Logo of eukcellPermissionsJournals.ASM.orgJournalEC ArticleJournal InfoAuthorsReviewers
Eukaryot Cell. Feb 2011; 10(2): 207–225.
PMCID: PMC3067405

Candida albicans Hap43 Is a Repressor Induced under Low-Iron Conditions and Is Essential for Iron-Responsive Transcriptional Regulation and Virulence [down-pointing small open triangle]


Candida albicans is an opportunistic fungal pathogen that exists as normal flora in healthy human bodies but causes life-threatening infections in immunocompromised patients. In addition to innate and adaptive immunities, hosts also resist microbial infections by developing a mechanism of “natural resistance” that maintains a low level of free iron to restrict the growth of invading pathogens. C. albicans must overcome this iron-deprived environment to cause infections. There are three types of iron-responsive transcriptional regulators in fungi; Aft1/Aft2 activators in yeast, GATA-type repressors in many fungi, and HapX/Php4 in Schizosaccharomyces pombe and Aspergillus species. In this study, we characterized the iron-responsive regulator Hap43, which is the C. albicans homolog of HapX/Php4 and is repressed by the GATA-type repressor Sfu1 under iron-sufficient conditions. We provide evidence that Hap43 is essential for the growth of C. albicans under low-iron conditions and for C. albicans virulence in a mouse model of infection. Hap43 was not required for iron acquisition under low-iron conditions. Instead, it was responsible for repression of genes that encode iron-dependent proteins involved in mitochondrial respiration and iron-sulfur cluster assembly. We also demonstrated that Hap43 executes its function by becoming a transcriptional repressor and accumulating in the nucleus in response to iron deprivation. Finally, we found a connection between Hap43 and the global corepressor Tup1 in low-iron-induced flavinogenesis. Taken together, our data suggest a complex interplay among Hap43, Sfu1, and Tup1 to coordinately regulate iron acquisition, iron utilization, and other iron-responsive metabolic activities.

Iron is the fourth most abundant element in the Earth's crust, and the transition states of iron endow it with chemical properties essential to many biological processes. Iron has been confirmed to be crucial for all organisms, with only two exceptions (1, 66). The metal plays a structural or functional role in a variety of proteins responsible for DNA synthesis, respiration, electron transport, oxygen transport/storage, and many central metabolic pathways (18). Excess iron content, however, does lead to deleterious oxidative damage as a result of the Fenton reaction, in which free ferrous iron reacts with H2O2 or lipid peroxide to generate free radicals (28,30). Therefore, a precise regulation system for cellular iron homeostasis is necessary to maintain the intracellular level of iron in a balanced state between the minimal requirement and cytotoxicity.

Based on recent studies of fungal eukaryotes, a finely tuned system for the maintenance of iron homeostasis was identified (48). This system requires apparatuses for iron sensing, iron transportation, and storage. The iron-sensing mechanism can act as an iron-responsive regulator to modulate the activities or expression of downstream effectors and proteins involved in iron uptake/utilization. In other words, a metabolic remodeling event occurs in response to iron repletion or depletion during the process of iron homeostasis (45). When iron levels are low, the iron-responsive activator in some yeast species is activated and induces the expression of genes encoding components for iron acquisition, whereas the expression of genes encoding iron-dependent proteins is repressed. In this way, the limited amount of iron may be used more efficiently by the cells for preventing cellular exhaustion, e.g., the iron can be used by vital enzymes that are involved in DNA synthesis, replication, repair, and transcription (18). In addition, the iron-responsive repressor in most other fungi represses genes involved in iron uptake or iron transport under high-iron conditions. This regulatory strategy avoids the detrimental consequences of iron overload.

In Saccharomyces cerevisiae, iron deficiency leads to activation and cytosol-to-nucleus translocation of two iron-regulatory activators, Aft1 and Aft2 (3, 13, 65, 76, 90). Under iron-deprived conditions, Aft1 and Aft2 directly induce the expression of iron uptake genes to increase the acquisition of external iron sources and withdraw the deposited iron from intracellular vacuoles. Furthermore, the withdrawal of stored iron and the decrease in iron use are also regulated by the gene product of CTH2, which is also one of the Aft1/Aft2 target genes. Cth2 is a CCCH-type zinc finger mRNA-binding protein and can couple with its iron-resistant paralog, Cth1, to bind coordinately to the AU-rich 3′ untranslated region (UTR) of the target mRNA, which leads to its degradation (11, 67, 68). Notably, these target mRNAs are mostly transcribed from genes that encode iron-dependent proteins, and thereby, degradation of these mRNAs decreases the consumption and facilitates the efficient usage of limited amounts of iron. Although Pfam prediction indicates that the Aft-type proteins (PF08731) exist in most hemiascomycota yeasts, only S. cerevisiae and Kluyveromyces lactis Aft orthologs have so far been demonstrated to possess iron-responsive transcriptional activity via the PuCACCC binding motif in the promoter region of the iron regulon (12).

The second regulatory mechanism for the expression of iron-responsive genes is mediated by GATA-type transcriptional repressors, which usually contain one or two Cys2/Cys2-type zinc finger domains separated by a conserved cysteine-rich region (26). The first fungal GATA-type repressor, Urbs1, was identified in the basidiomycete Ustilago maydis (86). Since then, orthologs of Urbs1 in ascomycetes with similar functions have been described, such as Penicillium chrysogenum Srep (25), Neurospora crassa Sre (93), Schizosaccharomyces pombe Fep1 (64), and Pichia pastoris Fep1 (58). Pathogenic fungi also have homologs of Urbs1, including Candida albicans Sfu1 (49), Aspergillus nidulans and Aspergillus fumigatus SreA (27, 60, 81), Histoplasma capsulatum Sre1 (10), and the basidiomycota Cryptococcus neoformans Cir1 (44). These repressors function under iron-replete conditions and negatively regulate the expression of genes encoding iron assimilation proteins, especially those involved in siderophore uptake and other reductive iron uptake apparatuses. Furthermore, loss of the GATA factor Cir1 attenuates the virulence of C. neoformans (44).

Recently, a third mode of transcriptional regulation in response to iron states was demonstrated as a negative modulation under iron-depleted conditions. In S. pombe, php4+ encodes a negative regulatory subunit of the heteromeric CCAAT-binding complex (CBC) composed of Php2/Php3/Php5 (Php2/3/5) (54). The expression of php4+ is partially repressed by the GATA-type repressor Fep1 when iron is sufficient, but it is upregulated in response to iron deficiency. Activated Php4 acts as a transcriptional repressor associated with the constitutively expressed Php2/3/5 complex to repress the gene expression of iron-dependent proteins, especially those involved in the mitochondrial electron transport chain, tricarboxylic acid (TCA) cycle, iron-sulfur (Fe/S) cluster assembly, and heme biosynthesis (56, 57). Moreover, Aspergillus HapX has been identified as a Php4 ortholog by sequence similarity (84). In A. nidulans, HapX is also controlled by SreA; the iron-dependent heme biosynthesis is repressed by HapX, together with the CBC factors HapB/HapC/HapE (HapB/C/E), and the siderophore biosynthesis pathway also requires HapX (34). Moreover, HapX and HapB/C/E are essential for the growth of A. nidulans under iron depletion. Deletion of hapX also leads to significant attenuation of virulence in a mouse model of invasive aspergillosis (79). In C. albicans, previous studies using DNA microarray analysis suggested that the expression of HAP43 (orf19.681), which encodes the ortholog of Php4/HapX, is repressed by Sfu1 under high-iron conditions (49). Experimental evidence demonstrating the roles of C. albicans Hap2 and Hap3 is, however, still lacking. Moreover, C. albicans Hap5 was first revealed as a repressor of mitochondrial electron transport components (43) and was further demonstrated to govern the iron starvation-mediated induction of the gene encoding ferric reductase (Frp1). Hap43 may also partially participate in the Hap-mediated regulation of FRP1 (2). We do not, however, have a detailed understanding of the roles of C. albicans Hap43 in iron-responsive gene regulation and virulence.

In this study, we characterized the function of C. albicans Hap43. We first created a hap43Δ mutant and showed that loss of HAP43 leads to a defect in iron-dependent cell growth. We further hypothesized that a pathogen unable to grow in an iron-deprived environment would lose its virulence to the host because of the iron-withholding defenses within the host body (8) and demonstrated that deletion of HAP43 did indeed attenuate the virulence of C. albicans in a mouse model. Moreover, the impeded growth of the hap43Δ strain under low-iron conditions did not result from its inability to acquire iron but was possibly due to disruptions of iron-responsive transcriptional remodeling in genes encoding iron-dependent proteins. Interestingly, one-hybrid assays revealed that Hap43 becomes a repressor when the environmental iron cannot support the minimal requirement for the growth of C. albicans. This iron-dependent switch in the activity of Hap43 is possibly controlled by low-iron-induced nuclear accumulation of Hap43. Finally, Hap43 was essential for iron starvation-induced flavinogenesis. Taken together, our findings highlight the significance of C. albicans Hap43 in the virulence and transcriptional reprogramming of iron homeostasis, particularly under iron-depleted conditions.


Yeast strains and growth conditions.

All C. albicans strains used in this study are listed in Tables 1 to to3.3. Cells were cultivated in YPD medium (1.0% yeast extract, 2.0% meat peptone, and 2.0% glucose), synthetic complete (SC) medium (0.67% yeast nitrogen base [YNB] with ammonium sulfate, 2.0% glucose, and 0.079% complete supplement mixture), or synthetic minimal (SM) medium (0.67% yeast nitrogen base with ammonium sulfate and 2.0% glucose). Plates were prepared with 1.5% agar for YPD-based medium and 2.0% agar for YNB-based medium. For the selection or growth of certain strains, SMU (SM plus 80 μg/ml uridine) or YPDNou (YPD plus 200 μg/ml nourseothricin; Werner BioAgents, Jena, Germany) medium was used. For the induction of the SAP2 promoter and the MAL2 promoter, yeast carbon base (YCB)-bovine serum albumin (BSA) (23.4 g/liter yeast carbon base, 4 g/liter of BSA fraction V, pH 4.0) medium and YPM (1.0% yeast extract, 2.0% meat peptone, and 2% maltose) medium were used, respectively (72). Cells were cultivated at 30°C with shaking at 180 rpm.

Table 1.
Strains used in gene deletion, iron-related assays, and RT-PCRa
Table 3.
Strains used in confocal microscopy and Western blotting
Table 2.
Strains used in one-hybrid assays

Iron-dependent growth analysis.

Cells were grown in acidic noniron medium (NIM) or YPD-based medium. NIM was prepared as described previously (49). Briefly, NIM contains 0.17% YNB without Fe and Cu, 0.079% complete supplement mixture, 2% glucose, 0.5% ammonium sulfate, and 0.25 μM CuSO4 and is supplemented with 100 μM basophenanthrolinedisulfonate disodium salt (BPS; Sigma), a ferrous iron chelator, to restrict the iron content of the medium. To support minimal cell growth, 10 μM ferrous ammonium sulfate (FAS) was added to make the low-iron medium (LIM). Accordingly, high-iron medium (HIM) was prepared by adding 10-fold excess of FAS (100 μM). The second limited-iron medium was prepared by adding BPS to the YPD medium to restrain the free iron. YPD was defined as an iron-rich medium, whereas YPD plus 200 μM BPS was defined as an iron-poor medium. For pre-iron starvation, overnight cultures in YPD medium were diluted 100- to 1,000-fold into NIM or YPD plus 400 μM BPS and then grown at 30°C with shaking for 20 to 24 h to achieve a steady state. For spot assays, iron-starved cells were harvested by centrifugation (Eppendorf 5810R centrifuge; A-4-62 rotor; 1,500 × g; 5 min; 25°C) and serially diluted to the desired cell densities with sterile double-distilled water (ddH2O). Each dilution was spotted onto agar plates (5 μl/spot) and incubated at 30°C for 1 day or longer as indicated.

