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Vol. 16, Issue 10, 4792-4813, October 2005
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Department of Microbiology and Immunology, University of California-San Francisco, San Francisco, CA 94143-0414
Submitted May 18, 2005;
Revised June 30, 2005;
Accepted July 11, 2005
Monitoring Editor: Mark Solomon
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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4-fold upon nitrosative stress. Because our microarray clones were generated from random fragments of genomic DNA, they did not necessarily correspond to H. capsulatum open reading frames. To identify induced genes, we used high-density oligonucleotide tiling arrays to determine the genomic boundaries and coding strand of 153 RNS-induced transcripts. Homologues of these genes in other organisms are involved in iron acquisition, energy production, stress response, protein folding/degradation, DNA repair, and .NO detoxification. Ectopic expression of one of these genes, a P450 nitric oxide reductase homologue, was sufficient to increase resistance of H. capsulatum to RNS in culture. We propose that H. capsulatum uses the pathways identified here to cope with RNS-induced damage during pathogenesis. |
INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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; Eissenberg and Goldman, 1994). Subsequently, yeast cells use host phagocytic cells as vehicles to spread to multiple organs of the reticuloendothelial system such as the spleen, liver, lymph nodes, and bone marrow. In patients with disseminated disease, a variety of additional organs can be colonized (Eissenberg and Goldman, 1991).
As described above, naïve macrophages are susceptible to colonization by H. capsulatum. However, upon induction of a cell-mediated immune response, macrophages become activated and gain the ability to restrict H. capsulatum replication (Newman, 1999). The anti-Histoplasma activity of activated murine macrophages is dependent on production of .NO (Lane et al., 1994; Nakamura et al., 1994; Newman, 1999).
A number of studies indicate that .NO and related reactive species are important antimicrobial effectors produced by macrophages (for reviews on RNS and pathogenesis, see Fang, 1999, 2004; Nathan and Shiloh, 2000; Shiloh and Nathan, 2000; Missall et al., 2004). To produce .NO, macrophages induce the transcription of inducible nitric oxide synthase (NOS2, also known as iNOS). Whereas NOS2 mRNA levels rise moderately during initial infection, a much more significant induction occurs upon interferon (IFN)-
stimulation of macrophages by T-cells (Shiloh and Nathan, 2000). Nos2p converts arginine into L-citrulline and the free radical .NO. .NO readily combines with thiols, metals, and reactive oxygen species (such as superoxide, O2-·) to form multiple reactive species, collectively referred to as reactive nitrogen species (RNS) (Nathan and Shiloh, 2000; Fang, 2004). RNS react with many cellular components, including transition metals, lipids, thiol moieties, and DNA bases. These perturbations result in DNA and membrane damage, inhibition of replication and respiration, and inactivation of other cellular enzymes.
Whereas cell culture experiments suggest that .NO is required to restrict H. capsulatum replication in activated macrophages (Lane et al., 1994; Nakamura et al., 1994; Newman, 1999), the inhibitory effect of .NO on H. capsulatum is fungi-static rather than fungicidal (Nakamura et al., 1994). This result suggests that H. capsulatum is able to resist killing by .NO in host cells. Additionally, the immune system is not able to completely eliminate H. capsulatum; instead, yeast cells remain latent in the host for many years (Eissenberg and Goldman, 1991, 1994; Bullock, 1993). Because the ability of H. capsulatum to withstand nitrosative stress is likely to contribute to pathogenesis, the goal of this work was to uncover candidate genes that might contribute to the response of this organism to RNS. Because molecular genetic tools in H. capsulatum are still limited, we took a functional genomics approach to identify H. capsulatum genes that are induced in response to treatment with .NO donors. We identified the first set of candidate genes and genetic pathways that H. capsulatum may use to cope with nitrosative stress.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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) or on HMM agar plates supplemented with 5 mg/ml bovine serum albumin (BSA). H. capsulatum strain G217B ura5- (a kind gift of William Goldman; Woods and Goldman, 1992) was grown in HMM broth supplemented with 0.2 mg/ml uracil (Sigma-Aldrich, St. Louis, MO). After transformation with URA5-containing plasmids, G217B ura5- was grown in HMM broth or on HMM agarose plates with no uracil supplementation. H. capsulatum cultures were passaged every 2-3 d at 1:25 dilution. Liquid cultures were grown at 37°C under 5% CO2 on an orbital shaker. Plates were grown at 37°C under 5% CO2 in a humidified incubator.
Cell Collection for Microarray Analysis
For each microarray sample, 100 ml of cells were harvested by vacuum filtration, frozen in liquid nitrogen, and stored at -80°C until RNA preparation.
Dipropylenetriamine NONOate (DPTA NONOate) Dose-Response Experiments
DPTA NONOate (Cayman Chemical, Ann Arbor, MI) was resuspended in 4°C 10 mM NaOH to a final concentration of 1 M. The exact concentration of the stock solution was determined by measuring absorption at 252 nm. Because DPTA NONOate is very stable in alkaline solutions, nitric oxide release was not initiated until addition of the stock to HMM, pH 7.5.
The half-life of DPTA NONOate under our experimental conditions was determined using a protocol modified from Hrabie et al. (1993). A 1 M stock of DPTA NONOate/10 mM NaOH was diluted to a final concentration of 50 mM in HMM and placed in a 37°C incubator with 5% CO2. Spectrophotometric measurements at 252 nm were taken every 30 min for the first 2 h and then every 30-90 min for the next 6 h. A final A252 reading taken at 50 h was used as the A
measurement. ln(A - A) versus time was plotted to determine the rate constant for the reaction. According to the equation t1/2 = 0.6931 (1/k), where k equals the rate constant, the half-life under our experimental conditions was calculated to be 90 min.
