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Summary

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. Supplementary material
  9. References
  10. Supporting Information

The suf and isc operons of Escherichia coli have been implicated in Fe–S cluster assembly. However, it has been unclear why E. coli has two systems for Fe–S cluster biosynthesis. We have examined the regulatory characteristics and mutant phenotypes of both operons to discern if the two operons have redundant functions or if their cellular roles are divergent. Both operons are similarly induced by hydrogen peroxide and the iron chelator 2,2′-dipyridyl, although by different mechanisms. Regulation of the isc operon is mediated by IscR, whereas the suf operon requires OxyR and IHF for the response to oxidative stress and Fur for induction by iron starvation. Simultaneous deletion of iscS and most suf genes is synthetically lethal. However, although the suf and isc operons have overlapping functions, they act as distinct complexes because the SufS desulphurase alone cannot substitute for the IscS enzyme. In addition, suf deletion mutants are more sensitive to iron starvation than isc mutants, and the activity of the Fe–S enzyme gluconate dehydratase is diminished in the suf mutant during iron starvation. These findings are consistent with the model that the isc operon encodes the housekeeping Fe–S cluster assembly system in E. coli, whereas the suf operon is specifically adapted to synthesize Fe–S clusters when iron or sulphur metabolism is disrupted by iron starvation or oxidative stress.


Introduction

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. Supplementary material
  9. References
  10. Supporting Information

Fe–S clusters function as key cofactors in multiple cellular processes (reviewed in Beinert and Kiley, 1999; Frazzon and Dean, 2003). In vitro assembly of Fe–S clusters in proteins can occur spontaneously but in vivo Fe–S biogenesis requires specific accessory proteins (reviewed in Frazzon and Dean, 2003). Early studies of Fe–S cluster assembly in the complex nitrogenase enzyme of Azotobacter vinelandii identified two major components required for cluster building, NifS and NifU (Jacobson et al., 1989). NifS is part of a large family of pyridoxal-phosphate dependent cysteine desulphurases that mobilize sulphur from l-cysteine (Zheng et al., 1993). NifU was found to be an Fe–S cluster scaffold upon which the nascent cluster can be constructed before its transfer to an apo-protein (Yuvaniyama et al., 2000). Homologues of these two core components can be found in organisms ranging from bacteria to humans.

The Gram-negative bacterium Escherichia coli contains three gene products that are homologous to NifS, namely IscS, SufS (also known as CsdB) and CsdA (also known as CSD or YgdJ). IscS and the NifU-like scaffold protein IscU are encoded by the iscRSUA operon and are required for assembly of a variety of Fe–S enzymes in E. coli (Flint, 1996; Schwartz et al., 2000; Urbina et al., 2001; Kato et al., 2002). The co-expressed IscA protein can form Fe–S clusters in vitro and may be an alternative scaffold or an iron handling protein (Krebs et al., 2001; Ollagnier-De-Choudens et al., 2001). Deletion mutants of isc possess a variety of growth defects as the result of the loss of Fe–S enzyme activity and the disruption of sulphur metabolism (Kambampati and Lauhon, 1999; Takahashi and Nakamura, 1999; Lacourciere et al., 2000; Schwartz et al., 2000; Tokumoto and Takahashi, 2001; Mihara et al., 2002).