Gene manipulation.

All deletion strains were generated from SC5314 using the SAT1 flipper method (72). The primers used are listed in Table 4. The HAP43 deletion cassette was constructed as follows. An ApaI-XhoI DNA fragment composed of the 5′ flanking region (nucleotides −816 to −64) of HAP43 was amplified from the SC5314 genome with the primer pair Orf19.681UP-1-ApaI and Orf19.681UP-2-XhoI. A SacII-SacI fragment composed of the 3′ flanking region (nucleotides +1949 to +2228) of HAP43 was amplified from the SC5314 genome with the primer pair Orf19.681DOWN-1-SacII and Orf19.681DOWN-2-SacI. These HAP43 5′ and 3′ flanking regions were cloned into the pSFS1A vector in order at the indicated restriction sites to generate pSFS1A-35fCaHap43. The plasmid pSFS2A-35fCaHap43 was constructed by replacing the XhoI-SacII fragment in pSFS1A-35fCaHap43 with the SAT1-PMAL2-FLIP cassette from pSFS2A. For the construction of the DNA fragment used in the HAP43 reintegration, an ApaI-XhoI fragment composed of the HAP43 promoter together with the full-length HAP43 coding sequence was amplified from the SC5314 genome with the primer pair Orf19.681UP-1-ApaI and CaHap43-ORF-2-XhoI. This fragment was then used to replace the 5′ flanking sequence of pSFS2A-35fCaHap43 to generate pSFS2A-35fCaHap43-ORF.

Table 4.
Primers and oligonucleotides used in strain constructions

C. albicans strains were transformed by electroporation as described previously (72) with some modifications. Cells from 5 ml of overnight culture in YPD medium were collected by centrifugation (Eppendorf 5810R centrifuge; A-4-62 rotor; 1,500 × g; 5 min; 25°C) and resuspended in sterile lithium buffer (8 ml of ddH2O, 1 ml of 10× Tris-EDTA [TE] [pH 8.0], and 1 ml of 1.0 M lithium acetate [pH 7.5]). After incubation at 30°C for 1 h with shaking, 250 μl of 1.0 M dithiothreitol (DTT) was added to the cell suspension and incubated for 30 min at 30°C with shaking. Then, the cells were washed sequentially with 30 ml of ddH2O, 10 ml of ice-cold ddH2O, and 5 ml of ice-cold 1.0 M sorbitol; resuspended in 500 μl 1.0 M sorbitol; and kept on ice until it was used. Deletion and reintegration cassettes from pSFS1A-35fCaHap43 (or pSFS2A-35fCaHap43) and pSFS2A-35fCaHap43-ORF, respectively, were excised by ApaI/SacI digestion and purified. Approximately 1 to 2 μg of linear DNA fragments was mixed with 50 μl of electrocompetent cells, and the cells were transformed with the Gene Pulser Xcell electroporator (Bio-Rad; 1,800 V; 200 Ω; 25 μF). After electroporation, the cells were immediately washed with 1 ml of 1.0 M sorbitol, resuspended in 1 ml fresh YPD medium, and incubated for 3 to 4 h at 30°C with shaking. For the selection of cells with the genome-integrating cassette, the transformants were spread onto YPDNou agar plates and grown at 30°C for 1 to 2 days. Nou-resistant colonies were selected for verification by PCR. To pop out the integrated SAT1-FLIP cassette from the HAP43 locus, the MAL2 promoter or SAP2 promoter was induced by YPM (22) or YCB-BSA (38) medium, respectively. Briefly, cells were inoculated into 5 ml YPM or YCB-BSA medium and grown for 20 to 24 h at 30°C with shaking. Overnight-cultured cells were then plated on YPM or YCB-BSA agar plates and incubated at 30°C for 2 days. Single colonies were then screened for Nou resistance by streaking them on both YPD and YPDNou plates, and the Nou-sensitive strains were picked for PCR diagnostics. Two rounds of deletion cassette integration and excision were performed to manipulate two loci of HAP43 during the process of HAP43 deletion or HAP43 reconstitution. Homozygous hap43Δ strains were created from SC5314, whereas HAP43 reconstituted strains were created from one hap43Δ strain (Table 1).

Similar procedures were used for the deletion of SFU1. Briefly, an ApaI-XhoI fragment corresponding to the 5′ flanking region (nucleotides −927 to −83) of C. albicans SFU1 was amplified from the SC5314 genome with the primer pair CaSfu1up-1-ApaI and CaSfu1up-2-XhoI, whereas a SacII-SacI fragment corresponding to the 3′ flanking region (nucleotides +1555 to +2042) was amplified from the SC5314 genome with the primer pair CaSfu1down-1-SacII and CaSfu1down-2-SacI. The SFU1 5′ and 3′ flanking regions were cloned into pSFS1A to generate pSFS1A-35fCaSfu1. For the construction of the DNA fragment used in the SFU1 reintegration, an ApaI-XhoI fragment composed of the SFU1 promoter region together with the full-length SFU1 coding sequence was amplified from the SC5314 genome with the primer pair CaSfu1up-1-ApaI and CaSfu1-ORF-2-XhoI to replace the 5′ flanking sequence of pSFS1A-35fCaSfu1, generating pSFS2A-35fCaSfu1-ORF. These ApaI-SacI fragments were excised from pSFS1A-35fCaSfu1 and pSFS2A-35fCaSfu1-ORF and used in yeast transformation to generate SFU1 homozygous deletion strains and reconstituted strains, respectively, as described above. All strains were verified by diagnostic PCR. For a successful deletion, the intra-open reading frame (ORF) PCR yielded no product, whereas a parental SC5314 yielded a matched product.

Southern blotting.

Genomic DNA isolation for C. albicans was performed as described previously (72) with modifications. Cells from 5 ml of overnight cultures in YPD medium were spun down, resuspended in 200 μl breaking buffer (10 mM Tris-Cl [pH 7.5], 100 mM NaCl, 1 mM EDTA, 2% Triton X-100, 1% SDS), and transferred to a microcentrifuge tube containing 0.3 g glass beads (Sigma) and 200 μl of PCIA (phenol-chloroform–isoamyl alcohol [25:24:1], 0.1% 8-quinolinol, pH 7.0). The cells were lysed by vortexing them at maximum speed for 5 min. The lysates were mixed with 200 μl TE buffer (pH 8.0) and centrifuged at maximum speed for 5 min (Eppendorf 5413D centrifuge; F45-24-11 rotor; 25°C). DNA in the aqueous phase was precipitated with 95% ethanol, pelleted by centrifugation (Eppendorf 5413D centrifuge; F45-24-11 rotor; maximum speed; 5 min; 25°C), dissolved in 400 μl TE buffer (pH 8.0) containing 3.0 μl of 10-mg/ml RNase A, and incubated at 37°C for 5 min. The RNase-treated genomic DNA was precipitated with 95% ethanol, pelleted by centrifugation (Eppendorf 5413D centrifuge; F45-24-11 rotor; maximum speed; 5 min; 25°C), and dissolved in 100 μl ddH2O. Blotting was performed using a standard method with some modifications (77). Approximately 20 μg of genomic DNA was digested with XbaI overnight at 37°C, separated on a 0.8% agarose gel with 1:10,000 SYBR Safe gel stain dye (Invitrogen), transferred onto a positively charged nylon membrane (Pall Corporation) by the alkaline transfer method, and fixed by baking it at 80°C for 2 h. The membrane was hybridized with [α-32P]dCTP-labeled HAP43 5′ flanking fragments using prehybridization buffer (6× SSC [1× SSC is 0.15 M NaCl plus 0.015 M sodium citrate], 5× Denhardt's reagent, 0.5% SDS, 100 μg/ml salmon sperm DNA, and 50% formamide) at 42°C overnight. The membrane was then washed and visualized by autoradiography.

Virulence assay.

Forty female BALB/c mice (7 weeks old) were obtained from BioLasco Taiwan Co., Ltd., and group housed (5 mice per cage) for 1 week before experiments. The C. albicans cells for inoculations were grown in SC medium overnight at 30°C and then subcultivated in SC medium to the early log phase. Yeast cells were harvested by centrifugation (Eppendorf 5810R centrifuge; A-4-62 rotor; 1,500 × g; 5 min; 25°C), washed once with phosphate-buffered saline (PBS), and resuspended in PBS at a density of 5.0 × 107 cells/ml. The cell suspension (100 μl; final, 5.0 × 106 cells) was injected via the lateral tail vein. Ten mice were used for each C. albicans strain. The injected mice were monitored for clinical symptoms of distress and for survival twice per day for 2 weeks. The animal studies were approved by the Institutional Animal Care and Use Committees, National Tsing Hua University and Animal Technology Institute Taiwan, Taiwan. The log rank test was used to assess the differences in survival between groups of mice. A P value of <0.05 was considered statistically significant.

For kidney sections, organs extracted from the mice on the third day postinfection were formaldehyde fixed and embedded in paraffin. Tissue sections were stained with periodic acid Schiff (PAS) to visualize C. albicans cells.

One-hybrid analysis.

Plasmids and strains used in one-hybrid assays were created as described previously (75) with some modifications. BamHI-linearized pCR-lacZ and pCR-OPlacZ were integrated into CAI8 at the ade2::hisG locus to generate strains CCR1 and COP1, respectively. The S. aureus LexA operator linked with penta-Gcn4 binding sequences [SaLexA-(GCRE)5] was introduced on a double-stranded oligonucleotide (Table 4) between the PstI and SalI sites in the pCR-lacZ vector to generate pCR-OPGCRElacZ. BamHI-linearized pCR-OPGCRElacZ was integrated into the CAI8 genome to create the COPGCRE-5 strain. All C. albicans strains carrying integrated pCR-lacZ and its derivatives were selected for the plasmid-borne ADE2 marker in SMU medium.

To fuse the lexA sequence of pCIplexA (75) in frame, a HindIII-MluI fragment composed of the LexA coding sequence was amplified from pCIplexA with the primer pair SaLexA-HindIII-1 and SaLexA-MluI-2. This fragment contains a primer-introduced GGT sequence at the C terminus and was cloned into pCIplexA between HindIII and MluI sites to generate pCIplexA-F1. In contrast to pCIplexA, the linker sequence 5′-GGTCC-3′ located between the LexA coding sequence and the MluI site was changed to 5′-GGT-3′ in pCIplexA-F1. The HAP43, GCN4, and NRG1 ORFs were amplified by PCR (Table 4) from the SC5314 genome and subcloned into pCIplexA-F1 between the MluI and PstI sites to create pCIplexA-F-CaHap43-3, pCIplexA-F-CaGcn4-21, and pCIplexA-F-CaNrg1-18, respectively. Strains CC1, CG21, CN18, and CH43-3 were generated by transforming CCR1 with StuI-linearized pCIplexA-F1, pCIplexA-F-CaGcn4-21, pCIplexA-F-CaNrg1-18, and pCIplexA-F-CaHap43-3, respectively. Strains OC1, OG21, ON18, and OH43-3 were generated by transforming COP1 with the four plasmids mentioned above. In addition, the OGRH43 and COGRE strains were generated by transforming COPGCRE-5 with pCIplexA-F-CaHap43-3 and pCIplexA-F1, respectively. Integration of pCIplexA-F1 and its derivatives into the C. albicans genome at the RPS1 locus was selected by the plasmid-borne URA3 marker using SM medium. Integrations were confirmed by PCR diagnostics (Table 4).

