Characterization of the Bat proteins in the oxidative stress response of Leptospira biflexa
© Stewart et al.; licensee BioMed Central Ltd. 2012
Received: 30 July 2012
Accepted: 10 December 2012
Published: 13 December 2012
Leptospires lack many of the homologs for oxidative defense present in other bacteria, but do encode homologs of the Bacteriodes aerotolerance (Bat) proteins, which have been proposed to fulfill this function. Bat homologs have been identified in all families of the phylum Spirochaetes, yet a specific function for these proteins has not been experimentally demonstrated.
We investigated the contribution of the Bat proteins in the model organism Leptospira biflexa for their potential contributions to growth rate, morphology and protection against oxidative challenges. A genetically engineered mutant strain in which all bat ORFs were deleted did not exhibit altered growth rate or morphology, relative to the wild-type strain. Nor could we demonstrate a protective role for the Bat proteins in coping with various oxidative stresses. Further, pre-exposing L. biflexa to sublethal levels of reactive oxygen species did not appear to induce a general oxidative stress response, in contrast to what has been shown in other bacterial species. Differential proteomic analysis of the wild-type and mutant strains detected changes in the abundance of a single protein only – HtpG, which is encoded by the gene immediately downstream of the bat loci.
The data presented here do not support a protective role for the Leptospira Bat proteins in directly coping with oxidative stress as previously proposed. L. biflexa is relatively sensitive to reactive oxygen species such as superoxide and H2O2, suggesting that this spirochete lacks a strong, protective defense against oxidative damage despite being a strict aerobe.
Molecular oxygen freely diffuses across bacterial membranes and can give rise to damaging reactive oxygen species (ROS) such as superoxide radicals (O2−), hydrogen peroxide (H2O2), and hydroxyl radicals (OH·). These highly reactive molecules lead to a variety of harmful effects within the bacterial cell, including inactivation of Fe-S-containing proteins and damage to DNA and to lipids, in some bacteria. For aerobic microorganisms the presence of these toxic species is by nature unavoidable and they have therefore evolved a variety of protective enzymes to preemptively detoxify ROS.
The enteric bacteria have been intensively studied for their response to ROS (recently reviewed by ). In contrast, leptospires lack a number of the enzymes used by enteric bacteria to combat oxidative damage  and are also more susceptible to H2O2-mediated killing than other microorganisms . Nascimento and colleagues speculated that the Bat proteins of L. interrogans might partially compensate for the shortage of oxidative stress proteins by providing an additional line of defense against oxidative damage .
The Bat proteins were first identified by Tang and co-workers in a transposon mutagenesis screen of the anaerobe Bacteroides fragilis. The transposon inserted into the fourth of five contiguous open reading frames and resulted in reduced levels of aerotolerance and pathogenicity. The genes were designated bat, for Bacteriodes aerotolerance genes, and were shown to comprise an operon. The mutant phenotype could be partially complemented by the addition of reducing agents and the Bat proteins were proposed to directly reduce oxidatively-damaged proteins in the periplasm or, alternatively, to help create a reduced environment in the periplasm by exporting reducing power equivalents. Interestingly, anaerobic growth did not restore the growth rate to that of wild-type and the addition of reducing agents also increased growth of the wild-type strain, although not as dramatically as it did for the mutant.
Recently, two bat homologs in Francisella tularensis were inactivated and the bat mutants were shown to have a reduced ability to replicate in macrophage cells and were also attenuated for virulence in a mouse model . The specific function of the Bat proteins, however, was not determined in F. tularensis. Genome sequences have identified homologs in a wide variety of other prokaryotes, including all families that comprise the phylum Spirochaetes (Brachyspiraceae, Leptospiraceae, and Spirochaetaceae). Although conserved in all branches of the Spirochaetes, the number and combination of bat homologs vary by species. However, the function of the Bat proteins in spirochetes or in any other species has not been elucidated.
Although pathogenic leptospires also contain bat homologs and are more resistant to peroxide exposure than the saprophyte L. biflexa[3, 6], the pathogenic spp. are notoriously recalcitrant to targeted allelic exchange. Since L. biflexa is more amenable to genetic manipulation than pathogenic species, it serves as a model organism for genetic studies in leptospires. Therefore, we used L. biflexa to investigate the function of the Bat proteins and to better understand the response of leptospires to oxidative stress. Here, we report the engineered deletion of the three contiguous L. biflexa bat genes and characterization of the mutant phenotype and oxidative stress response.