Two-day-old H. capsulatum yeast cultures were diluted to 4 x 107 cells/ml and grown for 24 h until they reached mid-logarithmic phase. Two aliquots of cells representing untreated, zero-minute time points were harvested. The remaining cells were split into eight cultures. DPTA NONOate (1 M) resuspended in NaOH was added to six of the cultures to result in final concentrations of 0, 0.1, 0.5, 0.8 1, or 2 mM DPTA NONOate. Because the DPTA NONOate stock contains NaOH, we controlled for NaOH addition by supplementing the first five of these cultures with 10 mM NaOH to give a final concentration of 20 µM NaOH, which matches the NaOH concentration in the 2 mM DPTA NONOate sample. An equivalent volume of water was added to the seventh culture. To the last culture, 1 M DPTA NONOate was added to a final concentration of 5 mM DPTA NONOate/50 µM NaOH. Samples were harvested 2 h after treatment for subsequent RNA analysis. Spectrophotometric readings at 600 nm were taken in triplicate at 0, 2, 5, and 25 h after treatment for 0, 0.5, 1, and 2 mM DPTA NONOate. Spectrophotometric readings at 600 nm were taken for the remaining cultures at 0, 2, and 25 h after treatment. Serial dilutions of cells were prepared in triplicate for each culture at 0, 2, and 25 h after treatment. Dilutions were plated in duplicate on HMM agar plates containing bovine serum albumin (BSA). Colony forming units (CFUs) were assessed 12 d later.
DPTA NONOate Kinetic Time Course
DPTA NONOate (1 M) stock and cell cultures were prepared as described above. An aliquot of cells was harvested before the beginning of the time course. DPTA NONOate was added to a final concentration of 1.2 mM DPTA NONOate/12 µM NaOH. Samples were harvested 0.17, 0.5, 1, 2, 5, and 25 h after treatment. A mock treatment time course was performed after addition of 12 µM NaOH.
S-Nitrosoglutathione (GSNO) Dose-Response Experiment
Two-day-old H. capsulatum yeast cultures were diluted to 3 x 107 cells/ml and grown for 24 h until they reached mid-logarithmic phase. Two aliquots of cells representing untreated, zero-minute time points were harvested. The remaining cells were split into five cultures. GSNO (Axxora Life Sciences, San Diego, CA) resuspended in HMM was added to four of the cultures to result in final concentrations of 0.2, 0.4, 1, or 5 mM GSNO. As a control, an equivalent volume of HMM was added to the fifth culture. Spectrophotometric readings at 600 nm were taken in triplicate at 0, 2, 5, 10, and 25 h after GSNO addition. Serial dilutions of cells were prepared in triplicate for each culture at 0, 2, and 25 h after treatment. Dilutions were plated in duplicate on HMM agar plates containing BSA. CFUs were assessed 12 d later.
Environmental Stress Experiments
For each environmental stress experiment H. capsulatum was diluted to 1.8 x 107 cells/ml and grown for 48 h. An aliquot of cells was harvested before treatment, and then each time course was initiated by the addition of diamide, menadione, dithiothreitol (DTT), or sorbitol. For the diamide time course, 1 M diamide (Sigma-Aldrich) was added to the culture to result in a final concentration of 1.5 mM. Samples were removed at 10, 20, 30, 40, 60, and 90 min after treatment. For the menadione time course, 1 M menadione (Sigma-Aldrich) was added to the culture to result in a final concentration of 1 mM. Samples were removed at 10, 20, 30, 50, 80, and 340 min after treatment. For the DTT time course, 1 M DTT (Roche Diagnostics, Indianapolis, IN) was added to the culture to result in a final concentration of 2.5 mM. Samples were removed at 30, 60, 120, 240, and 480 min after treatment. For the sorbitol time course, an equal volume of 2 M sorbitol in HMM (Sigma-Aldrich) was added to the culture, resulting in a final concentration of 1 M sorbitol. Samples were removed at 5, 15, 30, and 60 min after treatment.
Log Phase Time Course
H. capsulatum was diluted to 1.8 x 107 cells/ml and grown for 48 h. Samples were harvested at 0, 15, 30, 60, 180, 240, 480 min, and 24 h after treatment.
Cell Lysis and RNA Isolation
Total RNA was isolated using a guanidine thiocyanate lysis protocol (Hwang et al., 2003). Poly-adenylated (poly-A) RNA was isolated from total RNA using an Oligotex mRNA kit (QIAGEN, Valencia, CA) or an oligonucleotide-dT cellulose (Ambion, Austin, TX) column. The 2 mM DPTA NONOate total RNA sample from dose-response experiment 2 was lost during manipulations.
Probe Preparation, Microarray Hybridization, and Data Acquisition for Shotgun Genomic Microarray
Fluorescently labeled cDNA was made by incorporating amino-allyl dUTP during reverse transcription of poly-A RNA with oligonucleotide-dT and random hexamers. Cy5 or Cy3 dyes (GE Healthcare, Piscataway, NJ) were coupled to the amino-allyl group as described previously (DeRisi et al., 1997). For each sample, the cDNA was coupled to Cy5. All poly-A RNAs from a given time course or dose-response experiment were combined to prepare a reference cDNA pool, which was coupled to Cy3. A Cy5-labeled cDNA probe was mixed with its corresponding Cy3-labeled cDNA reference and competitively hybridized on a shotgun genomic microarray. Arrays were scanned using either a GenePix 4000A or GenePix 4000B scanner (Axon Instruments/Molecular Devices, Union City, CA).
Shotgun Genomic Microarray Data Analysis
Arrays were analyzed using GenePix PRO version 3.0 (Axon Instruments/Molecular Devices), NOMAD (http://derisilab5.ucsf.edu/NOMAD), CLUSTER (Eisen et al., 1998), and Java Treeview 1.0.8 (available at http://sourceforge.net/project/showfiles.php?group_id=84593). Each spot on the microarray corresponds to a random genomic fragment that will be referred to hereafter as a microarray clone. To eliminate clones with low signal, we did not analyze clones for which the sum of the medians for the 635- and 532-nm channels was 
500 intensity units. Because all samples were hybridized against reference pools, each ratio measurement was normalized relative to its respective untreated zero-minute time point by dividing the expression ratio for each clone in a sample by the expression ratio in its respective untreated zero-minute time point. For each clone in each dose-response experiment, the mean of the expression ratios from the duplicate untreated, zero-minute samples was calculated and used for the normalization described here.
Cluster analysis was performed on the two DPTA NONOate experiments and one GSNO dose-response experiment. We identified 714 induced clones and 964 repressed clones as 4-fold changed in at least one sample (compared with the zero-minute sample) and present in 80% of the samples. Further analysis was performed on clones that were induced in nitrosative stress. 678/718 clones were induced 2-fold in at least two dose-response experiments compared with their zero-minute sample. Another 17 clones were induced 4-fold in at least one dose-response sample versus the zero-minute sample but had data for <80% of the samples. The resultant set of 695 RNS-induced clones is shown in Figure 2.