The expression level of the β-galactosidase reporter in the one-hybrid strains was assayed by X-Gal (5-bromo-4-chloro-3-indolyl-β-d-galactopyranoside) overlay assays and liquid β-galactosidase (β-Gal) assays as described previously (74) with some modifications. For overlay assays, overnight-grown colonies on agar plates were lysed with chloroform for 5 min, air dried for 10 min after the chloroform was decanted, and overlaid with X-Gal–agarose (1% agarose, 0.25 or 0.5 mg/ml X-Gal, 0.1 M sodium phosphate buffer [pH 7.0]). After the gel solidified, the plates were incubated at 30 or 37°C overnight until the blue color developed. For liquid assays, cells at an optical density at 600 nm (OD600) of ~20 were resuspended in 300 μl Z buffer with β-mercaptoethanol (β-ME). Cells in 100 μl of the suspension were lysed by adding 15 μl of 0.1% SDS and 30 μl chloroform and vortexing them for 15 s. Then, the suspensions of lysed cells were incubated with 200 μl of 4-mg/ml ONPG (o-nitrophenyl-β-d-galactopyranoside) (Sigma) in potassium phosphate buffer (pH 7.0) at 37°C for 30 min. The reaction was stopped by adding 0.5 ml of 1.0 M Na2CO3(aq). The cell debris was spun down, and the absorbance of supernatants at 420 and 550 nm was determined with a spectrophotometer. The β-galactosidase levels were displayed as Miller units from at least three independent experiments.

Fluorescence microscopy.

To observe the translocation of green fluorescent protein (GFP)-tagged Hap43 in response to external iron levels, the S. aureus lexA sequence on pCIplexA-F-CaHap43-3 was replaced with the PCR product of the Candida-adapted GFP sequence from pNIM1 (62) to generate pCIpGFP-F-CaHap43-1. To generate strain CGH43-2, the StuI-linearized plasmid was integrated into the RPS1 locus of CAI4. Integrations were confirmed by PCR using primers listed in Table 4. The expression of GFP-HAP43 in this system was driven by the ACT1 promoter, which is constitutively active independently of the iron levels (49, 71). For the induction of GFP-Hap43 nuclear translocation, an iron-depleted culture (YPD plus 400 μM BPS) was prepared at a seeding density of 0.5 OD600/ml, and the cells were grown to mid-log phase (~5 OD600/ml) at 30°C with shaking. The cells were fixed with 3% formaldehyde in the culture medium at room temperature for 2 h with shaking, washed, and resuspended in PBS. The fixed cells were subsequently stained with DAPI (4′,6-diamidino-2-phenylindole) (Sigma; 10 μg/ml in PBS) at room temperature for 10 min. The stained cells were mounted on glass slides coated with 1% agarose, covered with a coverslip, and examined under a Carl Zeiss LSM 510 confocal microscope equipped with HeNe, argon visible-light, and diode lasers. GFP-Hap43 was excited by the argon laser at 488 nm, and DAPI was excited at 405 nm.

RNA isolation and quantitative real-time PCR.

Total RNA was extracted by the phenol-chloroform method. Briefly, mid-log-phase cells from subcultures grown under appropriate conditions were harvested by centrifugation (Eppendorf 5810R centrifuge; A-4-62 rotor; 3,000 × g; 5 min; 25°C), washed once with ddH2O, and stored at −80°C. Cells (OD600, ~20) were thawed on ice and resuspended in 500 μl ice-cold lysis buffer (0.1 M Tris-Cl [pH 7.5], 0.1 M LiCl, 2% β-ME, 0.01 M EDTA, and 5% SDS). The mixtures were transferred to a microcentrifuge tube containing 500 μl PCIA (pH 4.5) and 0.3 g glass beads (Sigma). The cells were lysed by vortexing them at maximum speed for 5 min. Supernatants were obtained by centrifugation (Eppendorf 5413D centrifuge; F45-24-11 rotor; maximum speed; 5 min; 4°C), extracted two times with 200 μl PCIA, and transferred into 1 ml of 99% ethanol for precipitation. Nucleic acid pellets were spun down, washed with 70% ethanol, air dried, and then resuspended in 100 μl nuclease-free water (Ambion). DNA contaminants were removed with a Turbo DNase-free kit (Ambion). The overall quality of the RNA was analyzed by agarose gel electrophoresis.

For cDNA synthesis, 4.0 μg pure RNA was reverse transcribed using 400 U Moloney murine leukemia virus (MMLV) reverse transcriptase (Promega) with 1.0 μg oligo(dT)18 primer in a 50-μl reaction mixture according to the manufacturer's instructions. Quantitative real-time PCR was carried out with the 7500 real-time PCR system (Applied Biosystems). The primers used are listed in Table 5. Briefly, each 20-μl reaction mixture contained 80 ng cDNA, 10 μl Power SYBR green PCR master mixture (Applied Biosystems), and 300 nM each forward and reverse primer. The reactions were performed with 1 cycle at 95°C for 10 min, followed by 40 repeated cycles at 95°C for 15 s and 60°C for 1 min. The EFB1 transcripts were used as an internal control for RNA input and quality after reverse transcription (78), and the ACT1 transcripts were used as an endogenous control for the quantitative PCR.

Table 5.
Primers used in RT-PCR and quantitative real-time PCR

Protein extraction and immunoblotting.

The protein extraction protocol was adapted from the instructions for the EasySelect Pichia Expression Kit (Invitrogen). Briefly, cells at an OD600 of about 20 to 30 were harvested by centrifugation (Eppendorf 5810R centrifuge; A-4-62 rotor; 3,000 × g; 5 min; 25°C), washed once with ddH2O, and stored at −80°C before lysis. Cell pellets were resuspended in 300 μl ice-cold breaking buffer (1 mM EDTA, 5% glycerol, 1 mM phenylmethylsulfonyl fluoride [PMSF], 1/1,000-diluted protease inhibitor cocktail [Sigma], 50 mM monobasic sodium phosphate [pH 7.4]) plus 0.3 g glass beads (Sigma). The cells were lysed by vortexing them at maximum speed for 5 min and then chilled on ice for another 5 min. The supernatants were separated from the beads and cell debris by centrifugation (Eppendorf 5413D centrifuge; F45-24-11 rotor; maximum speed; 5 min; 4°C) and stored at −80°C. Whole-cell extracts were quantified with the Pierce BCA Protein Assay Kit.

Equal amounts of whole-cell extract from each sample were subjected to 8% SDS-PAGE and subsequently transferred to a polyvinylidene difluoride (PVDF) membrane (Pall Corporation) with an enhanced chemiluminescence (ECL) semidry blotter (TE 77; Amersham). The membranes were blocked with 3% nonfat milk in TBST (TBS with 0.1% Tween 20) at room temperature for 2 h. For immunodetection, mouse monoclonal anti-GFP IgG (632380 [Clontech] or NB600-601SS [Novus Biologicals, Inc.]) diluted 1:2,000 in TBST (0.1% Tween 20, 1% BSA) was used. After hybridization with primary antibodies, the membranes were washed with TBST three times (25°C; 5 min each) and incubated with horseradish peroxidase (HRP)-conjugated goat anti-mouse or anti-rabbit IgG (Santa Cruz Biotechnology). Chemiluminescence was developed with ECL reagents (Perkin Elmer) and visualized by exposure to X-ray film.

Colorimetric quantification assay for determination of intracellular iron contents.

Intracellular iron levels of C. albicans were quantified by the BPS-based colorimetric method (83) with some modifications. Cells were harvested by centrifugation (Eppendorf 5810R centrifuge; A-4-62 rotor; 3,000 × g; 5 min; 25°C), washed with ddH2O, and resuspended in 500 μl of 3% nitric acid (J. T. Baker). The cell suspensions were boiled for 2 h to digest the cells completely, and cell debris was removed by centrifugation (Eppendorf 5413D centrifuge; F45-24-11 rotor; maximum speed; 5 min; 25°C). The supernatants containing iron (400 μl each) were collected and mixed with 160 μl of 38-mg/ml sodium ascorbate (Sigma), 320 μl of 1.7-mg/ml BPS, and 126 μl of 4 M ammonium acetate. Chelating reaction mixtures were incubated at room temperature for 5 min. The OD535 of the BPS-Fe complex was recorded with a spectrophotometer against blanks containing all reagents except the cells. To eliminate the nonspecific absorbance, the OD680 was subtracted from the OD535. The values for the iron content were adjusted by normalization according to the number of digested cells. Cell numbers were indicated by OD600 using the following formula: (OD535 − OD680)/(OD600).

In the 30-min iron uptake assay, the iron uptake activities of C. albicans strains were evaluated by measuring the increase in intracellular iron content after 30 min of iron uptake. Briefly, for pre-iron starvation, C. albicans cells were grown in YPD medium plus 400 μM BPS for at least 24 h, while the cells without the pre-iron starvation treatment were grown in YPD medium without BPS. These stationary-phase cells were harvested (Eppendorf 5810R centrifuge; A-4-62 rotor; 1,500 × g; 5 min; 25°C), washed with ddH2O, and resuspended in fresh YPD medium at a cell density of 2.0 OD600/ml. At time zero (T0), cells at an OD600 of ~10 were harvested immediately, washed with ddH2O, and stored at −80°C until they were used. Uptake reactions were then facilitated at 30°C for 30 min with shaking, and cells at T30 were harvested as described above. Intracellular iron was measured by the colorimetric method and displayed in arbitrary units (AU). The iron uptake activity of each C. albicans strain was determined by the difference between the intracellular iron levels at T0 (AU0 min) and T30 (AU30 min). Relative increases in intracellular iron within 30 min (Δ30–0 min) were calculated by subtracting the average AU0 min from the average AU30 min. The percentage relative to the wild type without iron starvation (% to ΔWT) was shown as indicated.

In the growth-dependent iron uptake assay, cells with or without pre-iron starvation treatment were inoculated at a cell density of 0.5 OD600/ml into fresh YPD (high-iron) medium or YPD plus 200 μM BPS (low-iron) medium. All inoculated cultures were grown at 30°C with shaking for 5 h to allow 3 or 4 generations of cell growth. After the incubation, cells were harvested, and the intracellular iron was measured.

2,3,5-Triphenyltetrazolium chloride (TTC) reduction overlay assay.

To evaluate cell surface reductase activity, the overlay assay with TTC (Sigma) was performed as described previously (44) with some modifications. Briefly, YPD-grown cells were harvested and diluted with sterile ddH2O to a cell density of 1.0 OD600/ml. Each dilution was spotted onto YPD agar plates (10 μl/spot) and incubated at 30°C for 1 day. Then, the colonies were overlaid with agarose (1.5% in PBS) containing 0.1% TTC. The TTC reduction reactions were performed at 30°C, and the development of red color on the colony spots was photographed. To inhibit electron transfer from respiration during the TTC reduction, 20 μg/ml antimycin A (Sigma) was added to the agarose overlay.

Phleomycin sensitivity assay.

Phleomycin sensitivity was assessed by spotting 10-fold serial dilutions of cells (5 μl/spot) onto YPD agar plates containing 50 μg/ml phleomycin (Zeocin; Invitrogen), with incubation at 30°C for 1 day. To restrict the growth-inhibitory effect of phleomycin on C. albicans, 100 μM BPS was added to the plates.

Flavin secretion assay.

For the induction of flavin secretion, C. albicans cells were iron starved in NIM at 30°C with shaking for 2 days. Supernatants from stationary-phase cultures were collected by centrifugation (Eppendorf 5413D centrifuge; F45-24-11 rotor; maximum speed; 5 min; 25°C). Two peaks at 360 nm and 450 nm can be observed in the flavin UV-visible spectra (53). The flavin content of each supernatant was quantified at the absorption maximum of 446.3 nm (47). The supernatants were photographed under UV light. Data were collected from at least three independent experiments.