The function of the sufABCDSE operon encoding SufS is less clear. SufA and SufS are homologous to IscA and IscS, respectively, but the suf operon does not contain a direct homologue to the Fe–S scaffold NifU. SufC is a soluble ATPase similar to those found associated with transporters (Rangachari et al., 2002; Nachin et al., 2003). The SufB and SufD proteins do not have assigned functions, although they do interact with SufC to form a complex (Rangachari et al., 2002; Nachin et al., 2003; Outten et al., 2003). SufE was recently shown to be a novel sulphur transfer protein that enhances SufS cysteine desulphurase activity by accepting sulfane sulphur (Loiseau et al., 2003; Ollagnier-De-Choudens et al., 2003; Outten et al., 2003). The SufBCD complex further stimulates SufE enhancement of SufS by an unknown mechanism (Outten et al., 2003). The suf operon was originally identified based on loss of the Fe–S cluster in the putative iron reductase FhuF in sufD or sufS deletion strains (Patzer and Hantke, 1999). FhuF is necessary for utilization of ferrioxamine B or rhodotorulic acid as sole iron sources (Muller et al., 1998), and the sufS and sufD mutants showed an inability to grow using these exclusive iron sources. Subsequently it was discovered that suf mutants of Erwinia chrysanthemi have similar limitations in their ability to utilize certain iron sources (Nachin et al., 2003). Recent work also has shown that, in certain media, suf mutants in E. coli are sensitive to phenazine methosulphate, a redox active reagent that can generate superoxide as well as oxidize protein thiols and intracellular iron, and that these defects are due to decreased activity of some Fe–S enzymes (Nachin et al., 2003).

The third NifS-like gene product, CsdA, can donate sulphur for molybdopterin synthesis in vitro but there have been no published phenotypes for csdA mutants (Leimkuhler and Rajagopalan, 2001). Interestingly, csdA is located on the chromosome adjacent to ygdK, which is homologous to SufE.

Recently it was discovered that the iscRSUA and sufABCDSE operons are induced during exposure to hydrogen peroxide (H2O2) in E. coli (Zheng et al., 2001). The only known regulator of isc is IscR, a transcriptional repressor co-expressed with the other isc genes. Repression of isc expression by IscR is relieved when a 2Fe−2S cluster present in IscR is damaged or fails to form, and a non-polar deletion in iscR causes a 17-fold increase in basal expression of the isc promoter (Schwartz et al., 2001). Induction of the suf promoter by H2O2 was found to be dependent on OxyR, the global regulator of the cellular response to H2O2 stress (Zheng et al., 2001). The expression of sufS– and sufD–lacZ fusions also were found to be regulated by Fur, a metalloregulatory protein that controls transcription of genes involved in iron metabolism (Patzer and Hantke, 1999). A subsequent DNA microarray study in E. coli confirmed that the suf operon is part of the Fur regulon in E. coli (McHugh et al., 2003). The suf system in E. chrysanthemi is also regulated by Fur in response to iron availability (Nachin et al., 2001).

The fact that the isc and suf operons both contain NifS homologues and both have phenotypes that connect them to Fe–S cluster assembly raises the question of why E. coli has two loci for Fe–S cluster assembly. The two operons may have distinct roles, such as is the case with the nif and isc operons of A. vinelandii, where the nif operon is specialized for the construction of nitrogenase whereas the isc operon assembles clusters for other Fe–S enzymes (Jacobson et al., 1989; Zheng et al., 1998). The isc and suf operons do contain different accessory genes that are co-expressed with the NifS homologue, indicating some likelihood of differential function. Alternatively, the two operons may be regulated differently to provide similar functions under distinct conditions in the cell. To gain more insight into these possibilities, we examined the regulatory characteristics and the mutant phenotypes of the isc and suf operons. Based on reported iron phenotypes for suf mutant strains and on the results presented here, we propose that the suf operon in E. coli is an Fe–S cluster assembly pathway specifically adapted for constructing Fe–S clusters under iron limiting conditions.

Results

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. Supplementary material
  9. References
  10. Supporting Information

Strong suf and isc induction by H2O2 and 2,2′-dipyridyl

To assess whether the suf or isc operons are differentially induced by H2O2, we carried out primer extension assays to measure suf and isc transcript levels in cells exposed to a range of H2O2 concentrations. Both operons were similarly responsive to H2O2, with the highest induction occurring at 300 µM H2O2(Fig. 1A). We also examined the suf and isc mRNA levels in cells treated with various amounts of the intracellular iron (II) chelator 2,2′-dipyridyl. The basal levels of the suf transcript appear to be lower than the basal levels of the isc transcript, but the two operons showed similar fold induction by 2,2′-dipyridyl-mediated iron starvation (Fig. 1B). These results show that the induction of the suf and isc operons in response to H2O2 and 2,2′-dipyridyl is similar.

image

Figure 1. Regulation of suf and isc in response to H2O2 and 2,2-dipyridyl. A. Primer extension analysis of suf and isc transcription in cells exposed to varying concentrations of H2O2 for 5 min. B. Primer extension analysis of suf and isc transcription in cells exposed to varying concentrations of 2,2′-dipyridyl for 60 min.