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Sequence reads were aligned to the contigs of the H. capsulatum genome sequencing project (Genome Sequencing Center, http://genome.wustl.edu/projects/hcapsulatum) using BLASTN (Altschul et al., 1997). Alignments were required to have a maximum E-value of 1E-99 or high identity (>95%) over their length. Additional alignments were accepted upon manual curation. For clones with forward and reverse reads, the contig sequence between and including the sequence reads was designated the microarray clone sequence. Because the average size of the microarray clones was 1-2 kb (Hwang et al., 2003), the sequence of clones with only one end read was defined as the contig sequence beginning with the sequence read and continuing for 1 kb.
Some sequence reads aligned to multiple locations in the genome and were likely to represent repetitive sequence. All clone sequence reads were compared with a library of H. capsulatum repetitive sequences (provided by the Genome Sequencing Center) using BLASTN. Sequence reads with minimum alignment lengths of 100 base pairs and 80% identity were designated repetitive sequence. BLASTN was used to determine whether clones that could be located in the genome corresponded to transposon-related or rDNA sequences (Hwang et al., 2003). Of the 695 RNS-induced clones, 356 contained sequences identified as repetitive or transposon-related and three contained rDNA sequence.
In addition, we removed another 54 clones from further analysis. Twenty-one of these clones lacked sequence information, did not align to the genome sequence, or aligned to the genome sequence at more than one location but did not correspond to transposons/repeat sequences. The other 33 clones in this group were not pursued further because they clustered with the transposon/repeat clusters described in the text. These clusters showed induction in response to multiple stresses and therefore did not seem likely to represent a specific response to RNS.
Further annotation was performed on the remaining 282 array clones. Contigs with mapped array clones were subjected to BLASTX analysis against the nonredundant protein database (nr) maintained by the National Center for Biotechnology Information. As described in the Results, a total of 41 array clones could be unambiguously annotated. The remaining 241 clones were located in the genome sequence but could not be unambiguously annotated by BLASTX analysis for reasons described in the Results. The majority of these clones were pursued further as described below.
To determine how many of the 282 clones were specifically induced in RNS, the fold change of clones in each experiment was evaluated. Clones were designated as induced in RNS if they were up-regulated 4-fold in at least one RNS-treated sample compared with the zero-minute sample. Clones were designated as not induced in stress if they were <2-fold in all stress experiments (diamide, menadione, sorbitol, and DTT) and RNS controls (untreated dose-response 1 and 2, NaOH-treated dose-response 1 and 2, untreated GSNO dose response, and 12 µM NaOH time course). In addition, clones had to have data for 80% of the stress experiments to be considered in this analysis. Fifty-nine clones were up-regulated 4-fold in RNS (compared with the zero-minute sample) and <2-fold in stresses/controls. These were designated specifically induced by RNS.
Tiling Microarray Construction
Of the 241 unannotated clones described above, 198 were chosen for tiling analysis using high-density oligonucleotide tiling arrays (CombiMatrix, Mukilteo, WA). (All 241 could not be included because of space restrictions on the tiling arrays.) An additional 19 clones that had already been annotated were included as controls to verify that the tiling arrays identified RNS-induced transcripts. This brought the total number of tiled clones to 217.
Because some of the clones represented overlapping genomic regions, the 217 clones corresponded to 105 genomic regions. In addition, 19 genomic regions also were tiled for which we had additional data suggesting up-regulation in response to RNS, bringing the total number of tiled genomic regions to 124. The boundaries of each tiled region were chosen according to criteria described in the Results. The sequence of each tile was selected by CombiMatrix. The tiling oligonucleotides were between 20 and 50 base pairs in length. The oligonucleotide length was varied to minimize the range of melting temperatures; 99% of the tiles were separated by 9-10 base pairs relative to the contig sequence.
Tiling Microarray Probe Preparation, Hybridization, and Data Acquisition
Fluorescently labeled probes were generated as described previously with one modification; amino-modified C6 5'-labeled oligonucleotide-dT primer was used for reverse transcription. CombiMatrix microarrays were washed before use as described in the "shampoo" protocol (http://derisilab.ucsf.edu/core/resources/index.html). Cy5-coupled 0.8 mM or 0.5 mM DPTA NONOate (2-h time point) samples were competitively hybridized against Cy3-coupled NaOH (2-h time point) samples as described previously (DeRisi et al., 1997). Dose-response 1 samples were used for this experiment. Cy5-coupled 5 mM GSNO (2-h time point) samples were competitively hybridized against Cy3-coupled 0 mM GSNO (2-h time point) samples. Microarrays were hybridized 12-14 h at 55°C with constant orbital rotation on a Nutator (Clay Adams Brand, Fisher Scientific, Pittsburgh, PA).
Arrays were scanned using a GenePix 4000B scanner. To normalize the 635- and 532-nm signal, the 635- and 532-nm PMT levels were adjusted to give equivalent background signal from hybridization to the CombiMatrix quality control oligonucleotide QC-oligoB-AS-3, present 200 times on the array. Because QC-oligoB-AS-3 does not align to the H. capsulatum genome or mitochondrial sequence, binding by sample probes represents background and should be equivalent between the Cy3 and Cy5 probes. The data sets were analyzed using GenePix PRO version 3.0. The mean of the 635- and 532-nm median signal intensities for the 200 QC-oligoB-AS-3 spots was considered background and subtracted from the 635- and 532-nm median signal intensities, respectively.
Tiling Microarray Transcript Identification and Annotation
The 635- and 532-nm signal intensities were plotted as a function of tile position using the Generic Genome Browser (Gbrowse, http://www.gmod.org). Transcripts were chosen manually according to the following criteria: 1) at least five contiguous oligonucleotides with a minimum 635-nm intensity of 750 intensity units and 2) at least one tile with 635-nm intensity >1000 intensity units. Transcripts that seemed to have at least twofold higher 635-nm signal intensity verses 532-nm signal intensity were designated RNS-induced. We identified 135 RNS-induced transcripts from the 124 tiled genomic regions because some regions contained more than one RNS-induced gene. Transcripts that had <2-fold RNS induction were designated constitutive and not analyzed further. No transcripts were identified with higher 532- verses 635-nm signal intensity. One transcript was induced only in GSNO but not in DPTA NONOate, whereas three transcripts showed much stronger induction in DPTA NONOate than in GSNO. Some regions showed no induced transcripts, and microarray clones corresponding to these regions were annotated as "No Induction." The GBrowse pages showing the tiling results, identified transcripts, and their Aspergillus nidulans, Saccharomyces cerevisiae, and Schizosaccharamyces pombe homologues can be found in the Supplemental Materials.