C. albicans Hap43 contains a region that is highly conserved in the N termini of fungal HapX/Php4.

The 634-amino-acid protein encoded by C. albicans HAP43 was first reported to be similar to the AP1-like transcription factor (49). We performed BLAST searches using the Candida Genome Database (CGD) (http://www.candidagenome.org) and identified HAP43 as the homolog of Hap4/Php4/HapX. To elucidate possible functional domains conserved in Hap43, we compared the Hap43 protein sequence with those of Aspergillus and N. crassa HapX, S. pombe Php4, and S. cerevisiae Hap4 and Yap5. Notably, we calculated the pairwise identities and similarities with the MatGat program (9) and found that the overall similarities among the proteins were weak. Hap43 and A. nidulans HapX share 28.7% identity (42.3% similarity), whereas Hap43 and Php4 share only 17.2% identity (27.6% similarity). Nevertheless, a highly conserved region was revealed in the N termini of these proteins. In this region, Hap43 and A. nidulans HapX share 42.1% identity (61.7% similarity), whereas Hap43 and Php4 share 26.1% identity (47.5% similarity). Figure 1 A shows alignments of these proteins with residues 1 to 147 of Hap43 and highlights the two domains that are particularly well conserved in these N-terminal regions. The first domain is the putative Hap complex-interacting domain that is required for interaction of Hap4 with the CBC in yeast (5). Interestingly, all proteins but Yap5 possess this domain, implying the presence of a distinct molecular mechanism between the typical AP1-like bZip factor (i.e., Yap5) and CCAAT-related transcription factors. The second domain within the conserved N-terminal region is the basic region of the bZip domain. With the exception of S. cerevisiae Hap4, this region exists in all the compared proteins (Fig. 1A). Notably, all HapX homologs and C. albicans Hap43 possess both the putative Hap complex-interacting domain and the basic region of the bZip structure, which suggests that Hap43 may have a conserved function similar to that of HapX/Php4. In addition, a similar comparison between Aspergillus HapX and C. albicans Hap43 also highlighted three conserved cysteine-rich motifs in the central-to-C-terminal region (34).

Fig. 1.
Construction of hap43-null mutants. (A) The amino acid sequence of C. albicans Hap43 was aligned with those of Aspergillus species HapX, N. crassa HapX, S. cerevisiae Yap5, S. pombe Php4, and S. cerevisiae Hap4 using Clustal W (50). The highly conserved ...

The hap43Δ strain is unable to grow under low-iron conditions.

To determine whether C. albicans Hap43 plays a role in iron homeostasis, we deleted HAP43 using the SAT1 flipper method (Fig. 1B) (72). Successful construction of the hap43-null mutants and their reintegrated strains was verified by Southern blot analysis (Fig. 1C). The cell growth of isogenic mutants was compared with that of the parental wild-type SC5314 under low-iron conditions. Based on the previous investigation, a synthetic YNB-based medium containing 100 μM BPS, an iron chelator, was used as the NIM, and ferrous iron was added to support the growth of C. albicans. The medium supplied with 10 μM ferrous iron was used as low-iron conditions, whereas supplementation with 100 μM ferrous iron generated high-iron conditions (49). In contrast to the wild-type and heterozygous strains, hap43 homozygous deletion mutants under low-iron conditions (NIM plus 10 μM Fe2+) were severely defective in their growth, even though the incubation was continued for 3 days (Fig. 2 A, top) or longer (data not shown). This growth defect was rescued by supplying excess iron (100 μM Fe2+) (Fig. 2A, bottom).

Fig. 2.
Deletion of C. albicans HAP43 causes growth defects under low-iron conditions. (A) Cells of C. albicans wild-type, hap43 heterozygous and homozygous deletion, and HAP43 reconstituted strains were serially diluted and spotted onto NIM-based iron agar plates ...

Baek et al. have demonstrated that hap43Δ mutants cannot grow on iron-limiting agar plates using a nonfermentable carbon source (YPG, composed of 1% yeast extract, 2% peptone, and 3% glycerol) and 150 μM BPS (2). We tested the growth of hap43Δ mutants under similar conditions, but YPD (with glucose as the carbon source) was used instead of YPG. To restrict the free iron, cells were spotted onto YPD agar plates with 25, 50, 100, 200, 400, or 800 μM BPS. The defective growth of hap43 mutants on iron-depleted YPD agar plates was quite similar to that under the NIM-based conditions (Fig. 2B). Iron restriction with <100 μM BPS was unable to inhibit the growth of the tested strains. With 200 μM BPS, the wild-type strain displayed reduced growth, whereas the growth of the hap43Δ strain was severely impeded. On the plates containing >400 μM BPS, the growth of all strains was severely defective in spite of a longer incubation of 3 days (data not shown). These findings allowed us to define YPD (without BPS), YPD plus 200 μM BPS, and YPD plus 400 μM BPS as high-iron, low-iron, and iron-starved conditions, respectively. The growth defect of the hap43Δ strain was successfully rescued by native expression of HAP43 in the reconstituted strain (Fig. 2A and B). Taken together, we concluded that Hap43 is essential for the growth of C. albicans under either acidic (NIM-based; pH 4.2) or near-neutral (YPD-based; pH 6.5) low-iron conditions. Moreover, we also found that YPD-based iron-limiting medium (YPD plus 400 μM BPS) can support the minimal growth of the hap43Δ strain in liquid culture during the early to mid-log phase but is unable to sustain the growth of this mutant strain to the stationary phase (data not shown). Thus, we could easily collect sufficient hap43Δ cells at the mid-log phase in YPD-based iron-limited culture without the prolonged incubation that is needed for synthetic medium (NIM based) (data not shown). As a result, YPD-based conditions were used in most of our subsequent experiments for convenience.

Loss of HAP43 impedes virulence in a mouse model of disseminated candidiasis.

To resist microbial infections, the human body develops natural resistance by storing iron and maintaining an extremely low level of free ionic iron (1018 M) in tissue fluids (8). This tightly regulated low-iron environment not only supports the normal function of the immune system, but also inhibits the growth of invading or resident pathogens. Because Hap43 is essential for C. albicans growth under iron-restricted conditions, we speculated that Hap43 might play a role in the virulence of C. albicans. We tested this hypothesis in a mouse dissemination model of candidiasis (Fig. 3 A). After the intravenous injection of the tails of mice with 5 × 106 yeast cells in PBS, nearly all mice displayed symptoms of illness, such as ruffled hair, shivering, and lethargy, on the third day postinfection (data not shown). The wild-type and the hap43 heterozygous strains exhibited the highest virulence, leading to the rapid death of the mice beginning on the third and fourth days postinfection, and all mice in these two groups died within 8 days. In contrast, mice infected with the hap43 homozygous mutant began to die on the seventh day, and 60% of the mice remained alive 2 weeks postinfection. Moreover, all surviving mice recovered from the illness by the 15th day postinfection (data not shown). In addition, all mice that had been infected with the HAP43 reconstituted strain succumbed to infection within 13 days. Although the death rate of mice injected with the reconstituted strain was lower than that of mice injected with wild-type cells (P = 0.00527), the reintegration of HAP43 did restore the attenuated virulence of the hap43-null mutant (P = 1.25E−05) to a level that was not significantly different from the heterozygous (hap43Δ/HAP43) mutant (P = 0.0581).

Fig. 3.
Deletion of HAP43 attenuates C. albicans virulence. (A) Ten female BALB/c mice were injected via the tail vein with 5 × 106 C. albicans cells, including SC5314 (orange), a heterozygous hap43 deletion mutant (black), a homozygous hap43 deletion ...

Kidneys were also collected on the third day postinfection from both wild-type-infected and hap43Δ/hap43Δ strain-infected mice for histological examination. In the kidney sections examined, the wild-type cells were observed as filamentous structures (Fig. 3C and D). In contrast, no cells from the hap43-null mutant strain could be visualized (Fig. 3B), implying that there is a slow invasion of the mutant cells or that they are eliminated from the host tissues. These results support our hypothesis that Hap43 is essential for growth under iron-restricted conditions, such as those within the host, and contributes to C. albicans virulence.

The hap43 deletion mutant is not defective in iron acquisition.

To understand the nature of growth defects in the hap43Δ mutant under iron-restricted conditions, two possible hypotheses were considered. One was that C. albicans is unable to acquire sufficient iron from extracellular environments in the absence of Hap43. The other was that an irregular distribution of iron occurs within the cell when HAP43 is deleted. To distinguish these two possibilities, we used a 30-min iron uptake assay to quantify the net increase in intracellular iron in the hap43Δ mutant. Wild-type, sfu1Δ, and ftr1Δ strains were used as controls, as iron uptake genes are derepressed in the sfu1Δ mutant under high-iron conditions (49) and the ftr1Δ mutant is defective in iron uptake (71). For the cells without pre-iron starvation (i.e., cells grown in YPD plus 400 μM BPS for 24 h), both sfu1Δ and hap43Δ mutants exhibited a 40 to 50% increase in iron uptake in fresh YPD medium compared with the wild type, whereas the ftr1Δ mutant showed extremely low iron uptake activity (Fig. 4 A). Moreover, although iron starvation enhanced the iron uptake activity in all strains, including the ftr1Δ mutant, there were no significant differences in intracellular iron among iron-starved hap43Δ, sfu1Δ, and wild-type strains. This result implies that the hap43Δ mutant possesses a functional iron acquisition activity even when cells have been iron starved compared with the wild type.

Fig. 4.
The hap43Δ strain is not defective in iron acquisition under iron-depleted conditions. (A) The iron uptake activities of C. albicans were evaluated in a 30-min uptake assay in fresh YPD medium. Normal and iron-starved stationary-phase cells were ...

The sfu1Δ mutant exhibits an iron overload phenotype under high-iron conditions, and this is possibly due to its constitutive expression of iron uptake-related genes, including cell surface ferric reductases (49). Elevated ferric reductase activity as determined by the TTC reduction assay is also demonstrated in S. pombe fep1Δ (64) and C. neoformans cir1Δ (44) mutants. Therefore, to further study the iron uptake-related activity, we performed the TTC reduction assay to determine the ferric iron reduction activities of C. albicans strains. The sfu1Δ strain displayed a high level of cell surface reductase activity, as indicated by the dark-red formazan precipitate that formed on top of the colony spot (Fig. 4B). In contrast, the hap43Δ and ftr1Δ mutants showed no obvious differences in colony color compared with the wild type, indicating that high intracellular iron in the hap43Δ strain is not caused by elevated activity of ferric reductases. The contribution of ferric reductases in the TTC reduction was further confirmed by the addition of a respiration inhibitor, antimycin A, which eliminates the effect of mitochondrial electron transport. Antimycin A did not affect the pattern of formazan formation (Fig. 4B). This result is consistent with the previous finding that C. albicans ferric reductase activity is independent of mitochondrial respiration (46).

We also measured the intracellular iron levels of all strains using a growth-dependent iron uptake assay, as described in Materials and Methods. After cell growth in the high-iron medium for several generations, both the hap43Δ and sfu1Δ mutants accumulated higher levels of intracellular iron than did the wild type (Fig. 4C, gray bar). When cells were grown in the low-iron medium, however, they exhibited no significant differences from the wild type in the level of intracellular iron. In addition, cells without and with pre-iron starvation showed the same result. Although the iron uptake activity of the ftr1Δ mutant was quite low in the 30-min assay (Fig. 4A), the ftr1Δ strain gained a normal iron content compared with the wild type (Fig. 4C), unless it was pre-iron starved followed by growth in the low-iron medium. This is because Ftr1 is essential only under low-iron conditions, and the ftr1Δ strain can grow well under iron-sufficient conditions (71). Therefore, the ftr1Δ strain was omitted from the assay under iron-starved conditions (i.e., cells were pre-iron starved and then grown in the low-iron medium) because it did not grow after 5 h of incubation in this assay.

To further determine the intracellular iron contents, tests of cell sensitivity to phleomycin were also performed. Phleomycin belongs to an antibiotic family that requires iron to generate free radicals. Sensitivity to phleomycin reflects the level of free intracellular iron (44, 64). The accumulation of excess free iron within the sfu1Δ mutant led to its hypersensitivity to phleomycin, and this phenotype was alleviated by iron restriction with BPS (Fig. 4D). The hap43Δ strain, however, was not hypersensitive to phleomycin compared with the sfu1Δ strain, suggesting that excess iron within hap43Δ cells may not exist as a freely accessible form to react with phleomycin to cause cytotoxicity. Taken together, these studies not only substantiate the role of C. albicans Hap43 in iron homeostasis, but also exclude the possibility that the growth defect of the hap43Δ mutant under iron-depleted conditions (Fig. 2) is due to impaired iron acquisition.