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iscR is required for isc induction in response to H2O2 and 2,2′-dipyridyl

To ascertain if the induction of the isc operon in response to oxidative stress and iron starvation is mediated through the IscR repressor, we examined isc induction in response to H2O2 and 2,2′-dipyridyl in a mutant strain with a non-polar deletion of iscR. An iscR deletion strain shows constitutive expression of the isc operon and isc expression is not further induced in response to H2O2 or 2,2′-dipyridyl (Fig. 2A and B). This result indicates that the IscR repressor accounts for the transcriptional induction of the isc promoter in response to oxidative stress and iron starvation.

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Figure 2. Regulation of isc in response to H2O2 and 2,2-dipyridyl in an iscR mutant. Autoradiograph exposure times were optimized for each panel to visualize the isc transcript in the wild-type background, and therefore do not allow direct comparison between panels A and B. A. Primer extension analysis of isc transcription in wild-type and iscR mutant cells exposed to 1 mM H2O2 for 5 min. B. Primer extension analysis of isc transcription in wild-type and iscR mutant cells exposed to 100 µM 2,2′-dipyridyl for 60 min.

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Both OxyR and IHF are required for suf induction in response to H2O2

To further characterize the suf transcriptional response to H2O2 and 2,2′-dipyridyl, we defined the regulatory features of the suf promoter. It was previously shown that OxyR regulates suf in response to H2O2 and that OxyR binds to the suf promoter (Zheng et al., 2001). Interestingly, the OxyR binding site is located at −197 to −244, much further from the −35 promoter element than is typically seen with OxyR regulated genes (Fig. 3A). Examination of the suf promoter revealed a putative binding site for integration host factor (IHF) located at −139 to −151 (Fig. 3A). IHF, a DNA-bending protein, plays a key role in the regulation of a number of stress response genes and is thought to be important for the cellular adaptations that occur upon entry into stationary phase (Nash and Robertson, 1981; Stenzel et al., 1987; Altuvia et al., 1994; Ditto et al., 1994).

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Figure 3. Regulation of the suf promoter. A. The +1, −10 and −35 elements of the suf promoter are underlined, the start codon for sufA is shown in bold capital letters, the putative Fur site is indicated with asterisks under the sequence, and the putative IHF site is indicated with double asterisks under the sequence. The sequences protected by Fur, IHF and OxyR in DNase I footprinting assays are denoted with solid bars. B. Primer extension analysis of suf transcription in response to 1 mM H2O2 in oxyR, himA, and himD mutant strains.

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To test whether IHF is necessary for OxyR-dependent induction of the suf operon, we examined suf expression in strains lacking either himA or himD, which encode the subunits of IHF. Deletion of either himA or himD abolished the H2O2 induction of suf by OxyR (Fig. 3B). This suggests that the DNA bending activity of IHF may be required to bring the atypically distant OxyR site closer to RNA polymerase bound at the −10 and −35 promoter elements. There was still some residual activation of suf in response to H2O2 even in the oxyR, himA and himD deletion strains. Because Fe-Fur may be damaged during oxidative stress, the residual transcriptional activation of suf seen in the mutant strains (Fig. 3B) could be the result of loss of Fur repression of suf upon H2O2 exposure. Alternatively, the residual induction could be mediated by an as yet unidentified regulator (Lee et al., 2003).