The bat genes are distributed throughout the Spirochaetesand encode conserved protein motifs
Homologs of the bat genes are present in each family of the Spirochaetes (Additional file 1: Figure S1), although not in all species. In contrast to the 5 genes present in B. fragilis, L. biflexa contains 3 bat genes and the pathogenic leptospires contain 4 [2, 7–9]. However, the batB and batC genes are fused in L. biflexa, which does not appear to be the case for the pathogenic species, and explains the discrepancy in gene number. Fusions of bat coding regions also appear to have occurred in Borrelia burgdorferi and Spirochaeta thermophila (Additional file 1: Figure S1) and were also reported for F. tularensis type A strain Schu S4 .
Deletion of batgenes
Transcript analyses indicate independent promoters in the batgene cluster
Morphology and growth rate of batmutants are equivalent to wild-type
Growth rates of WT, ΔbatA, and ΔbatABD strains were compared during in vitro cultivation in EMJH liquid medium and also for colony formation on solid EMJH medium. No significant differences in growth rate were observed when cultured in liquid medium, regardless of whether the cultures were aerated or static (Figure 4B). Colony morphology and rate of formation were similar among all strains (data not shown).
As the mutant strains did not display an obvious growth defect compared to WT, we assessed the growth dynamics of both parent and mutant when cultured together in the same medium (Figure 4C). WT and Δbat-ABD strains were co-inoculated into the same cultures (performed in triplicate) and assessed daily to determine if population ratios changed over time. As shown in Figure 4C, relative proportions of each strain did not change significantly over time and this was statistically confirmed by two-way Analysis of Variance (ANOVA) with the Bonferroni post-test. Therefore, the Bat proteins do not significantly affect L. biflexa growth, either in pure culture or when the mutant is mixed with an equal density of WT cells.
Deletion of batgenes does not alter tolerance to oxidative stress
L. biflexalacks an inducible stress response to ROS
Differential proteomic analysis of WT and mutant strains detected changes in HtpG levels among membrane-associated proteins
Bat homologs are present in all families of the Spirochaetales (Additional file 1: Figure S1), despite the vast evolutionary divergence noted in this order . The retention of these proteins suggests they confer an evolutionary advantage to spirochetes, even though the environment and life cycle of these bacteria are incredibly diverse, ranging from free-living aerobic saprophytes (L. biflexa) and anaerobic thermophiles (Spirochaeta thermophila) to mammalian pathogens (L. interrogans and B. burgdorferi). L. borgpetersenii, purportedly undergoing genome reduction, retains the same number and order of bat genes as L. interrogans, again suggesting the Bat proteins provide an important function that prevents their elimination even in a decaying genome. One spirochete appears to be an exception to this theory – the obligate human pathogen and syphilis agent, Treponema pallidum, in which we were unable to identify any Bat homologs.
In addition to the wide-spread distribution of the Bat proteins in the Spirochaetales, MoxR and HtpG are also encoded in many spirochete genomes (data not shown). The MoxR chaperone is postulated to coordinate the metal ion into the Bat proteins MIDAS domain (Figure 1) . In the sequenced Leptospira genomes, moxR and htpG are located in the same contiguous gene cluster as the bats (Figure 2A) [2, 7–9]. However, Dieppedale et al. inactivated moxR in F. tularensis and their proteomic comparisons of wild-type to the moxR mutant did not identify changes in Bat protein levels . HtpG is a homolog of the eukaryotic heat shock protein Hsp90, but its function in bacteria is unclear and it has been reported to have different roles in different prokaryotes [19–22]. The arrangement of the 11 tandem ORFs in this cluster suggest they potentially form a large operon, but qRT-PCR analyses detected transcript from the ORFs downstream of the deleted bat genes. The presence of transcript from the downstream ORFs, regardless of the orientation of the inserted kanamycin-resistance cassette, implies that these genes can be independently transcribed (Figure 3). These data do not rule out the possibility of an additional promoter that drives expression of all 11 genes in an operon, but do support independent promoters for the genes downstream of the deleted regions.