We defined background signal intensity as the median signal intensity of all tiles outside the constitutive and RNS-induced transcript areas. On each of the arrays, the background signal was <300 intensity units, indicating that our signal cutoff of 750 was sufficiently above background for transcript identification.
RNS-induced transcripts were aligned by BLASTX to nr and the following predicted gene sets: A. nidulans (Broad Institute, Oct. 2003), Magnaporthe grisea (Broad Institute, Oct. 2003), Fusarium graminearum (Broad Institute, Sep. 2003), Neurospora crassa (Munich Information Center for Protein Sequences, June 2004), Cryptococcus neoformans (National Center for Biotechnology Information GenBank, Feb. 2005), Candida albicans (Biotechnology Research Institute/National Research Council Canada, Dec. 2004), S. cerevisiae (Saccharomyces Genome Database/National Center for Biotechnology Information GenBank, Mar. 2004), S. pombe (Sanger Institute, Mar. 2005), Yarrowia lipolytica (Genolevures, May 2004), and Phanerochaete chrysosporium (DOE Joint Genome Institute, Mar. 2004). Only alignments that corresponded to the correct strand, extended across 75% of the query length, and had a maximum E-value of E-7 were designated homologues. Two exceptions to these criteria are noted in the Results.
Oligonucleotides
All oligonucleotides used for standard molecular biology manipulations are listed in Supplemental Table 4.
Northern Analysis
Total RNA (5 µg) from DPTA NONOate dose-response 2 samples was used for Northern analysis (Hwang et al., 2003). NOR1 and ACT1 probes were PCR amplified from genomic DNA using gene-specific primers.
NOR1 5' Rapid Amplification of cDNA Ends (RACE), 3' RACE, and cDNA Sequencing
Poly-A RNA from 0-h untreated and 2-h 0.1 mM DPTA NONOate-treated cells (dose-response 1) was used to map the NOR1 transcript ends and coding sequence. The 5' and 3' ends of cDNAs were determined using the First-Choice RNA-ligase mediated (RLM)-RACE kit (Ambion, Austin, TX) per manufacturer's instructions. cDNA reverse-transcribed from NONOate treated cells (dose-response 1) was used as a template to amplify the NOR1 coding sequence (cds). All amplified products were cloned using TOPO-TA (Stratagene, La Jolla, CA) and sequenced using a combination of M13-forward, M13-reverse, and gene-specific primers as necessary.
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To construct the Gateway destination vector, a polylinker was introduced into the BamHI site of H. capsulatum plasmid pWU55 (Woods et al., 1998) creating pDG29AS. To replace the Podospora anserina URA5 with H. capsulatum URA5, pDG29AS was first enzymatically digested with EcoRI, and the ends of the larger fragment were filled using Klenow. A plasmid containing H. capsulatum URA5 was created by PCR amplifying H. capsulatum URA5 from G217B genomic DNA, cloning the amplified product into TOPO-TA, and sequencing the product as described above. This plasmid was digested with the blunt cutting enzyme, HpaI, and the resulting URA5 fragment was ligated with the prepared pDG29AS blunt-ended backbone creating pDG31AS. Finally, the Gateway destination vector pDG33 was created by ligating the Gateway vector conversion cassette reading frame A into HpaI-digested pDG31AS.
Gateway recombination of pDHM1 with pDG33 resulted in pDHM5, an H. capsulatum plasmid containing CRP1 promoter-NOR1 cds-CATB terminator, H. capsulatum URA5, and telomeric repeats flanking a kanamycin resistance cassette. Gateway recombination of pCC4 with pDG33 yielded pDG45, the vector control, lacking the NOR1 cds. pDHM5 and pDG45 were digested with PacI to expose the telomeric ends and transformed into G217B ura5- after gel purification. Prototrophic colonies were selected on HMM agar plates.
For ectopic expression experiments, 2-d cultures of G217B ura5- transformed with pDHM5 or pDG45 were diluted to 1.6 and 1.7 x 107 cells/ml, respectively. The cultures were grown for 24 h until they were in mid-logarithmic phase. Copper sulfate (10 mM; Fisher Scientific) was added to each culture to a final concentration of 10 µM. After 1.5 h, each culture was split into three. DPTA NONOate (1 M), prepared as described previously, and 10 mM NaOH were added to each culture to achieve a final concentration of 0, 1, or 2 mM DPTA NONOate/20 µM NaOH. Spectrophotometric readings at 600 nm were taken in triplicate 0 and 24 h after DPTA NONOate treatment.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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To examine the transcriptional profile of H. capsulatum in response to nitrosative stress, we used a shotgun genomic microarray. In brief, each of the 10,000 spots on the shotgun genomic microarray corresponds to a random fragment from the H. capsulatum genome. This array has been used to perform large-scale analysis of transcript abundance in the absence of a fully sequenced and annotated genome (Hwang et al., 2003). To determine the transcriptional response to nitrosative stress, poly-adenylated RNA isolated from each RNS-treated sample was subjected to microarray analysis and compared with controls as described in Materials and Methods.
Cluster analysis (Eisen et al., 1998), which groups array clones with common gene expression profiles, was used to compare the expression profiles observed in three RNS dose-response experiments (two DPTA NONOate experiments and one GSNO experiment; Figure 2). Overall, 714 clones were induced 4-fold in at least one dose-response sample compared with their relevant zero-minute sample, and 964 clones were repressed 4-fold in at least one dose-response sample compared with their relevant zero-minute sample. A comparison of the two DPTA NONOate dose-response experiments showed that a similar set of genes was induced in both experiments but not with the same dose dependence. Interestingly, no cluster was observed to be specific to a particular nitrosative stress, and in fact, the intensity of induction in GSNO was very similar to the second DPTA NONOate dose-response experiment. Subsequent analysis focused on array clones that were induced in at least two of the three RNS dose-response experiments compared with their zero-minute samples (695 clones). Because neither the shotgun microarray nor the H. capsulatum genome was annotated at the time of this work, it took considerable effort to determine the identity of transcripts that corresponded to each regulated clone from the microarray. Thus, we chose to focus on annotating only the induced (but not the repressed) clones, because these are the most likely to represent genes necessary for RNS response or resistance.