Independent binding of IHF, OxyR and Fur to the suf promoter

Although genetic evidence indicates that both Fur and IHF regulate suf, it was not known where these two regulatory factors bind to the suf promoter. To localize the IHF and Fur binding sites we carried out DNase I footprinting experiments. These assays showed that IHF binding protects from −121 to −164 of the suf promoter, centred at the predicted binding site between the OxyR site and the promoter (Fig. 3A and Fig. 4). At 100 nM and higher concentrations of IHF, some protection was observed at a second lower affinity site from +7 to −14, which partially overlaps the −10 promoter element (Fig. 4A and data not shown). We observed Fur binding to the predicted Fur box centred between the −10 and −35 promoter elements resulting in DNase I protection of the region from −3 to −32 (Fig. 3A and Fig. 4). No Fur binding was observed elsewhere in the suf promoter even at Fur concentrations of up to 1 µM (Fig. 4, data not shown). Oxidized OxyR bound at a single site located at −197 to −244, as has been previously reported (Zheng et al., 2001).

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Figure 4. DNase I footprinting assays of the suf promoter. The samples were run in parallel with Maxam–Gilbert G/A sequencing ladders. The regions protected by each protein are shown as a solid bar to the left of the footprint. Numbering of the protected regions are relative to +1 of the suf promoter. A. Binding of various amounts of oxidized OxyR, IHF and Fur to the top strand of the suf promoter followed by DNase I treatment. B. Binding of stepwise mixtures of the three proteins at 50 nM levels followed by DNase I treatment. The order of addition is indicated by the label order.

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To determine if binding of one regulatory protein to the suf promoter might interfere with subsequent binding of the other proteins to their mapped sites, we mixed different combinations of IHF, oxidized OxyR, and Fur with the suf promoter fragment before DNase I treatment (Fig. 4B). Prior binding of IHF to the suf promoter did not affect the OxyR or Fur footprint. Addition of OxyR before Fur also did not effect the footprints for either protein. Finally, mixing of IHF, OxyR and Fur did not significantly block or alter the footprints of any of the three proteins. These results indicate that each regulatory protein binds at a distinct site within the suf promoter independent of the other proteins.

Most suf gene products are needed to complement an iscS deletion

The isc operon of E. coli is important for Fe–S cluster assembly in a variety of enzymes, and the iscS deletion mutant shows a range of growth defects including a slow growth phenotype and several auxotrophies (Takahashi and Nakamura, 1999; Schwartz et al., 2000). In contrast, no obvious phenotypes for a sufABCDSE deletion mutant under normal growth in rich or minimal media have been reported. However, a recent study has shown that a plasmid expressing sufABCSE was required to construct an iscRSUA mutant in a sufABCDSE deletion background, indicating that deletion of the two operons is synthetically lethal (Takahashi and Tokumoto, 2002).

To test if individual suf gene products can substitute for deletion of iscS, we measured the co-transduction of an iscS::kanR mutation and a nearby Tn10 insertion into various suf mutant backgrounds. The results revealed that all suf mutations constructed, except sufA, were synthetically lethal with the iscS mutation (Table 1). The synthetic lethality assays suggest that the cysteine desulphurase SufS alone cannot directly substitute for IscS but requires the other suf accessory proteins. In addition, the synthetic lethality observed upon deletion of the sufE or sufBCD genes with iscS indicated that SufE and SufBCD are absolutely essential for proper suf function. Thus the suf gene products are likely to act as a co-dependent complex rather than as separate enzymes. This interpretation is consistent with a recent study that shows that SufB, SufC, and SufD interact as a biochemical complex and that one function for SufE and the SufBCD complex is to enhance the unusually low cysteine desulphurase activity of SufS (Outten et al., 2003). The synthetic lethality phenotype was complemented if the suf operon was provided in trans (Table 1).