Somewhat surprisingly, transcript from genes immediately following the deletion site had detectable levels of transcript, although these levels were significantly lower than WT levels. Specifically, transcript of batB was detected in the ΔbatA strain, even though the endogenous promoter is likely to be located in the deleted batA gene. However, batB transcript levels are over 10-fold lower in the ΔbatA strain compared to wild-type, suggesting that the kanamycin-resistance cassette upstream of batB may provide a weak, fortuitous promoter sequence. A similar result was also observed for htpG transcript in the Δbat-ABD strain; presumably, the htpG promoter would be located in the deleted region. The borrelial flgB promoter used to drive kan expression in the deletion of batABD is oriented in the same transcriptional direction as the endogenous genes (specifically, htpG) and read-through may account for the htpG transcript detected, albeit at a lower level than the endogenous promoter would produce.
The presence of a signal sequence, transmembrane helices and motifs for protein-protein interactions, also conserved in the Bat proteins of Leptospira (Figure 1), led Tang et al. to propose that the Bat proteins of B. fragilis formed a complex in the periplasm . Despite their putative cellular location, growth rate and morphology of L. biflexa were unaffected by the loss of these proteins (Figure 4). Nor could we demonstrate a protective role for the Bat proteins in coping with oxidative stress, as initially proposed for B. fragilis and subsequently hypothesized for other spirochetes [2, 14]. The wild-type and ΔbatABD mutant were equally susceptible to oxidative challenge by both peroxides and the superoxide generator paraquat (Figure 5), indicating that the Bat proteins do not contribute to L. biflexa’s limited ability to cope with oxidative damage. However, the lack of an observable phenotype for the bat mutants may relate to in vitro growth where the transcript levels for these genes is quite low relative to flaB or htpG transcript levels (Figure 3). It is conceivable that bat expression may increase under specific in vivo conditions of which we are unaware. Various microarray studies, however, did not detect any significant changes in bat transcript levels in pathogenic leptospires when in vitro conditions were altered to mimic in vivo environments [23–29].
We also examined the potential contribution of the Bat proteins to sensing ROS and inducing an oxidative stress response in L. biflexa. Enteric bacteria such as E. coli and Salmonella typhimurium have well-characterized oxidative stress responses that can be induced by the addition of sublethal levels of peroxide [15, 16] or superoxide [30–32]. However, pretreatment of exponentially growing L. biflexa cultures with either 1 μM H2O2 or 0.5 μM paraquat failed to confer a higher level of resistance to ROS when subsequently challenged with lethal levels (Figure 6). Therefore, it appears that L. biflexa does not have the same capability as enteric bacteria of inducing an oxidative stress response, at least under the conditions tested. L. biflexa lacks homologs for the two main regulators of the oxidative stress response in enteric bacteria (SoxR and OxyR), in support of this conclusion. However, Leptospira spp. do possess a PerR homolog (LEPBI_I2461 in L. biflexa), a negative regulator of peroxide defense first characterized in Gram positive bacteria (reviewed in ). Lo et al. reported a PerR transposon mutant of L. interrogans that resulted in an 8-fold increase in resistance to hydrogen peroxide over the wild-type . However, microarray data of this mutant did not report any significant changes in bat transcript, suggesting that these genes may not be under the regulatory control of PerR. It is still possible that the Bat proteins are involved in sensing ROS, but the cellular response they may direct remains enigmatic.
Surprisingly, even wild-type L. biflexa is highly susceptible to oxidative stress compared to B. burgdorferi (10 μM vs. 10 mM, respectively, for t-Butyl hydroperoxide)  or E. coli. The relative susceptibility of L. biflexa to oxidative damage may be due to the absence of some proteins capable of detoxifying ROS or repairing damaged proteins. For example, L. biflexa does not have recognizable homologs of glutathione reductase, thioredoxin 2, Ferric reductase, and others. However, L. biflexa does possess both superoxide dismutase (Sod) and KatG (a Hydroperoxidase I enzyme), two enzymes widely conserved among aerobic organisms for defense against ROS. Sod catalyzes the reduction of O2− to H2O2 and O2. Hydrogen peroxide is itself an oxidant that freely diffuses across membranes and can give rise to the highly reactive hydroxyl radical. Therefore, aerobic cells require a mechanism for detoxifying H2O2. Catalase or peroxidase enzymes usually fulfill this cellular function and a gene encoding KatG, which can have either activity, has been identified in the L. biflexa genome (LEPBI_I2495). Since catalase activity has not been detected in L. biflexa strains but peroxidase activity has [36–40], it seems likely that KatG is a peroxidase and provides a mechanism by which L. biflexa detoxifies H2O2, albeit not very effectively. L. biflexa also possesses alkyl hydroperoxide reductase homologs (LEPBI_I3008 & LEPBI_I3009) that may also detoxify H2O2. Superoxide dismutase may play an essential role in L. biflexa’s defense against oxidative stress, as we were unable to inactivate the sod gene, either by allelic exchange or by transposon mutagenesis (data not shown).