Annotation of Array Clones
The next step of our analysis was to determine which genes were represented by the 695 clones on the shotgun genomic microarray. Complete details of how the microarray clones were annotated can be found in Materials and Methods. To summarize, single-sequence reads that marked the boundaries of the genomic insert in each array clone were mapped onto contigs from the ongoing H. capsulatum genome sequencing project at the Washington University in St. Louis Genome Sequencing Center (http://genome.wustl.edu/projects/hcapsulatum). BLASTX analysis of the GSC contigs against the nr peptide sequence database maintained by the National Center for Biotechnology Information allowed us to locate homologues on the contig sequences. We determined the overlap between the location of microarray clones of interest and nr homologues relative to the genome sequence. Based on information from sequenced fungal genomes, we hypothesized that the minimum upstream regulatory sequence in H. capsulatum is 300 base pairs, and the minimum 3'-untranslated sequence is 200 base pairs. Therefore, if a microarray clone contained either >300 base pairs 5' of a region of nr homology or >200 base pairs 3' of a region of nr homology, it had the potential to overlap a second gene. Forty-one clones were unambiguously annotated because 1) the microarray clone and nr homologue completely overlapped or 2) the microarray clone and nr homologue did not completely overlap, but the remaining clone sequence was of insufficient size to contain a second gene. Two hundred seventy-four clones could not be annotated because 1) the microarray clone overlapped more than one nr homologue, 2) the microarray clone had the potential to overlap more than one nr homologue, or 3) the microarray clone did not overlap any nr homologues. In addition to the 41 clones mentioned above, 356 microarray clones were annotated as transposons/repeats because they aligned with either previously identified transposon-related sequences (Hwang et al., 2003) or H. capsulatum repeat families identified by the GSC. Another three microarray clones were annotated as rDNA because they aligned with previously identified rDNA sequence (Hwang et al., 2003). The remaining 21 clones could not be analyzed because they either lacked sequence information or could not be uniquely located in the genome sequence.
Comparison of Gene Expression Profiles of RNS-induced Clones in Other Stresses
Before proceeding with further annotation, we determined which clones were induced specifically in nitrosative stress. Work in S. cerevisiae and S. pombe demonstrated that a common set of genes changes in expression in response to all stresses. In contrast to this general environmental stress response, individual stresses also trigger a specific gene expression response (Gasch et al., 2000; Causton et al., 2001; Chen et al., 2003). To categorize the RNS-induced clones into those 1) induced in all stresses, 2) induced in a subset of stresses, and 3) induced specifically in nitrosative stress, we examined the transcriptional profiles of these clones in response to other stresses. To generate these data, H. capsulatum was exposed to a superoxide generator (menadione), a sulfhydryl oxidant (diamide), a disulfide reductant (DTT), and osmotic shock (sorbitol). Doses and time courses were chosen to match previous studies in S. cerevisiae (Gasch et al., 2000). To identify clones that change in expression during and after exponential growth, we determined the gene expression profile of log phase (this study) and stationary phase cultures (Hwang et al., 2003). In addition, a time course of 1 mM DPTA NONOate treatment (or mock treatment with 12 µM NaOH) consisting of 0-, 0.5-, 1-, 2-, 5-, and 25-h time points was performed to evaluate the kinetic effects of nitrosative stress.
Cluster analysis was used to compare RNS-induced array clones across all of the stresses (Figure 2 and Supplemental Table 1). The section of the cluster labeled "E" indicates 395 clones induced in all stresses as well as in response to RNS. We had previously annotated 356 induced clones as repeats/transposons, as described above. Three hundred and fifty of these clones are located in region E, constituting 89% (350/395) of the clones in this area. The induction of these clones in multiple stresses is interesting, but not surprising. Work in S. cerevisiae has shown that retrotransposons are induced when cells are stressed by nitrogen starvation or DNA-damaging agents (Bradshaw and McEntee, 1989; Morillon et al., 2000). The remainder of the clones in section E (33) were not annotated because these clones are not likely to represent pathways used by H. capsulatum to cope specifically with RNS-induced damage.
For our subsequent analysis, we focused on the remaining 282 clones that could be located in the genome sequence and were not repeats/transposons or rDNA. The majority of these clones are induced in RNS and minor subsets of the other stresses. Fifty-nine clones are induced only in RNS, but they do not cluster together into one group. The simplest interpretation of these data is that multiple transcriptional circuits are activated by nitrosative stress, although transcriptional profiling with a whole-genome array is necessary to definitively test this hypothesis.
Identification of RNS-induced H. capsulatum Transcripts Using Tiling Arrays
The BLASTX analysis described above allowed us to annotate only 41 of the 282 clones of interest. The remaining 241 clones defined genomic regions of interest that overlapped with an RNS-induced transcript of unknown identity. Because H. capsulatum has numerous small introns, the location of genes cannot be determined simply by finding open reading frames in the vicinity of microarray clones. Consequently, we used alternative methods to map RNS-induced transcripts.
We determined the precise location and coding strand of the remaining RNS-induced transcripts using high-density oligonucleotide tiling microarrays. Work in Arabidopsis thaliana, Escherichia coli, and Homo sapiens has demonstrated the utility of tiling arrays to map transcript locations (Shoemaker et al., 2001; Tjaden et al., 2002; Yamada et al., 2003; Schadt et al., 2004). In this type of array, both strands of a region of interest are represented on a microarray by a series of end-to-end oligonucleotides. Labeled cDNA probes representing the sequence of the 5' untranslated region (UTR), 3' UTR, and exons will bind to their complementary oligonucleotides on the microarray and generate an increase in intensity at these targets. In contrast, intron sequences and non-UTR intergenic sequences are not present in the cDNA probe, and thus oligonucleotides representing these areas will not bind the cDNA probes. Therefore, by hybridizing labeled cDNA probes to a tiling array, the boundaries of a transcript and its coding strand can be determined.