Hap43 is responsible for the repression of iron utilization genes under iron-deprived conditions.

To test the second possibility mentioned above, we performed quantitative gene expression analyses to determine whether Hap43 can function as a regulator of the expression of genes involved in iron homeostasis, especially when the extracellular iron levels are disturbed. S. cerevisiae utilizes multiple transcriptional and posttranscriptional mechanisms to optimize iron usage in response to iron deprivation (45) by inducing the expression of genes encoding iron acquisition apparatuses and of Cth1/Cth2, two proteins required for mRNA degradation. Cth1 and Cth2 are responsible for the downregulation of genes encoding iron-dependent proteins that engage in metabolic pathways, such as respiration, the TCA cycle, lipid synthesis, heme synthesis, biotin synthesis, and Fe/S cluster assembly. Similar iron-dependent metabolic remodeling occurs in S. pombe and A. nidulans (34, 57), and key negative regulators that participate in this process are Php4 and HapX, respectively. The existence of a similar remodeling system and the possible role of Hap43 in this iron-responsive regulatory mechanism have not, however, been investigated in C. albicans.

Accordingly, we selected several genes that are differentially expressed in response to iron limitation based on genome-wide microarray data (49) and examined their expression in the hap43 deletion background. We first confirmed that expression of HAP43 is upregulated in the sfu1Δ mutant under high-iron conditions (49) and that expression of SFU1 is also upregulated in the hap43Δ mutant independently of iron levels (data not shown); this suggests that there is reciprocal transcriptional regulation between the expression of HAP43 and that of SFU1. Interestingly, expression of HAP43 was higher under low-iron conditions than under high-iron conditions (data not shown), implying its essential role in iron-deprived states. This differential expression of C. albicans HAP43 has not been previously reported. A similar upregulation in response to low-iron conditions was, however, reported for S. pombe php4+ and A. nidulans hapX (34, 57).

Deletion of HAP43 led to the increased expression of ACO1, SDH2, CYC1, CCP1, and YHB1 under iron-deprived conditions (Fig. 5 A). These genes are all downregulated in response to iron deficiency (49). ACO1 and SDH2 encode aconitase and succinate dehydrogenase, respectively, and are both Fe-S cluster proteins involved in the TCA cycle. CYC1, CCP1, and YHB1 encode heme-containing proteins. Cyc1 is the cytochrome c responsible for transferring electrons from mitochondrial complex III to complex IV. The amino acid sequence of Ccp1 is similar to the N terminus of cytochrome c peroxidase, and Yhb1 is predicted to be a nitric oxide oxidoreductase. Our quantitative gene expression analysis was also applied to genes required for Fe-S cluster assembly. We analyzed four genes (ATM1, ISA1, ISU1, and YAH1) whose functions were predicted based on their gene homologs in S. cerevisiae (15, 51, 52, 73). These genes were all derepressed in the cells lacking Hap43 under low-iron conditions (Fig. 5B). ATM1 encodes a mitochondrial inner membrane ABC transporter that is possibly involved in exporting mitochondrially synthesized precursors of Fe-S clusters to the cytosol. ISA1 and ISU1 gene products are predicted to be a mitochondrial matrix protein and a scaffold protein required for the biogenesis of the Fe-S cluster, respectively. YAH1 encodes a ferredoxin homologous to S. cerevisiae Yah1, which has electron transfer activity involved in the Fe-S cluster biosynthesis. The 5′ upstream regions of theses potential Hap43 target genes (ACO1, SDH2, CYC1, CCP1, YHB1, ATM1, ISA1, ISU1, and YAH1) all contain multiple CCAAT motifs (data not shown), suggesting that Hap43/CBC may directly regulate the expression of theses genes.

Fig. 5.
Hap43 represses the transcription of genes encoding iron-dependent proteins under iron-deprived conditions. Quantitative real-time PCR was performed for selected iron-responsive genes. Cells were inoculated into high-iron (YPD) or low-iron (YPD plus 400 ...

Furthermore, other genes encoding iron-dependent proteins such as CYT1, CAT1, and BIO2, and one gene encoding a non-iron-carrying protein, Hem1, were analyzed (data not shown). The CYT1 gene product, cytochrome c1, is a heme-containing protein that constitutes a subunit of complex III in the electron transport chain. The CAT1 product is a catalase responsible for the breakdown of H2O2 to protect cells from oxidative stress, especially when iron is overloaded (89). BIO2 encodes an Fe-S cluster protein and is predicted to be a biotin synthase that participates in biotin biosynthesis. Hem1 is described as a 5-aminolevulinate synthase that catalyzes the first step of the heme biosynthetic pathway. A. nidulans hemA, the homolog of the C. albicans Hem1 gene, is repressed by HapX under low-iron conditions (34). Interestingly, no obvious differential expression of these four genes was observed in the absence of C. albicans Hap43 in response to iron deprivation (data not shown).

The expression of a ferric reductase gene, FRP1, was also quantified. A previous report suggested that expression of FRP1 is regulated by CBC factors and that Hap43 may be involved in this regulation (2). In contrast to the expression pattern of genes encoding iron-dependent proteins, FRP1 is upregulated (by 22-fold) in response to iron deficiency, whereas the deletion of HAP43 decreased the low-iron-induced activation (to only 10-fold) (Fig. 5C), implying a positive role for Hap43 in controlling some iron-responsive genes.

Iron levels affect the transcriptional activity of Hap43.

The data suggested that Hap43 plays a role in iron homeostasis as a repressor to downregulate the expression of genes encoding iron-dependent proteins when iron is not sufficient (Fig. 5A and B). The sequence homolog of Hap43 in budding yeast, Hap4, acts as an activator to regulate respiratory gene expression (21). Php4 in fission yeast is a negative regulatory subunit of the CBC complex under low-iron conditions (57). Similar repression activity of HapX, together with CBC factors, has also been reported (34). However, Hap43 plays a positive role in the regulation of FRP1 expression (Fig. 5C). Accordingly, to further investigate the role of C. albicans Hap43 in transcriptional regulation, the one-hybrid system was used (75). We fused the sequence of the Staphylococcus aureus LexA DNA-binding domain to the N terminus of Hap43 and constitutively expressed the fusion protein in the lacZ reporter-carrying strains COP1 and CCR1. The COP1-derived strains use a LexA operator and an ADH1 basal promoter to drive the lacZ reporter, whereas CCR1-derived strains lack the LexA operator. Both the X-Gal overlay assay and the quantitative β-galactosidase assay were performed. Strains that express LexA-Gcn4 and LexA-Nrg1 were included as controls for the activator and repressor, respectively. Although not as strong as the activity of LexA-Gcn4, the LacZ activity generated by LexA-Hap43 was about 3-fold higher than that of basal controls (strains without the LexA operator) in the YPD medium (Fig. 6 A). Because YPD medium represents an iron-sufficient condition (Fig. 2B), we then examined the activity of Hap43 via the X-Gal overlay assay under low-iron conditions by adding 200 μM BPS or under the excess-iron condition by adding 50 μM Fe2+ to YPD agar plates. The activation of the lacZ reporter by Hap43 completely disappeared when cells were grown on the low-iron agar plate, even when the plate was incubated at 37°C to enhance the development of the blue color (Fig. 6B). This result suggests that environmental iron can regulate the transcriptional activity of Hap43.

Fig. 6.
One-hybrid analysis demonstrated that Hap43 is an activator under iron-rich conditions and a repressor under iron-deficient conditions. LexA-Hap43 binds to the LexA operator (LexAOP) upstream of the basal promoter of the lacZ reporter gene and modulates ...

Moreover, we proposed that the availability of iron might alter Hap43 activity in a dose-dependent manner. To test this hypothesis, different concentrations of BPS from 25 to 800 μM were added to YPD to restrict iron availability, and β-galactosidase assays for expression of the reporter were conducted. The transactivation activity of Hap43 was maximal in the absence of the iron chelator BPS and gradually decreased in the presence of increasing levels of BPS (Fig. 6C). Interestingly, Hap43 activity declined to the basal level under low-iron conditions (200 μM BPS) and appeared lower than basal activity under iron-starved conditions (400 and 800 μM BPS).

From the expression analyses (Fig. 5) and one-hybrid assays (Fig. 6A to C), we were still unable to decide the biological role of Hap43 under high-iron conditions; however, they raised a strong possibility of Hap43 serving as a transcriptional repressor under low-iron conditions. To assess this possibility, we determined the repression activity of Hap43 in a more precise system. By introducing (GCRE)5, a penta-general control response element sequence, into the reporter strains between the S. aureus LexA operator and the ADH1 basal promoter, the expression of the lacZ reporter can be enhanced in a Gcn4-dependent manner (85). We adapted this idea to generate new reporter strains that exhibited basal levels of lacZ higher than that of the original LexA operator-only strain, allowing more sensitive and accurate evaluations of gene repression mediated by Hap43. As shown in Fig. 6D, the LexA-only strains containing lexAOP-(GCRE)5-lacZ showed a 30-fold increase in the basal level of reporter gene expression in comparison with the LexA operator-only strains (Fig. 6A) under either low- or high-iron conditions. Cells expressing the LexA-Hap43 fusion protein did, however, have a dramatic reduction in lacZ expression under low-iron conditions (Fig. 6D and E). Furthermore, the presence of Hap43 also resulted in LacZ activity higher than that in the LexA-only strains in YPD. This result is consistent with the data shown in Fig. 6A to C, which imply that Hap43 potentially plays a positive role in transcriptional regulation only under iron-rich conditions. Taken together, these data suggest that Hap43 regulates iron homeostasis by turning into a repressor in response to iron deficiency.

Iron limitation induces nuclear accumulation of Hap43.

To elucidate the possible mechanism by which Hap43 is regulated in response to external iron levels, we examined the cellular localization of Hap43 by confocal microscopy. To correlate these results with the data from the one-hybrid assays, we modified the Hap43 one-hybrid plasmid by replacing the LexA coding sequence with the sequence from Candida-adapted GFP. This construct was transformed into CAI4 to create the GFP-Hap43-expressing strains. The strains were grown in YPD overnight and subcultured in iron-depleted and iron-sufficient media (YPD with and without 400 μM BPS, respectively). After a 5-h culture period, GFP-Hap43 proteins accumulated in the nucleus only under iron-depleted conditions (Fig. 7 A). Similar results were seen when 400 μM BPS was used to treat cells for 2 h (data not shown), indicating that the nuclear accumulation of Hap43 is a process regulated by a response to environmental iron rather than by a growth-dependent mechanism. In addition, we excluded the possibility that the expression of GFP-HAP43 is regulated at the transcriptional level by using a constitutive promoter-driven expression system similar to the one used in the one-hybrid strains (see Materials and Methods). Thus, we focused on the regulation of Hap43 at the protein level. To evaluate the expression of GFP-Hap43 under low- or high-iron conditions, immunoblotting was used to detect GFP fusion proteins with monoclonal anti-GFP (Fig. 7B). The results indicated that the fusion proteins were expressed under both iron conditions. However, an unexpected supershift of the bands corresponding to GFP-Hap43 was observed on SDS-PAGE (from the predicted position at 97 kDa to ~120 kDa) independent of iron conditions. Moreover, more GFP-Hap43 was present under low-iron conditions. In summary, the iron-responsiveness of Hap43 is in part regulated by the cytonuclear localization.

Fig. 7.
Iron deficiency induces nuclear accumulation of Hap43. The C. albicans CAI4 strain was transformed with a GFP-fused Hap43 under the control of the ACT1 promoter. Cells expressing GFP-Hap43 were grown in YPD overnight and inoculated into iron-depleted ...

Iron starvation induces production of flavin molecules only in the presence of Hap43.