Finally, we employed a proteomic comparison of wild-type and mutant spirochetes to identify L. biflexa proteins whose expression may be altered due to the loss of the Bat proteins. Two-dimensional differential gel electrophoresis of protein lysates from the wild-type and the ΔbatABD strain identified HtpG as the sole protein in the ΔbatABD strain that had significantly reduced levels compared to the wild-type (Figure 7). Altered levels of HtpG were detected in the membrane-associated protein fraction, but not the soluble fraction (data not shown), although HtpG does not have any recognizable signal or lipidation sequences. However, Lo et al. also reported that HtpG associated with the membrane fraction in their analyses of temperature effects on protein levels in L. interrogans. In our analysis, HtpG was downregulated approximately 4-fold in the ΔbatABD mutant relative to the WT, and this decrease corresponded to the 3.8-fold decrease in htpG transcript levels observed by qRT-PCR (Figure 3), discussed above. Although HtpG protein is lower in the mutant, this variation did not produce a phenotype in the conditions tested here.
L. biflexa has a relatively small repertoire of enzymes for defense against ROS, and it may depend on the activities of Sod and KatG to survive oxidative assault. During in vitro growth, bat transcript levels are relatively low and deletion of the bat loci did not detectably alter morphology, growth rate, or the ability to survive oxidative stress. Despite the proposed role for the Bat proteins in directly combating oxidative damage in spirochetes, the data presented here do not support this. Although we cannot exclude a role for the Bat proteins in sensing oxidative stress in L. biflexa, perhaps as a signaling complex in the periplasm, Bat function remains elusive.
Bacterial strains used in this study
L. biflexa serovar Patoc strain Patoc I (kindly provided by Dr. Dave Haake and Dr. Marija Pinne) was cultured at 30°C with shaking at 150 RPM in EMJH medium (Fisher Scientific, Pittsburgh, PA) [41, 42]. Plating medium included 1.2% wt/vol Noble agar (final concentration) (Fisher Scientific) and plates were incubated at 30°C, inverted and sealed with parafilm. Kanamycin, when needed, was added to the medium at a final concentration of 20 μg/mL. Escherichia coli strains TOP10 (Invitrogen, Carlsbad, CA) or NEB5α (New England Biolabs, Ipswich, MA) were used for all plasmid manipulations.
Construction of L. biflexamutant strains
Oligonucleotides used in this study
Sequence (5′ – 3′)
Amplification of batABD + flanking sequences
Amplification of batABD + flanking sequences
Amplification of batA + flanking sequences
Amplification of batA + flanking sequences
Construction of allelic-exchange plasmids
Construction of ΔbatABD allelic-exchange plasmid
Construction of ΔbatA allelic-exchange plasmid
Amplification of kan
Amplification of kan
Amplification of batA probe
Amplification of batA probe
Amplification of batB/D probe
Amplification of batB/D probe
RNA isolation and quantitative RT-PCR analysis
Total RNA was isolated from 10 mL cultures of exponentially growing L. biflexa cells using TRIzol reagent (Invitrogen). Cells were pelleted at 7,000 RPMs in 15 mL Falcon 2059 tubes and the pellet resuspended in 5.0 mL TRIzol. After incubation at room temperature for 2.5 min with vigorous shaking, 1 mL of chloroform was added, mixed and incubated for a further 2.5 min. The suspension was centrifuged again and the aqueous phase removed to a new Falcon tube and the RNA precipitated by addition of 5 mL isopropanol. Following a 10 minute incubation (room temperature), RNA was pelleted, washed in 75% ethanol and dissolved in 100 μL of RNase-free water. DNA was removed by treating with Turbo DNase (Ambion, INC. Austin, TX) following the manufacturer’s recommendations.