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). For this region, we tiled 1 kb upstream and downstream of the defined coding sequence (Figure 3G). In total, 540.85 kb of genomic DNA were tiled, requiring 21,536 tiles. As described in Materials and Methods, the tiling arrays represented 217 microarray clones and 124 genomic regions. Labeled cDNAs from existing RNS-treated samples were hybridized to the tiling microarrays. In each case, the RNS-treated sample was labeled with Cy5, and the corresponding uninduced sample was labeled with Cy3. The median intensity of the 635-nm channel (red) and 532-nm channel (green) was plotted as a function of oligonucleotide position for the plus and minus strands. To verify that the arrays were correctly and accurately identifying transcribed regions, the signal from the AOX1 control region was examined. The transcript boundaries predicted by the tiling data matched the published sequence of the longest AOX1 transcript remarkably well (Figure 3G). Figure 3, B, D, and F, show specific examples of data obtained for the categories of clones defined in Figure 3, A, C, and E, respectively. These data confirmed that H. capsulatum RNS-induced transcripts could be identified using the tiling microarrays. Intensity graphs for each of the defined transcripts are shown in Supplemental Materials.
Each of the tiled regions was examined for RNS-induced transcripts. We chose a minimum signal cutoff of 750 intensity units to qualify a tile as potentially representing part of a transcript because the mean intensity of tiles on the noncoding strand and intergenic regions was <300 intensity units in each of the arrays examined. Because we observed rare isolated oligonucleotides that gave a signal intensity >750 intensity units, we designated genomic regions as transcripts if they had 1) at least five contiguous oligonucleotides with a minimum 635-nm intensity of 750 and 2) at least one tile with intensity >1000. The first base pair of the 5'-induced tiling oligonucleotide and the last base pair of the 3'-induced tiling oligonucleotide were arbitrarily designated as the transcript boundaries, although the actual ends of the transcript are likely to fall within the relevant tiling oligonucleotide. In some cases, the signal intensity did not decline to baseline at the ends of the tiled region, indicating that the tiled region did not contain the entire transcript (Figure 3B). These transcripts are indicated as "partial" in Table 1 and Supplemental Tables 2 and 3. In each case, the strand of the induced transcript also was noted. Transcripts that were expressed at equivalent levels in the RNS-treated and untreated samples were not pursued. Only 14 tiled regions did not contain any transcripts by our criteria.
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In sum, a total of 153 RNS-induced transcripts were identified using the combination of shotgun genomic and tiling microarrays described above. We named these transcripts and submitted them to GenBank.
Annotation of RNS-induced Transcripts
For each of the identified transcripts, homologues were sought that would provide insight into gene function. BLASTX analysis was conducted on each identified transcript against the nr peptide database and predicted gene sets from a variety of fungal species (A. nidulans, M. grisea, F. graminearum, N. crassa, S. pombe, S. cerevisiae, C. albicans, C. neoformans, Y. lipolytica, and P. chrysosporium). A transcript was annotated as having a homologue if 1) the H. capsulatum transcript extended across 75% of the homologue's length, 2) the E-value was less than E-7 (with 2 exceptions for homologues with sequences <100 amino acids), and 3) the transcript was on the same strand as the homologue. Fifty-three percent of the identified transcripts had a named homologue in another organism, 25% were homologous to "hypothetical" proteins in another organism, and 22% had no identifiable homologues in any organism (Table 1 and Supplemental Tables 2 and 3). For the 149 previously unidentified H. capsulatum genes, we named them based on their putative homologue in another organism. In cases where this nomenclature scheme was not applicable, we named genes nitrosative stress-induced transcript (NIT). All annotations were applied to the cluster comparing RNS-induced clones to other environmental stresses (Figure 2)
Clusters of Induced Clones Suggest Particular Pathways Are Triggered by Exposure to RNS
The ultimate goal of this work is to identify genes and pathways that play an important role in the ability of H. capsulatum to respond to nitrosative stress. Genes that function in the same pathway are often transcriptionally coregulated and thus may be grouped together by Cluster analysis of transcriptional data over a series of conditions (Hughes et al., 2000). Because we used a shotgun genomic microarray that did not cover the entire genome, we did not have all genes represented on the microarray, and therefore, we could not generate all relevant clusters. Thus, some genes of interest clearly fall into clusters, whereas others do not. Here, we describe both RNS-induced pathways as defined by groups of genes with similar function that cluster together as well as individual genes that do not fall into obvious clusters at this time.
Iron Acquisition
Cluster A, shown in Figures 2 and 4, has homologues of several genes involved in iron acquisition in other fungi. This cluster contains a homologue of L-ornithine monooxygenase (H.c. LOM1; Hwang and Rine, personal communication), an enzyme that performs the first committed step in siderophore production. Siderophores are small molecules secreted by microbes to scavenge iron from the environment. In Ustilago maydis, the L-ornithine monooxygenase and nonribosomal peptide synthetase required for siderophore production are grouped together in the genome (Yuan et al., 2001). Our iron acquisition cluster also contains a nonribosomal peptide synthetase (H.c. NPS1), which, similar to U. maydis, is grouped together in the genome with H.c. LOM1. Two other genes of unknown function from this cluster are adjacent to NPS1 and LOM1 in the genome. The cluster analysis indicates these neighboring genes are coregulated with LOM1 and NPS1, suggesting that they may function in siderophore production as well. This iron acquisition cluster also includes a putative ferric chelate reductase (H.c. NIT1) that may function either to remove iron from siderophores after uptake of siderophore/Fe3+ complexes, or to reduce iron to Fe2+ outside of the cell before iron import (De Luca and Wood, 2000). H.c. STR1, a homologue of a siderophore iron transporter from S. pombe, also is induced upon RNS exposure but is present in a different cluster (Table 1).
Up-regulation of iron acquisition genes in response to nitrosative stress has been observed in a number of organisms (Crawford and Goldberg, 1998; Mukhopadhyay et al., 2004; Hromatka et al., 2005; Sarver and DeRisi, 2005). In E. coli, .NO inactivates the transcription factor Fur, which normally represses iron acquisition genes under high iron conditions. Thus, in nitrosative stress, Fur-regulated repression is relieved, and iron acquisition genes are induced (D'Autreaux et al., 2002; Mukhopadhyay et al., 2004). Induction of H. capsulatum iron acquisition genes may reflect .NO-induced modification of an unidentified iron-regulated transcriptional repressor.