C. albicans can secrete flavin in response to iron limitation. This activity is controlled by the general transcriptional corepressor Tup1. Flavin production by the tup1Δ strain is elevated in spite of the increased iron in the YPD medium (47). Because Tup1 is also a negative regulator of genes involved in iron homeostasis, we proposed that other iron-responsive repressors might also participate in the process of flavinogenesis. To test this hypothesis, the regulatory roles of Sfu1, Hap43, and Tup1 in low-iron-induced flavinogenesis were examined. For the convenience of observational and spectrometric analyses, transparent YNB-based NIM and SC medium were used instead of a YPD-based medium to induce the production of flavin. Sfu1 had no effect on low-iron-induced flavinogenesis compared with the wild-type strain (Fig. 8). Hap43 was, however, essential for the iron-responsive process of flavin production. In addition, Tup1 was required to sustain the production of flavin under iron-depleted conditions (NIM). Deletion of TUP1 led to a 50% reduction in flavin production (Fig. 8). These findings suggest that Hap43 may cooperate with Tup1 to regulate transcriptional and metabolic remodeling, especially under conditions of iron depletion.

Fig. 8.
Hap43 is essential for the flavinogenesis induced by iron starvation. (A) The absorbance at 446.3 nm of each supernatant from the stationary-phase culture in NIM (Fe) or SC medium (Fe+) is displayed. This measurement is directly correlated with ...


Iron is essential for almost all organisms. Both iron deficiency and iron overload cause deleterious consequences for living cells. C. albicans maintains iron homeostasis by remodeling transcriptional programs to regulate iron-responsive genes. Besides the high-iron-specific repressor Sfu1 (49), we have shown that the low-iron-specific repressor Hap43 is also important for the regulation of C. albicans iron homeostasis.

Hap43 contains a conserved N-terminal region composed of a putative CBC-interacting domain and a bZip domain. It functions more similarly to HapX/Php4 than to yeast Hap4 and another bZip transcription factor, Yap5. Both Hap4 and Yap5 lack either one of the conserved structures described above. Apart from Hap43, another two possible Hap43 homologs (Hap41 and Hap42) exist in C. albicans (43). Interestingly, C. albicans Hap41 and Hap42 contain the putative CBC-interacting domains, like S. cerevisiae Hap4, but lack the bZip structure (Fig. 1A). Hap41 and Hap42 display lower identity/similarity to A. nidulans HapX (19.1%/33.7% and 14.2%/26.7%, respectively) than does Hap43. Moreover, based on the work of Baek et al. (2), Hap41 is not required for the limiting iron-restricted binding of the FRP1 promoter and makes no contribution to low-iron-dependent growth (YPG medium plus 100 μM BPS). Both Hap5 and Hap43, however, do play essential roles in low-iron cell growth. Therefore, we doubt that Hap41 and Hap42 are required for adaptation to iron-deficient environments. An analysis of the functions of Hap41/Hap42 is currently under way.

Accumulated evidence has suggested that iron is strongly correlated with microbial infections, including those caused by zoopathogenic and opportunistic fungi (35, 36, 82). In addition, iron-restricting therapies are effective in treating fungal infections in mouse models (39,42). Several iron-related proteins contribute to the virulence of C. albicans and other pathogenic fungi (31, 32, 44, 71, 79, 80). In our study, we provided the first example in C. albicans of an iron-responsive transcription factor, Hap43, which is essential under low-iron conditions and can contribute to virulence in the disseminated infection (Fig. 3). Notably, most of the gene deletions described above lead to defects in iron uptake, and this lack of capability may consequently decrease the fitness of C. albicans to survive within the host. Furthermore, cryptococcal Cir1 is reported to regulate many virulence factors, such as capsule and melanin production and the capability to survive at 37°C, as well as iron acquisition (24). Nevertheless, in C. albicans, it seems that the administration of iron uptake is not the major causative factor of Hap43-mediated virulence, based on our results (Fig. 4). Instead, hap43Δ cells had a higher intracellular iron content under iron-sufficient conditions, as did the sfu1Δ cells (Fig. 4C), and after overnight iron starvation than the wild type and all other strains tested (Fig. 4A, 0 min). In addition, the hap43Δ mutant showed no increased ferric reductase activity and phleomycin sensitivity, as did the sfu1Δ strain, implying that different molecular and biochemical events occur in the absence of Hap43 and consequently cause the accumulation of a higher intracellular iron content.

Among fungal HapX/Php4 homologs, apart from C. albicans Hap43, only A. fumigatus HapX has been shown to be required for virulence so far (79). The attenuated virulence in the hapXΔ strain is possibly caused by the general dysregulation of gene expression essential for metabolic adaptation to iron deficiency, the accumulation of toxic metabolites, and/or the decreased expression of possible virulence determinants. In C. albicans, we found that the deletion of HAP43 does not affect the morphogenesis of C. albicans in hypha-inducing spider medium (data not shown), but the contribution of Hap43 to the regulation of other virulence factors, such as secreted proteases, lipases, adhesins, and biofilm formation, is still unclear. Nevertheless, we demonstrated that Hap43 is essential for the repression of many genes that encode iron-dependent proteins, especially those involved in the TCA cycle, respiration, and Fe-S cluster assembly (Fig. 5A and B), indicating that Hap43 possesses general functions similar to those of Aspergillus HapX (34) and S. pombe Php4 (57).

According to our hypothesis and experimental evidence (Fig. 4), we have de-emphasized the regulation of iron uptake genes in the hap43Δ mutant and have instead focused on the role of Hap43 in the transcriptional repression of genes encoding iron-dependent proteins. Hap43 acted as a positive transcription factor under iron-rich conditions, and this activating activity decreased with the decline of iron availability (Fig. 6A, B, and C). We also provided evidence that low-iron status converts Hap43 to a transcriptional repressor (Fig. 6D). The iron responsiveness of Hap43 can be interpreted in two possible ways. One is that Hap43 loses its ability to be an activator under low-iron conditions, and the other is based on a molecular conversion of Hap43 from an activator to a repressor in response to iron deficiency. Some bacterial regulators of metal homeostasis also possess this contrary activity to activate or repress metal assimilation or consumption (16, 37). S. pombe Php4 does not, however, exhibit activation activity under high-iron conditions (57). Therefore, we cannot exclude the possibility that the activating activity of Hap43 is an artifact that results from the constitutive expression of HAP43 as controlled by the ACT1 promoter or from the possible absence of CBC (Hap2, Hap3, and Hap5 [Hap2/3/5]) factors that were replaced by the LexA DNA-binding domain in the one-hybrid assays. Furthermore, although the N-terminal LexA seems to have no influence on the activity of Hap43 under low-iron conditions, we do not know whether LexA affects the activity, stability, structure, or transcriptional partners of Hap43 under high-iron conditions, even though the level of Hap43 is lower than that under low-iron conditions. Deletion of HAP43 only partially decreased the expression of FRP1 in response to low-iron conditions (Fig. 5C). In the study of Baek et al. (2), the interaction between the FRP1 promoter and the protein extracts from the hap43Δ strain were assayed by electrophoretic mobility shift assay (EMSA). They showed that a band was only slightly reduced (not completely absent) in the iron-starved hap43Δ cells compared with its expression in the wild type. Our explanation for this finding is that Hap43 may play an indirect role by negatively regulating an unidentified repressor that controls the expression of FRP1. Therefore, the absence of Hap43 leads to the elevation of the unidentified repressor and results in reduced FRP1 induction in response to iron deprivation. Moreover, the deletion of HAP43 led to accumulation in the cell of phleomycin-inaccessible iron (Fig. 4D). To explain this observation, we hypothesize that the absence of Hap43 elevated the level of iron-dependent proteins by derepressing gene expression (Fig. 5A and B), leading to an increase in iron binding by these resultant proteins and a decrease in phleomycin-inaccessible iron. This hypothesis is able to explain only the higher intracellular iron content in the iron-starved hap43Δ mutant (Fig. 4A, at 0 min) but cannot explain why hap43Δ cells maintain a high level of iron when grown under iron-sufficient conditions (Fig. 4C).

Although the detailed mechanism for the regulation of Hap43 activity is not clear, Hap43 dispersed throughout the cell under conditions of iron sufficiency and accumulated in the nucleus when iron levels were low. A. nidulans HapX (34) and S. pombe Php4 (55) display similar nucleocytoplasmic shuttling in response to iron levels. Some evidence suggests that the mechanisms that regulate HapX/Php4/Hap43 translocalization and their activities in target gene repression are much more complicated. For example, the iron-induced export of S. pombe Php4 from the nucleus relies on a physical interaction with a monothiol glutaredoxin, Grx4. Deletion of grx4+ leads to the nuclear retention of Php4 and sustained repression of its target gene, even when iron is sufficient (55). In the case of yeast Aft1, iron-mediated inhibition requires two glutaredoxins, Grx3 and Grx4 (61, 69). Loss of both Grx3 and Grx4, but not loss of just one of them, causes the sustained nuclear localization of Aft1 and constitutive expression of Aft1 target genes. In C. albicans, there are at least four putative glutaredoxin genes (GRX1, GRX3, TTR1, and GRX5) annotated in the CGD, but the expression of only GRX1 is induced in response to low-iron conditions (49). We generated a C. albicans grx1Δ strain but found that there was no significant effect on the expression of ISA1, ATM1, CCP1, and YHB1 when GRX1 was deleted (data not shown). Therefore, determining the role of each glutaredoxin in the iron-responsive regulation of Hap43 activity requires further investigation. Furthermore, translocation of Php4 depends on the Leu-rich nuclear export signal (93LLEQLEML100) and the exportin Crm1 (55). In C. albicans, a putative Leu-rich nuclear export signal (129LVNTINKLKV138) was predicted in Hap43. This signal overlaps with part of the leucine zipper sequence (129LVNTINKLKVENQFLVKNLEQL150 [structural leucines are underlined]) of Hap43 (Fig. 1A), and it provides a potential domain for the iron-responsive regulation of Hap43.

There is a link between iron metabolism and flavinogenesis in eukaryotic cells (17). For example, in the riboflavin-overproducing yeast Pichia guilliermondii, flavinogenesis is coregulated with reductive iron assimilation (19). In addition, iron can transcriptionally repress riboflavin synthesis in Pichia (4). A deficiency in mitochondrial frataxin, which is involved in iron trafficking and storage, leads to dysregulation of riboflavin biosynthesis (70). In the case of bacteria, secreted flavins may be advantageous for iron acquisition by acting as electron shuttles to facilitate the reduction of insoluble ferric ions in the surrounding environment or to assist in the release of iron from ferrisiderophores (14, 87). However, the physiological function of inducible flavinogenesis remains to be elucidated. Recently, flavinogenic Candida species have been characterized (47, 88, 91). In C. albicans, the iron starvation-induced production of flavin is regulated by Tup1, which was originally identified as a corepressor and functions as a global regulator of morphogenesis, metabolism, phase switching, and mating (6, 59, 63, 92). Deletion of TUP1 causes an elevation of flavin production that is independent of iron levels in YPD medium (47). Interestingly, in our study, we found that Tup1 exerted a positive effect on low-iron-induced flavinogenesis in the acidic defined medium (YNB based). The iron-responsive induction of flavinogenesis is present in the tup1Δ strain, but this induction shows a 50% reduction in comparison with the wild-type strain. Taken together, these findings suggest potential cooperative roles for Hap43 and Tup1 in the regulation of iron-dependent metabolism, such as flavinogenesis.