RNA was converted to cDNA using the High-Capacity cDNA Reverse Transcription kit (Applied Biosystems, Foster City, CA); reaction mixtures consisted of 1 μg RNA and were converted to cDNA per the manufacturer’s recommendations. The cDNA samples were diluted 1:20 with water and 2 μL used for subsequent quantitative PCR (qPCR) reactions. All samples were analyzed in triplicate. TaqMan Universal PCR Master Mix kit (Applied Biosystems) and PCR conditions were as previously described . L. biflexa genomic DNA ranging from 106 cells to 10 cells (in 10-fold serial dilutions) was used to generate a standard curve using the Ct values from the flaB primer/probe set. This standard curve was then used to interpolate the number of transcript copies from the Ct values generated from gene-specific primer/probe sets. The resulting transcript levels were then normalized to 104 copies of flaB transcript. Negative controls lacking reverse transcriptase were included to demonstrate that all genomic DNA had been degraded and did not contribute to the signal.
Electron microscopy, growth rate analysis and oxidative stress assays
Bacterial suspensions from cultures grown in EMJH media were prepared for scanning electron microscopy (SEM) essentially as described previously . Samples were lightly sputtered with iridium and examined on a model SU-8000 scanning electron microscope operated at 2 kV (Hitachi High Technologies America, Pleasanton, CA). Images were digitized using the on-board frame card according to the manufacturer’s specifications.
For transmission electron microscopy (TEM), bacteria were prepared as described previously for imaging by microwave-assisted processing . Grids were examined using a model H-7500 transmission electron microscope, operated at 80 kV (Hitachi). Digital images were captured and recorded using a model HR100 camera system (Advanced Microscopy Techniques, Danvers, MA).
Growth rate comparisons were performed in quadruplicate. Five mL cultures were inoculated at 105 cells/mL from a starter culture grown to between 5 × 108 to 1 × 109 cells/mL, as determined by counting with Petroff-Hauser counting chambers. All cultures were incubated at 30°C; aerated cultures were shaken at 150 RPM. Cell densities were measured by optical density at 420 nm in a spectrophotometer.
Co-growth comparisons of wild-type and mutant strains were similarly tested with each strain inoculated at 105 cells/mL in the same culture (for a combined concentration of 2 × 105 cells/mL). Aliquots were removed daily from triplicate cultures, counted and diluted appropriately for colony formation on non-selective EMJH agar plates. PCR was performed on 24–30 colonies per plate to enumerate wild-type and mutant cells by amplifying a fragment of batB (wild-type) and the kanamycin-selectable marker (mutant).
Oxidative stress assays were also performed similarly. Peroxide-treated cultures were first diluted to 103 cells/mL and peroxides were then added at specified concentrations and incubated for approximately 2 ½ hours, after which 100 μL samples were removed from each culture and spread on EMJH agar plates. After 4–6 days of incubation at 30°C, plates were removed and colony counts used to calculate viable cells. A similar strategy was followed for assessing whether an oxidative stress response could be induced in L. biflexa; quadruplicate cultures of 103 cells/mL were exposed to a sublethal level of H2O2 (1μM) for 3 hrs with aeration, followed by the addition of specified concentrations of peroxide and a further incubation for 3 hours. Aliquots of 100 μL were removed and spread on EMJH agar plates to determine viable cell counts. Superoxide sensitivity was determined by diluting triplicate cultures to 5 × 106 cells/mL and exposing to various concentrations of the superoxide-generating molecule paraquat (Sigma-Aldrich, St. Louis, MO) with incubation for 24 hrs. Cell viability was determined by counting motile cells using a Petroff-Hauser chamber with darkfield microscopy. To determine if L. biflexa produces an oxidative stress response to superoxide, triplicate cultures of 5 × 106 cells/mL were pre-exposed to 0.5 μM paraquat for 2.5 hrs followed by addition of specific concentrations of paraquat. Cultures were further incubated for 24 hrs and cell viability assessed as described above.
Two-dimensional differential in-gel electrophoresis (2D-DIGE) and protein identification
L. biflexa isolates were grown to a cell density of ~1 × 109 cells/ml and harvested by centrifugation (10,000 × g, 10 min, 23°C). Cell pellets were rinsed in PBS and lysed in PBS supplemented with 1 X Complete Protease Inhibitor (Roche Applied Science) by 3 passes through a French pressure cell (16,000 lb/in2). Cell lysates were further fractionated into soluble and membrane-associated proteins by ultracentrifugation (100,000 × g 1 h, 4°C). The membrane-associated protein pellet was rinsed with PBS and suspended in PBS+PI with the aid of a glass tissue homogenizer (Kontes Glass Co.,Vineland, NJ). Protein concentrations were determined by a modified Lowry protein assay with bovine serum albumin as a standard.