Energy Production
Genes involved in energy production also are induced upon RNS exposure as seen in cluster B (Figures 2 and 5. Reactive nitrogen species are known to inhibit components of the electron transport chain [complexes I, II, III, and IV] [Brown, 1999]). Exposure to RNS may trigger up-regulation of genes that encode the components themselves or genes that allow bypass of standard complexes of the electron transport chain. We observed induction of homologues of cytochrome c oxidase subunit VIb (H.c. COX12) and cytochrome c (H.c. CYC1), which are standard components of the electron transport chain used to produce ATP for cellular processes. We also observed induction of a protein homologous to a mitochondrial succinate-fumarate transporter (H.c. SFC1), which, in other organisms, transports succinate into the mitochondrial matrix either for use by the tricarboxylic acid (TCA) cycle or to donate electrons to complex II (Palmieri et al., 1997; Joseph-Horne et al., 2001). A phosphate transporter homologue (H.c. MCP2) also was induced in response to RNS. In other organisms, this protein transports phosphate into the mitochondrial matrix for use in ATP production by the ATP synthase that is coupled to the electron transport chain (Laloi, 1999). Alternative components of the electron transport chain, such as alternative oxidase (H.c. AOX1), were also up-regulated. In other organisms, Aox1p is able to perform the terminal reduction of O2 to water and bypass complexes III and IV of the electron transport chain (Joseph-Horne et al., 2001). Unlike complex IV, Aox1p is refractory to inhibition by .NO; thus, its up-regulation may help to ensure that energy production can continue in the presence of .NO (Joseph-Horne et al., 2001). Alternatively, induction of Aox1p may relieve the oxidative stress associated with inhibition of complex IV (Joseph-Horne et al., 2001). Additionally, a homologue of alternative NADH dehydrogenase (H.c. NDH1) is also up-regulated upon RNS exposure but is found in a different cluster. In other organisms, this protein facilitates transfer of electrons from NADH to ubiquinone, the shuttle protein between complexes I and III (Joseph-Horne et al., 2001), thus allowing electrons from NADH to bypass complex I and enter the electron transport chain. In addition to maintaining electron flow, this process regenerates oxidized NAD+. As long as sufficient quantities of NAD+ exist, energy could continue to be produced via glycolysis in the absence of a functioning electron transport chain.
Other RNS-induced genes outside this cluster also may function in energy production. For example, several genes function in the metabolism and oxidation of lipids. Homologues of genes involved in fatty acid transport and activation (H.c. FAT1 and H.c. FAA1) as well as a gene involved in fatty acid 
-oxidation (H.c. FOX2) are up-regulated (Table 1) (van Roermund et al., 2003). Fatty acid -oxidation produces acetyl-CoA, which can be used in the TCA cycle directly or in the glyoxylate pathway to produce succinate. Succinate can be transported into the mitochondria, possibly via H.c. SFC1 (see above), where it can be used in the TCA cycle or the electron transport chain. Because the -oxidation of fatty acids represents the highest energy-yielding pathway in the cell, up-regulation of these genes may reflect alternate pathways of energy production induced by the cell during nitrosative stress.
Stress Response
A small cluster of clones all corresponding to a homologue of the S. pombe transcription factor atf1 is shown in cluster C (Figures 2 and 6). S. pombe Atf1p is the primary transcription factor downstream of the Sty1 stress response pathway (Chen et al., 2003). This pathway operates through Wis1p, a mitogen-activated protein kinase kinase. We also observed that the H. capsulatum homologue of WIS1 is induced by RNS (Figure 5). In S. pombe, this pathway responds to a variety of stresses, including DNA damage, heat shock, oxidative stress, and high osmolarity (Hohmann, 2002). Atf1p induces transcription of a significant fraction of the core environmental stress response in S. pombe (Chen et al., 2003). We observed induction of H. capsulatum ATF1 in response to all stresses tested, including RNS exposure (Figure 6). This uniform pattern of induction suggests that the H. capsulatum Atf1p may function in a core environmental stress response.
Protein Folding and Degradation
A gene cluster with many homologues of genes involved in protein folding and degradation is also induced in response to RNS (cluster D; Figures 2 and 7). We have identified homologues of S. cerevisiae CDC37, STI1, YDJ1, and YME1 as well as the Arabidopsis thaliana UFD1-like protein. S. cerevisiae Cdc37p, Sti1p, and Ydj1p are known to associate with heat-shock protein (Hsp)70p and Hsp90p to assist protein folding (Cyr et al., 1994; Bohen et al., 1995; Lee et al., 2004). We also observed induction of an Hsp70 family member (H.c. HSP70) in response to nitrosative stress, although this gene was located in a different cluster. S. cerevisiae Ydj1p also is involved in endoplasmic reticulum (ER)-associated degradation (ERAD), the process whereby misfolded proteins in the ER are removed to the cytoplasm for degradation by the proteasome (Huyer et al., 2004; Youker et al., 2004). Whereas the function of the A. thaliana UFD1-like protein has not been elucidated, S. cerevisiae Ufd1p also participates in ERAD by recognizing polyubiquitinated proteins and targeting them to the proteasome (Ye et al., 2001; Bays and Hampton, 2002). S. cerevisiae Yme1p functions in mitochondrial degradation of misfolded proteins (Nakai et al., 1995). Homologues of other RNS-induced genes outside this cluster also seem to be involved in proteasomal degradation (Table 1). For example, the homologue of H.c. CUT8 targets proteins to the proteasome for degradation in S. pombe (Tatebe and Yanagida, 2000). We also observed induction of ubiquitin (H.c. UBI1) and a ubiquitin-conjugating enzyme (H.c. UBC1) in response to RNS. Interestingly, the protein degradation cluster shows little induction in response to stresses other than RNS with the only exception being diamide, a stress known to induce protein unfolding (Figure 7). Thus, induction of these genes is relatively specific to RNS exposure in H. capsulatum, and likely reflects the cell's attempt to deal with unfolded proteins generated by nitrosative stress.
Regulation of RNS-induced Genes in Other Stresses
Large-scale gene expression profiling over a variety of conditions allowed us to identify clusters of genes that were induced in nitrosative stress, as described above. In addition, we also were able to examine the response of these clusters under other stresses. Notably, two of the RNS-response clusters, iron acquisition and energy production, were down-regulated upon oxidative treatment (Figures 4 and 5). This transcriptional regulation is consistent with the known roles these proteins play in the production of oxidative species. Iron produces reactive oxygen species (ROS) by reaction with oxygen and hydrogen peroxide (De Luca and Wood, 2000), and respiration produces low concentrations of the ROS superoxide (Liu, 1997). Down-regulation of these two classes of genes under oxidative stress may help H. capsulatum avoid damage by limiting ROS production.