Table 1. . Synthetic lethality of iscS and most suf deletion mutants.
StrainCo-transduction frequency (iscS::kanR and zff208-Tn10)
Wild type81%
sufABCDSE 0
sufA79
sufBCD 0
sufB 0
sufD 0
sufSE 0
sufE 0
sufABCDSE/pBADsufABCDSE71

suf mutants are more sensitive to iron starvation than isc mutants

As both suf and isc were induced by iron starvation, we compared the ability of suf and isc deletion mutants to grow under iron starvation conditions. Stationary phase cultures were diluted directly into media containing the iron chelator 2,2′-dipyridyl and allowed to grow for 14 h. Although initial growth rates were similar for the wild-type and sufABCDSE mutant strains, with time, at 2,2′-dipyridyl concentrations of 200 µM or greater, the sufABCDSE mutant strain showed a marked decrease in growth compared to the wild-type strain (Fig. 5A and data not shown). The sufABCDSE mutant also was more sensitive to iron starvation than an iscS mutant despite the pleiotropic growth defects observed for the iscS mutant strain (Fig. 5A). The iron starvation phenotype of the sufABCDSE mutant was observed in a variety of media compositions but was most pronounced in minimal media with gluconate as the carbon source. The iron starvation phenotype could be complemented by the suf operon expressed from a plasmid or by the addition of iron to the media (data not shown). All suf mutants tested showed the same iron starvation phenotype with the exception of sufA, paralleling the results we observed with the synthetic lethality phenotype (Fig. 5B). When exponential phase cultures of the sufABCDSE deletion strain were exposed to 2,2′-dipyridyl, they also showed sensitivity to the iron chelator (data not shown). However, the growth defect exhibited by the exponential phase cells was less severe than the defect observed for the stationary phase cultures.

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Figure 5. Iron starvation sensitivity of suf mutants. Stationary phase cultures of wild-type, iscS::kanR , and sufABCDSE mutant strains (A); wild-type and various suf mutant strains (B); and wild-type, fhuF::kanR, and sufABCDSE mutant strains (C) were diluted into M9 gluconate minimal media containing increasing levels of 2,2′-dipyridyl. Final OD600 measurements were taken after 14 h of growth at 37°C in 2,2′-dipyridyl containing media. Error bars are ± one standard deviation for experiments carried out in triplicate.

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To determine if overexpression of the isc operon might compensate for the sufABCDSE deletion under iron starvation conditions, we compared the sensitivity of a non-polar iscR and sufABCDSE double mutant strain with that of wild-type and sufABCDSE single mutant strains when stationary phase cultures were diluted directly into gluconate minimal media containing 2,2′-dipyridyl. The constitutive overexpression of iscSUA that occurs in an iscR deletion strain (Fig. 2 and Schwartz et al., 2001) was not sufficient to rescue the sufABCDSE phenotype as the iscR sufABCDSE double mutant exhibited the same sensitivity to 2,2′-dipyridyl as the sufABCDSE single mutant (data not shown).

One phenotype reported for sufD or sufS mutant strains is the loss of the FhuF Fe–S centre (Patzer and Hantke, 1999). FhuF, a putative iron reductase, is needed for the utilization of ferrioxamine B or rhodotorulic acid as iron sources (Muller et al., 1998). The exact range of FhuF substrates has not been determined, and it is possible that the FhuF ferric reductase may play a role in releasing iron from other siderophore complexes. To test whether the iron starvation phenotype of the suf mutants could be attributed to loss of FhuF activity, we deleted fhuF and compared the iron starvation phenotype of the fhuF::kanR strain with that of the sufABCDSE deletion strain. The iron starvation phenotype of the suf mutant was more severe than that of the fhuF mutant, which grew as well as the wild-type strain under these conditions (Fig. 5C). Thus the iron starvation sensitivity of a suf deletion strain was not solely caused by loss of FhuF activity, but was the result of a more general growth defect when iron is limiting.

Activity of the Fe–S enzyme 6-phosphogluconate dehydratase is reduced during iron starvation in a suf mutant

Although iron starvation sensitivity also was observed in glucose and succinate minimal media (data not shown), the effects of 2,2′-dipyridyl were most pronounced in gluconate minimal media. Similarly, phenazine methosulphate sensitivity was most pronounced for suf mutants grown with gluconate as the carbon source (Nachin et al., 2003). The enhanced phenotypes seen in gluconate media may be the result of the labile nature of the 4Fe−4S cluster in 6-phosphogluconate dehydratase (Gardner and Fridovich, 1991). Loss of phosphogluconate dehydratase activity would block the Entner–Doudoroff pathway and restrict growth with gluconate as the sole carbon source.