For DIGE analysis of membrane-associated proteins, 50 ug of L. biflexa wild-type or the ΔbatABD isolate was labeled with either 400 pmol Cy3 or Cy5 (CyDye minimal dye labeling kit, GE Healthcare) for 30 min on ice. As an internal control, a mixture of 25ug of the wild-type and 25 ug of the ΔbatABD samples were labeled with Cy2 for 30 min on ice. All labeling reactions were performed in DIGE labeling solutions consisting of 7 M Urea, 2M Thiourea, and 4% CHAPS in 10 mM Tris (pH 8.5). The labeling reaction was quenched by adding 1 ul of 10 mM lysine and incubating for 10 min on ice. To ensure that observed differences were not due to artifacts from preferential dye binding to proteins, several coupled samples were labeled by dye switching. Labeled proteins were stored at −20°C in the dark until isoelectric focusing.
Cy-dye labeled samples for comparison were mixed and DTT and IPGphore 3–10 buffer were added at final concentrations of 100 mM and 1.0%, respectively. The volume of each set was brought to 350 ul with isoelectric focusing solution C4TT  and applied to 18 cm 3–10 non-linear IPG strips (GE Healthcare). Strips were focused using the following parameters: 12 hr rehydration, 500 V for 1 hr, 1000 V for 1 hr, 1500 V for 1 hr, 4000 V for 1 hr, and 8000V for 60,000 Vhr. Once focusing was complete, strips were stored at −80°C until equilibrated and separated in the second dimension by standard SDS-PAGE using 8-16% gradient gels (Jule, Inc., Milford, CT) .
After separation of DIGE-labeled strips by SDS-PAGE, gels were scanned in the glass plates using a three laser Typhoon 9400 variable mode imager (GE Healthcare, Piscataway, NJ) at 200 microns. Differences in protein spots were quantified using DeCyder 2-D Differential Analysis Software v7.2. Protein spots of interest were excised and processed for mass spectrometry as previously described . Dried peptides were sent to the Protein Chemistry section of the NIAID Research Technologies Branch, NIH for identification as described below.
The recovered peptides were re-suspended in 5 ul of Solvent A (0.1% formic acid, 2% acetonitrile, and 97.9% water). Prior to mass spectrometry analysis, the re-suspended peptides were chromatographed directly on column, without trap clean-up. The bound peptides were separated at 500 nl/min generating 80–120 Bar pressure, using an AQ C18 reverse phase media (3 u particle size and 200 u pore) packed in a pulled tip, nano-chromatography column (0.100 mm ID × 150 mm L) from Precision Capillary Columns, San Clemente, CA. The chromatography was performed in-line with an LTQ-Velos Orbitrap mass spectrometer (ThermoFisher Scientific, West Palm Beach, FL) and the mobile phase consisted of a linear gradient prepared from solvent A and solvent B (0.1% formic acid, 2% water, and 97.9% acetonitrile) at room temperature. Nano LC-MS (LC-MS/MS) was performed with a ProXeon Easy-nLC II multi-dimensional liquid chromatograph and temperature controlled Ion Max Nanospray source (ThermoFisher Scientific) in-line with the LTQ-Velos Orbitrap mass spectrometer.
Mass calibration was performed as needed with the positive ion Cal Mix prepared as described by Thermo-Scientific and monitored by routine analysis of a 10 femtomole stock sample of BSA digest. Typical acceptable results for this analysis would yield a 2800 – 3300 Mascot score, 75 – 85% coverage and 0 - +/−4 ppm error when submitted to the Mascot server using Proteome Discoverer 1.3 using the Swiss Prot-Trembl data base.
Computer controlled data dependent automated switching to MS/MS by Xcalibur 2.1 software was used for data acquisition and provided the peptide sequence information. Data processing and databank searching were performed with PD 1.3 and Mascot software (Matrix Science, Beachwood, OH).
The authors gratefully acknowledge the generous gifts of strains and advice from David Haake and Marije Pinne. We also thank Joe Hinnebusch and Frank Gherardini for critical reading of the manuscript; Dan Sturdevant, Kevin Lawrence and Julie Boylan for technical advice and helpful discussions, Jeff Skinner at Bioinformatics and Computational Biosciences Branch for statistical analysis, and Scott Samuels’ lab for technical advice on RNA isolation.
This research was supported by the Intramural Research Program of the NIH, NIAID.
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