To summarize, a simplified model of Hap43-mediated regulation in response to iron availability is proposed (Fig. 9). In this model, Sfu1 represses excess iron assimilation in coordination with the corepressor Tup1 by repressing the expression of genes encoding iron uptake proteins under iron-rich conditions. Under iron-deficient conditions, nuclear-accumulated Hap43 represses the expression of SFU1 to relieve the depression of iron acquisition and also represses the expression of iron-dependent genes to reapportion the usage of restricted iron. Furthermore, the expression of HAP43 is partially repressed by Sfu1 when iron is sufficient to activate the expression of genes encoding iron-dependent proteins. Hap43 also positively regulates low-iron-induced flavinogenesis independently or cooperates with Tup1-mediated regulation. These complicated and interacting functions may contribute to the normal growth and virulence of C. albicans within the host. Although Hap43 appeared to cooperate with CBC factors in a study of FRP1 transcription regulation (2), deletion of the CBC subunit HAP2, HAP3, or HAP5 generated some phenotypes that are inconsistent with that observed in the hap43Δ strain (33). For example, hap2Δ and hap5Δ strains are hyperresistant to rapamycin and caffeine, in contrast to the wild-type, hap43Δ, and hap3Δ strains (33). Additionally, the growth defect of hap43Δ under low-iron conditions was more severe than that of hap2Δ, hap3Δ, and hap5Δ in Homann et al. (33) and based on our unpublished data. These inconsistencies indicate that not all cell functions mediated by Hap43 rely on cooperation with CBC factors. Therefore, more investigations are required to understand the role of each CBC factor in Hap43-mediated iron homeostasis.

Fig. 9.
A simple model of Hap43-mediated iron metabolism is proposed. When cells encounter a shift from iron sufficiency to iron deficiency, the expression of HAP43 is released from repression by Sfu1. In turn, Hap43 is induced to repress the expression of SFU1 ...


This work was supported by grants NSC95-2311-B-007-022-MY3, NSC98-2627-B-007-015, and NSC99-2627-B-007-007 from the National Science Council, Taiwan (to C.-Y.L.).

We thank Alistair J. P. Brown for the kind gift of one-hybrid materials (pCIplexA, pCR-lacZ, pCR-OPlacZ, and CAI8). We also thank Joachim Morschhäuser for the generous gifts of SAT1-FLIP gene deletion cassettes (pSFS1A and pSFS2A) and the GFP template from pNIM1. We thank Yu-Ting Chen for technical assistance with confocal microscopy.


[down-pointing small open triangle]Published ahead of print on 3 December 2010.


