughly investigated bothin the yeasts S. cerevisiae (for a review, see reference 59) and S.pombe (53). It is well established that S. cerevisiae Ras2p sig-nals through Cdc42 to promote pseudohyphal growth (43),which involves a marked polarization of the actin cytoskeleton

FIG. 9. Subcellular localization of GapA-GFP. (A) Subcellular localization of GapA-GFP in young germlings, shortly after germ tubeemergence. Bar, 10 �m. (B) Subcellular localization of GapA-GFP in longer germlings. Bar, 10 �m. (C) Marked polarization at the hyphal tip(GFP-stained and Nomarski images shown). Bar, 5 �m. (D) GapA-GFP localization to septa. GFP and Calcofluor images are shown. Bars, 5 �m.



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(9). In S. cerevisiae yeast cells, Ras2p is involved in cell cycle-dependent actin polarization and in the localization of Cdc42pto the bud tips (32). Thus, Cdc42 would be a likely candidateto mediate the effects of gapA� on actin organization. How-ever, unlike yeasts, filamentous fungi contain Rac homologues,and these have been shown to act in combination with Cdc42to regulate different aspects of cell morphology in Penicilliummarneffei and C. neoformans (see the introduction; also seereferences 6 and 51). Therefore, we anticipate that the eluci-dation of the specific Ras effector(s) acting at each morpho-genetic or developmental decision will require intensive re-search combining genetic, biochemical, and (as illustratedhere) subcellular localization studies.

Involvement of a Ras GAP in fungal morphogenesis was firstreported for the Schizophyllum commune protein Gap1 (70).Deletion of the cognate gene mainly affects the sexual cycle,and thus, the dikaryon between two �gap1 strains is unable toyield fertile basidiospores. However, the pleiotropic phenotypeof the mutant strongly suggests that Gap1 is also involved inpolarity maintenance.

Spatial regulation of Ras during fungal morphogenesis hasbeen reported for Ras1 of S. pombe, where the Efc25 GEFregulates Cdc42-dependent Ras signaling from endomem-branes to determine elongated cell morphology, whereas theSte6 GEF regulates MAPK-dependent Ras signaling from theplasma membrane to control mating (53, 57). The subcellularlocalization of GapA in growing hyphae strongly suggests thatdownregulation of Ras signaling in a plasma membrane do-main located at the hyphal tip plays an important role inpolarized hyphal growth. Such local downregulation of Ras byGAP would provide a second way by which spatially restrictedRas signaling might be achieved.

ACKNOWLEDGMENTS

We thank Ricardo Ehrlich, Alberto Rosa, and Ana Ramon forsupport and laboratory facilities in Oruguay; Elena Reoyo for techni-cal assistance; Eduardo Espeso and Herb Arst for critical reading ofthe manuscript; Javier Valdez-Taubas and Christine Drevet for helpfuldiscussions; Lidia Araujo and Olga Rodrıguez-Galan for help withactin immunofluorescence detection; Bernard Labedan and JorgeGraneri for help with phylogeny and statistical analyses, respectively;Stephen Osmani and Berl Oakley for plasmids and strains; and oneanonymous referee for helpful suggestions.

This work was supported by the DGCYT (Spain) through grantBIO2006-0556 to M.A.P., the International Foundation for Science(Stockholm, Sweden) through a grant to L.G. (Uruguay), and theFrench CNRS, the Universite Paris-Sud, and the Institut Universitairede France through grants to C.S. Cooperation between the UniversiteParis-Sud and the Universidad de la Republica was supported byECOS-Sud project 00B01 to L.G. and C.S. L.H. was partially sup-ported by the Direction des Relations Internacionales (Universite Par-is-Sud) and by the Agencia Espanola de Cooperacion Internacional(Spain).

ADDENDUM IN PROOF

A recent paper (A. Virag, M. P. Lee, H. Si, and S. D. Harris, Mol.Microbiol. 66:1569–1596, 2007) addresses the role of Cdc42 and Racproteins in the regulation of A. nidulans hyphal morphogenesis.

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