Homologues of Known RNS-Response or Stress-Response Genes
Some of the interesting RNS-response genes did not fall into specific clusters, yet still had intriguing connections to nitrosative stress response, as described below. Additionally, novel genes and genes with homologues of unknown function also were identified. Table 1 lists all of the identified RNS-induced transcripts that have named homologues. These transcripts are grouped according to the function of their homologous gene. RNS-induced transcripts homologous to "hypothetical" genes in other organisms are listed in Supplemental Table 2, and novel RNS-induced transcripts (with no homologues) are listed in Supplemental Table 3.
Some of the transcripts in Table 1 have homologues that have been observed to be necessary for or up-regulated during RNS response in other organisms. For example, the C. albicans transcriptional response to RNS stress includes up-regulation of a copper transporting P-type ATPase and an alternative oxidase (Hromatka et al., 2005), which we also observed. A copper transporting P-type ATPase also was induced in the E. coli response to RNS (Mukhopadhyay et al., 2004). In addition, the C. albicans as well as the S. cerevisiae RNS-induced transcriptional response includes up-regulation of a catalase (Hromatka et al.; 2005; Sarver and DeRisi, 2005). Screens in Mycobacterium tuberculosis for mutants unable to grow after RNS exposure identified proteins involved in proteasomal degradation (Darwin et al., 2003). Moreover, treatment of M. tuberculosis with inhibitors of the proteasome resulted in increased susceptibility to RNS, suggesting that protein degradation is necessary for the response of M. tuberculosis to nitrosative stress. Finally, the mammalian homologue of H.c. DBP2, p68, was strongly induced in human and murine keratinocytes upon .NO stimulation (Kahlina et al., 2004).
Homologues of other genes in Table 1 have intriguing connections to RNS stress. First, response to DNA damage is likely to be a key element of response to nitrosative stress. We observed that homologues of S. cerevisiae WSS1 (H.c. NIT2) and YNG2 (H.c. YNG2) were induced in response to RNS. In S. cerevisiae, disruption of these genes results in increased sensitivity to DNA damage, suggesting they have a role in DNA repair (Choy and Kron, 2002; O'Neill et al., 2004). A homologue of U. maydis Hmp1 (H.c. HMP1) also is induced. This protein exhibits cruciform binding activity thought to be necessary for the resolution of Holliday structures during homologous recombination (Dutta et al., 1997), which can be important for DNA repair. Second, we observed induction of a homologue of indoleamine-2,3-dioxygenase (H.c. IDO1), an enzyme in the kynurenine pathway. This protein modifies tryptophan during the first step in NAD+ synthesis (Panozzo et al., 2002). NAD+ is required for energy production and DNA repair, among other cellular processes. The up-regulation of IDO1 may allow H. capsulatum to increase levels of NAD+ during nitrosative stress. Alternatively, H. capsulatum could be using IDO1 to deplete tryptophan and thereby impair macrophage .NO production: in culture, depletion of tryptophan prevents NOS2 induction by IFN- in macrophages (Chiarugi et al., 2003). Additionally, a metabolite of indoleamine-2,3-dioxygenase has been shown to down-regulate NOS2 expression and inhibit Nos2p activity in a macrophage cell line (Sekkai et al., 1997; Oh et al., 2004). Finally, whereas homologues of H. capsulatum arginase (H.c. CAR1) are part of the glutamine biosynthesis pathway, other interesting possibilities exist for the role of arginase in RNS response. One of the products of the arginase reaction is L-ornithine (http://www.genome.jp/dbget-bin/www_bget?enzyme+3.5.3.1). L-ornithine is utilized by Lom1p during siderophore biosynthesis as discussed previously. By producing L-ornithine, arginase could facilitate siderophore biosynthesis and iron acquisition. Alternatively, because Nos2p uses arginine as a substrate to produce .NO, reduction of environmental levels of arginine by arginase could reduce macrophage Nos2p activity. Studies in Helicobacter pylori suggest that H. pylori arginase inhibits .NO production by macrophages by depleting extracellular arginine concentrations (Gobert et al., 2001). It is possible that H. capsulatum induces arginase in response to nitrosative stress to reduce further .NO production by host cells.
H. capsulatum P450 Nitric Oxide Reductase (P450nor) Is Sufficient to Provide Increased RNS Resistance In Vitro
In other organisms, genes involved in .NO detoxification have been observed to be up-regulated during nitrosative stress and to be required for RNS resistance. For example, flavohemoglobins, such as E. coli Hmp, are able to convert .NO into nitrate under aerobic conditions and into nitrous oxide under anaerobic conditions (reviewed in Poole and Hughes, 2000). Studies in a range of organisms including E. coli, S. typhimurium, S. cerevisiae, C. albicans, and C. neoformans have shown that deletion of flavohemoglobin results in .NO sensitivity (Crawford and Goldberg, 1998; Membrillo-Hernandez et al., 1999; Liu et al., 2000; de Jesus-Berrios et al., 2003; Ullmann et al., 2004; Hromatka et al., 2005; Sarver and DeRisi, 2005). Flavorubredoxin and NADH:(flavo)rubredoxin oxidoreductase, exemplified by E. coli NorVW, are capable of reducing .NO under anaerobic conditions (Gardner et al., 2002). Deletion of these enzymes in E. coli also results in RNS sensitivity (Gardner et al., 2002; Hutchings et al., 2002; Justino et al., 2005). BLASTX searches of the recently assembled H. capsulatum genome sequence failed to identify homologues to either of these classes of enzymes. H. capsulatum does have a homologue to GSNO reductase, an enzyme that contributes to the nitrosative stress response in other organisms by detoxifying GSNO (Liu et al., 2001; de Jesus-Berrios et al., 2003). Although the H. capsulatum GSNO reductase is transcriptionally induced upon GSNO addition (our unpublished data), the H. capsulatum gene has no apparent start codon, and thus its role in combating nitrosative stress is unclear. The only obvious candidate for a .NO detoxifying enzyme observed in these experiments was a P450nor (Figure 8A). P450nor was originally identified in F. oxysporum where it detoxifies .NO to nitrous oxide (N2O) during the process of denitrification (Shoun and Tanimoto, 1991; Nakahara et al., 1993).
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