1. Archibald F. 1983. Lactobacillus plantarum, an organism not requiring iron. FEMS Microbiol. Lett. 19:29–32
2. Baek Y.-U., Li M., Davis D. A. 2008. Candida albicans ferric reductases are differentially regulated in response to distinct forms of iron limitation by the Rim101 and CBF transcription factors. Eukaryot. Cell 7:1168–1179 [PMC free article] [PubMed]
3. Blaiseau P. L., Lesuisse E., Camadro J. M. 2001. Aft2p, a novel iron-regulated transcription activator that modulates, with Aft1p, intracellular iron use and resistance to oxidative stress in yeast. J. Biol. Chem. 276:34221–34226 [PubMed]
4. Boretsky Y. R., et al. 2005. Positive selection of mutants defective in transcriptional repression of riboflavin synthesis by iron in the flavinogenic yeast Pichia guilliermondii. FEMS Yeast Res. 5:829–837 [PubMed]
5. Bourgarel D., Nguyen C. C., Bolotin-Fukuhara M. 1999. HAP4, the glucose-repressed regulated subunit of the HAP transcriptional complex involved in the fermentation-respiration shift, has a functional homologue in the respiratory yeast Kluyveromyces lactis. Mol. Microbiol. 31:1205–1215 [PubMed]
6. Braun B. R., Johnson A. D. 1997. Control of filament formation in Candida albicans by the transcriptional repressor TUP1. Science 277:105–109 [PubMed]
7. Braun B. R., Johnson A. D. 2000. TUP1, CPH1 and EFG1 make independent contributions to filamentation in Candida albicans. Genetics 155:57–67 [PMC free article] [PubMed]
8. Bullen J. J., Rogers H. J., Spalding P. B., Ward C. G. 2006. Natural resistance, iron and infection: a challenge for clinical medicine. J. Med. Microbiol. 55:251–258 [PubMed]
9. Campanella J. J., Bitincka L., Smalley J. 2003. MatGAT: an application that generates similarity/identity matrices using protein or DNA sequences. BMC Bioinformatics 4:29. [PMC free article] [PubMed]
10. Chao L. Y., Marletta M. A., Rine J. 2008. Sre1, an iron-modulated GATA DNA-binding protein of iron-uptake genes in the fungal pathogen Histoplasma capsulatum. Biochemistry 47:7274–7283 [PubMed]
11. Ciais D., Bohnsack M. T., Tollervey D. 2008. The mRNA encoding the yeast ARE-binding protein Cth2 is generated by a novel 3′ processing pathway. Nucleic Acids Res. 36:3075–3084 [PMC free article] [PubMed]
12. Conde e Silva N., et al. 2009. KlAft, the Kluyveromyces lactis ortholog of Aft1 and Aft2, mediates activation of iron-responsive transcription through the PuCACCC Aft-type sequence. Genetics 183:93–106 [PMC free article] [PubMed]
13. Courel M., Lallet S., Camadro J. M., Blaiseau P. L. 2005. Direct activation of genes involved in intracellular iron use by the yeast iron-responsive transcription factor Aft2 without its paralog Aft1. Mol. Cell. Biol. 25:6760–6771 [PMC free article] [PubMed]
14. Coves J., Fontecave M. 1993. Reduction and mobilization of iron by a NAD(P)H:flavin oxidoreductase from Escherichia coli. Eur. J. Biochem. 211:635–641 [PubMed]
15. Craig E. A., Marszalek J. 2002. A specialized mitochondrial molecular chaperone system: a role in formation of Fe/S centers. Cell Mol. Life Sci. 59:1658–1665 [PubMed]
16. da Silva Neto J. F., Braz V. S., Italiani V. C., Marques M. V. 2009. Fur controls iron homeostasis and oxidative stress defense in the oligotrophic alpha-proteobacterium Caulobacter crescentus. Nucleic Acids Res. 37:4812–4825 [PMC free article] [PubMed]
17. Demain A. L. 1972. Riboflavin oversynthesis. Annu. Rev. Microbiol. 26:369–388 [PubMed]
18. Drakesmith H., Prentice A. 2008. Viral infection and iron metabolism. Nat. Rev. Microbiol. 6:541–552 [PubMed]
19. Fedorovich D., Protchenko O., Lesuisse E. 1999. Iron uptake by the yeast Pichia guilliermondii. Flavinogenesis and reductive iron assimilation are co-regulated processes. Biometals 12:295–300 [PubMed]
20. Fonzi W. A., Irwin M. Y. 1993. Isogenic strain construction and gene mapping in Candida albicans. Genetics 134:717–728 [PMC free article] [PubMed]
21. Forsburg S. L., Guarente L. 1989. Identification and characterization of HAP4: a third component of the CCAAT-bound HAP2/HAP3 heteromer. Genes Dev. 3:1166–1178 [PubMed]
22. Geber A., Williamson P. R., Rex J. H., Sweeney E. C., Bennett J. E. 1992. Cloning and characterization of a Candida albicans maltase gene involved in sucrose utilization. J. Bacteriol. 174:6992–6996 [PMC free article] [PubMed]
23. Gillum A. M., Tsay E. Y. H., Kirsch D. R. 1984. Isolation of the Candida albicans gene for orotidine-5′-phosphate decarboxylase by complementation of S. cerevisiae ura3 and E. coli pyrF mutations. Mol. Gen. Genet. 198:179–182 [PubMed]
24. Gross L. 2006. Iron regulation and an opportunistic AIDS-related fungal infection. PLoS Biol. 4:e427. [PMC free article] [PubMed]
25. Haas H., Angermayr K., Stöffler G. 1997. Molecular analysis of a Penicillium chrysogenum GATA factor encoding gene (sreP) exhibiting significant homology to the Ustilago maydis urbs1 gene. Gene 184:33–37 [PubMed]
26. Haas H., Eisendle M., Turgeon B. G. 2008. Siderophores in fungal physiology and virulence. Annu. Rev. Phytopathol. 46:149–187 [PubMed]
27. Haas H., Zadra I., Stoffler G., Angermayr K. 1999. The Aspergillus nidulans GATA factor SREA is involved in regulation of siderophore biosynthesis and control of iron uptake. J. Biol. Chem. 274:4613–4619 [PubMed]
28. Halliwell B., Gutteridge J. M. 1984. Oxygen toxicity, oxygen radicals, transition metals and disease. Biochem. J. 219:1–14 [PMC free article] [PubMed]
29. Halliwell B., Gutteridge J. M. C. 1992. Biologically relevant metal ion-dependent hydroxyl radical generation. An update. FEBS Lett. 307:108–112 [PubMed]
30. Hentze M. W., Muckenthaler M. U., Andrews N. C. 2004. Balancing acts: molecular control of mammalian iron metabolism. Cell 117:285–297 [PubMed]
31. Heymann P., et al. 2002. The siderophore iron transporter of Candida albicans (Sit1p/Arn1p) mediates uptake of ferrichrome-type siderophores and is required for epithelial invasion. Infect. Immun. 70:5246–5255 [PMC free article] [PubMed]
32. Hissen A. H., Wan A. N., Warwas M. L., Pinto L. J., Moore M. M. 2005. The Aspergillus fumigatus siderophore biosynthetic gene sidA, encoding L-ornithine N5-oxygenase, is required for virulence. Infect. Immun. 73:5493–5503 [PMC free article] [PubMed]
33. Homann O. R., Dea J., Noble S. M., Johnson A. D. 2009. A phenotypic profile of the Candida albicans regulatory network. PLoS Genet. 5:e1000783. [PMC free article] [PubMed]
34. Hortschansky P., et al. 2007. Interaction of HapX with the CCAAT-binding complex—a novel mechanism of gene regulation by iron. EMBO J. 26:3157–3168 [PMC free article] [PubMed]
35. Howard D. H. 1999. Acquisition, transport, and storage of iron by pathogenic fungi. Clin. Microbiol. Rev. 12:394–404 [PMC free article] [PubMed]
36. Howard D. H. 2004. Iron gathering by zoopathogenic fungi. FEMS Immunol. Med. Microbiol. 40:95–100 [PubMed]
37. Huang D. L., et al. 2008. The Zur of Xanthomonas campestris functions as a repressor and an activator of putative zinc homeostasis genes via recognizing two distinct sequences within its target promoters. Nucleic Acids Res. 36:4295–4309 [PMC free article] [PubMed]
38. Hube B., Monod M., Schofield D. A., Brown A. J. P., Gow N. A. R. 1994. Expression of seven members of the gene family encoding secretory aspartyl proteinases in Candida albicans. Mol. Microbiol. 14:87–99 [PubMed]
39. Ibrahim A. S., Edwards J. E., Jr., Fu Y., Spellberg B. 2006. Deferiprone iron chelation as a novel therapy for experimental mucormycosis. J. Antimicrob. Chemother. 58:1070–1073 [PubMed]
40. Ibrahim A. S., et al. 2007. The iron chelator deferasirox protects mice from mucormycosis through iron starvation. J. Clin. Invest. 117:2649–2657 [PMC free article] [PubMed]
41. Ibrahim A. S., Gebremariam T., French S. W., Edwards J. E., Jr., Spellberg B. 2010. The iron chelator deferasirox enhances liposomal amphotericin B efficacy in treating murine invasive pulmonary aspergillosis. J. Antimicrob. Chemother. 65:289–292 [PMC free article] [PubMed]
42. Ibrahim A. S., Spellberg B., Edwards J., Jr 2008. Iron acquisition: a novel perspective on mucormycosis pathogenesis and treatment. Curr. Opin. Infect. Dis. 21:620–625 [PMC free article] [PubMed]
43. Johnson D. C., Cano K. E., Kroger E. C., McNabb D. S. 2005. Novel regulatory function for the CCAAT-binding factor in Candida albicans. Eukaryot. Cell 4:1662–1676 [PMC free article] [PubMed]
44. Jung W. H., Sham A., White R., Kronstad J. W. 2006. Iron regulation of the major virulence factors in the AIDS-associated pathogen Cryptococcus neoformans. PLoS Biol. 4:e410. [PMC free article] [PubMed]
45. Kaplan J., McVey Ward D., Crisp R. J., Philpott C. C. 2006. Iron-dependent metabolic remodeling in S. cerevisiae. Biochim. Biophys. Acta 1763:646–651 [PubMed]
46. Knight S. A., Dancis A. 2006. Reduction of 2,3-bis(2-methoxy-4-nitro-5-sulfophenyl)-2H-tetrazolium-5-carboxanilide inner salt (XTT) is dependent on CaFRE10 ferric reductase for Candida albicans grown in unbuffered media. Microbiology 152:2301–2308 [PubMed]
47. Knight S. A. B., Lesuisse E., Stearman R., Klausner R. D., Dancis A. 2002. Reductive iron uptake by Candida albicans: role of copper, iron and the TUP1 regulator. Microbiology 148:29–40 [PubMed]
48. Kornitzer D. 2009. Fungal mechanisms for host iron acquisition. Curr. Opin. Microbiol. 12:377–383 [PubMed]
49. Lan C. Y., et al. 2004. Regulatory networks affected by iron availability in Candida albicans. Mol. Microbiol. 53:1451–1469 [PubMed]
50. Larkin M. A., et al. 2007. Clustal W and Clustal X version 2.0. Bioinformatics 23:2947–2948 [PubMed]
51. Lill R., Muhlenhoff U. 2005. Iron-sulfur-protein biogenesis in eukaryotes. Trends Biochem. Sci. 30:133–141 [PubMed]
52. Lill R., Muhlenhoff U. 2006. Iron-sulfur protein biogenesis in eukaryotes: components and mechanisms. Annu. Rev. Cell Dev. Biol. 22:457–486 [PubMed]
53. Macheroux P. 1999. UV-Visible spectroscopy as a tool to study flavoproteins. Methods Mol Biol. 131:1–7 [PubMed]
54. McNabb D., Tseng K., Guarente L. 1997. The Saccharomyces cerevisiae Hap5p homolog from fission yeast reveals two conserved domains that are essential for assembly of heterotetrameric CCAAT-binding factor. Mol. Cell Biol. 17:7008–7018 [PMC free article] [PubMed]
55. Mercier A., Labbe S. 2009. Both Php4 function and subcellular localization are regulated by iron via a multistep mechanism involving the glutaredoxin Grx4 and the exportin Crm1. J. Biol. Chem. 284:20249–20262 [PMC free article] [PubMed]
56. Mercier A., Pelletier B., Labbe S. 2006. A transcription factor cascade involving Fep1 and the CCAAT-binding factor Php4 regulates gene expression in response to iron deficiency in the fission yeast Schizosaccharomyces pombe. Eukaryot. Cell 5:1866–1881 [PMC free article] [PubMed]
57. Mercier A., Watt S., Bahler J., Labbe S. 2008. Key function for the CCAAT-binding factor Php4 to regulate gene expression in response to iron deficiency in fission yeast. Eukaryot. Cell 7:493–508 [PMC free article] [PubMed]
58. Miele R., Barra D., Bonaccorsi di Patti M. C. 2007. A GATA-type transcription factor regulates expression of the high-affinity iron uptake system in the methylotrophic yeast Pichia pastoris. Arch. Biochem. Biophys. 465:172–179 [PubMed]
59. Murad A. M., et al. 2001. Transcript profiling in Candida albicans reveals new cellular functions for the transcriptional repressors CaTup1, CaMig1 and CaNrg1. Mol. Microbiol. 42:981–993 [PubMed]
60. Oberegger H., et al. 2002. Identification of members of the Aspergillus nidulans SREA regulon: genes involved in siderophore biosynthesis and utilization. Biochem. Soc. Trans. 30:781–783 [PubMed]
61. Ojeda L., et al. 2006. Role of glutaredoxin-3 and glutaredoxin-4 in the iron regulation of the Aft1 transcriptional activator in Saccharomyces cerevisiae. J. Biol. Chem. 281:17661–17669 [PubMed]
62. Park Y.-N., Morschhauser J. 2005. Tetracycline-inducible gene expression and gene deletion in Candida albicans. Eukaryot. Cell 4:1328–1342 [PMC free article] [PubMed]
63. Park Y. N., Morschhauser J. 2005. Candida albicans MTLalpha tup1Delta mutants can reversibly switch to mating-competent, filamentous growth forms. Mol. Microbiol. 58:1288–1302 [PubMed]
64. Pelletier B., Beaudoin J., Mukai Y., Labbe S. 2002. Fep1, an iron sensor regulating iron transporter gene expression in Schizosaccharomyces pombe. J. Biol. Chem. 277:22950–22958 [PubMed]
65. Philpott C. C., Protchenko O. 2008. Response to iron deprivation in Saccharomyces cerevisiae. Eukaryot. Cell 7:20–27 [PMC free article] [PubMed]
66. Posey J. E., Gherardini F. C. 2000. Lack of a role for iron in the Lyme disease pathogen. Science 288:1651–1653 [PubMed]
67. Prouteau M., Daugeron M.-C., Seraphin B. 2008. Regulation of ARE transcript 3′ end processing by the yeast Cth2 mRNA decay factor. EMBO J. 27:2966–2976 [PMC free article] [PubMed]
68. Puig S., Askeland E., Thiele D. J. 2005. Coordinated remodeling of cellular metabolism during iron deficiency through targeted mRNA degradation. Cell 120:99–110 [PubMed]
69. Pujol-Carrion N., Belli G., Herrero E., Nogues A., de la Torre-Ruiz M. A. 2006. Glutaredoxins Grx3 and Grx4 regulate nuclear localisation of Aft1 and the oxidative stress response in Saccharomyces cerevisiae. J. Cell Sci. 119:4554–4564 [PubMed]
70. Pynyaha Y. V., et al. 2009. Deficiency in frataxin homologue YFH1 in the yeast Pichia guilliermondii leads to missregulation of iron acquisition and riboflavin biosynthesis and affects sulfate assimilation. Biometals 22:1051–1061 doi: 10.1007/s10534-009-9256-x [PMC free article] [PubMed]
71. Ramanan N., Wang Y. 2000. A high-affinity iron permease essential for Candida albicans virulence. Science 288:1062–1064 [PubMed]
72. Reuß O., Vik Å., Kolter R., Morschhäuser J. 2004. The SAT1 flipper, an optimized tool for gene disruption in Candida albicans. Gene 341:119–127 [PubMed]
73. Rouault T. A., Tong W. H. 2005. Iron-sulphur cluster biogenesis and mitochondrial iron homeostasis. Nat. Rev. Mol. Cell Biol. 6:345–351 [PubMed]
74. Rupp S. 2002. LacZ assays in yeast. Methods Enzymol. 350:112–131 [PubMed]
75. Russell C. L., Brown A. J. P. 2005. Expression of one-hybrid fusions with Staphylococcus aureus lexA in Candida albicans confirms that Nrg1 is a transcriptional repressor and that Gcn4 is a transcriptional activator. Fungal Genet. Biol. 42:676–683 [PubMed]
76. Rutherford J. C., Jaron S., Ray E., Brown P. O., Winge D. R. 2001. A second iron-regulatory system in yeast independent of Aft1p. Proc. Natl. Acad. Sci. U. S. A. 98:14322–14327 [PMC free article] [PubMed]
77. Sambrook J., Russell D. W. 2001. Molecular cloning: a laboratory manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY
78. Schaller M., Schafer W., Korting H. C., Hube B. 1998. Differential expression of secreted aspartyl proteinases in a model of human oral candidosis and in patient samples from the oral cavity. Mol. Microbiol. 29:605–615 [PubMed]
79. Schrettl M., et al. 2010. HapX-mediated adaption to iron starvation is crucial for virulence of Aspergillus fumigatus. PLoS Pathog. 6:e1001124. [PMC free article] [PubMed]
80. Schrettl M., et al. 2004. Siderophore biosynthesis but not reductive iron assimilation is essential for Aspergillus fumigatus virulence. J. Exp. Med. 200:1213–1219 [PMC free article] [PubMed]
81. Schrettl M., et al. 2008. SreA-mediated iron regulation in Aspergillus fumigatus. Mol. Microbiol. 70:27–43 [PMC free article] [PubMed]
82. Sutak R., Lesuisse E., Tachezy J., Richardson D. R. 2008. Crusade for iron: iron uptake in unicellular eukaryotes and its significance for virulence. Trends Microbiol. 16:261–268 [PubMed]
83. Tamarit J., Irazusta V., Moreno-Cermeño A., Ros J. 2006. Colorimetric assay for the quantitation of iron in yeast. Anal. Biochem. 351:149–151 [PubMed]
84. Tanaka A., Kato M., Nagase T., Kobayashi T., Tsukagoshi N. 2002. Isolation of genes encoding novel transcription factors which interact with the Hap complex from Aspergillus species. Biochim. Biophys. Acta 1576:176–182 [PubMed]
85. Tripathi G., et al. 2002. Gcn4 co-ordinates morphogenetic and metabolic responses to amino acid starvation in Candida albicans. EMBO J. 21:5448–5456 [PMC free article] [PubMed]
86. Voisard C., Wang J., McEvoy J. L., Xu P., Leong S. A. 1993. urbs1, a gene regulating siderophore biosynthesis in Ustilago maydis, encodes a protein similar to the erythroid transcription factor GATA-1. Mol. Cell. Biol. 13:7091–7100 [PMC free article] [PubMed]
87. von Canstein H., Ogawa J., Shimizu S., Lloyd J. R. 2008. Secretion of flavins by Shewanella species and their role in extracellular electron transfer. Appl. Environ. Microbiol. 74:615–623 [PMC free article] [PubMed]
88. Wang L., Chi Z., Wang X., Ju L., Guo N. 2008. Isolation and characterization of Candida membranifaciens subsp. flavinogenie W14-3, a novel riboflavin-producing marine yeast. Microbiol. Res. 163:255–266 [PubMed]
89. Wysong D. R., Christin L., Sugar A. M., Robbins P. W., Diamond R. D. 1998. Cloning and sequencing of a Candida albicans catalase gene and effects of disruption of this gene. Infect. Immun. 66:1953–1961 [PMC free article] [PubMed]
90. Yamaguchi-Iwai Y., Dancis A., Klausner R. D. 1995. AFT1: a mediator of iron regulated transcriptional control in Saccharomyces cerevisiae. EMBO J. 14:1231–1239 [PMC free article] [PubMed]
91. Yatsyshyn V. Y., Ishchuk O. P., Voronovsky A. Y., Fedorovych D. V., Sibirny A. A. 2009. Production of flavin mononucleotide by metabolically engineered yeast Candida famata. Metab. Eng. 11:163–167 [PubMed]
92. Zhao R., Lockhart S. R., Daniels K., Soll D. R. 2002. Roles of TUP1 in switching, phase maintenance, and phase-specific gene expression in Candida albicans. Eukaryot. Cell 1:353–365 [PMC free article] [PubMed]
93. Zhou L. W., Haas H., Marzluf G. A. 1998. Isolation and characterization of a new gene, sre, which encodes a GATA-type regulatory protein that controls iron transport in Neurospora crassa. Mol. Gen. Genet. 259:532–540 [PubMed]

Articles from Eukaryotic Cell are provided here courtesy of American Society for Microbiology (ASM)
PubReader format: click here to try


Related citations in PubMed

See reviews...See all...

Cited by other articles in PMC

See all...


  • Compound
    PubChem Compound links
  • MedGen
    Related information in MedGen
  • Pathways + GO
    Pathways + GO
    Pathways, annotations and biological systems (BioSystems) that cite the current article.
  • Protein
    Published protein sequences
  • PubMed
    PubMed citations for these articles
  • Substance
    PubChem Substance links
  • Taxonomy
    Related taxonomy entry
  • Taxonomy Tree
    Taxonomy Tree

Recent Activity

Your browsing activity is empty.

Activity recording is turned off.

Turn recording back on

See more...