To test this possibility, we measured the specific activity of phosphogluconate dehydratase in wild-type and sufABCDSE deletion strains diluted from stationary phase directly into gluconate minimal media containing 2,2′-dipyridyl (Fig. 6A). Phosphogluconate dehydratase activity decreased more rapidly in response to increasing amounts of 2,2′-dipyridyl in the suf mutant strain compared to the wild-type strain. The activity of a more stable Fe–S enzyme, glutamate synthase, decreased in response to iron starvation but the decrease was the same in the wild-type strain and in the sufABCDSE mutant (Fig. 6A). Decreased gluconate dehydratase activity in the sufABCDSE mutant was not simply the result of decreased levels of gluconate dehydratase protein since Western blot analysis showed similar protein levels in both wild-type and sufABCDSE mutant strains at 2,2′-dipyridyl levels below 250 µM (Fig. 6B).

image

Figure 6. Decreased 6-phosphogluconate dehydratase activity in response to iron starvation. A. Stationary phase cultures of wild-type and sufABCDSE mutant strains were diluted to OD600 = 0.25 directly in gluconate minimal media with various amounts of 2,2′-dipyridyl for 2 h at 37°C. Specific activities of phosphogluconate dehydratase and glutamate synthase were measured in cell extracts after exposure. Error bars are ± one standard deviation for triplicate reactions carried out from each extract. Specific activity is defined as µmol of product formed per min per mg of total protein. B. Levels of 6-phosphogluconate dehydratase protein in 20 µg aliquots from each of the cell extracts used in Fig. 6A were compared via immunoblot with antibodies specific for the enzyme. C. Wild-type and sufABCDSE mutant strains were grown in glucose minimal media at 37°C to mid-exponential phase, harvested, and washed with minimal salts. Cells were then suspended in gluconate minimal media containing 100 µM 2,2′-dipyridyl. Specific activity of phosphogluconate dehydratase was measured in cell extracts at various time-points after suspension.

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To determine if the suf defect in phosphogluconate dehydratase was due to decreased de novo synthesis of Fe–S clusters or to lack of Fe–S cluster repair, we grew wild-type and sufABCDSE deletion strains in glucose minimal media to ensure that transcription of the gluconate utilization pathway was fully repressed (Zwaig et al., 1973; Isturiz et al., 1986). Cells in exponential phase then were harvested and shifted into gluconate media containing 2,2′-dipyridyl so that only the activity of phosphogluconate dehydratase newly synthesized under iron starvation conditions would be measured. Phosphogluconate dehydratase activity increased more slowly and to a lesser degree in the sufABCDSE mutant compared to the wild-type strain at 100 µM 2,2′-dipyridyl, suggesting that de novo synthesis of the 4Fe-4S cluster in phosphogluconate dehydratase is impaired in the sufABCDSE mutant strain under iron starvation conditions (Fig. 6C).

Discussion

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. Supplementary material
  9. References
  10. Supporting Information

Our comparison of the regulation and mutant phenotypes of the isc and suf operons suggests overlapping as well as divergent roles for the two systems. Both operons encode Fe–S cluster assembly proteins and are induced by oxidative stress and iron starvation. The induction of isc by oxidative stress and iron starvation likely reflects a failure in the formation of the Fe–S cluster in the IscR repressor, a regulatory system designed to sense perturbations in the overall Fe–S cluster status of the cell (Schwartz et al., 2001). In contrast, suf is regulated by OxyR and Fur, which are optimized to sense hydrogen peroxide stress and iron starvation, respectively. Thus whereas isc induction reflects disruption of overall Fe–S cluster status and the inability of IscS, IscU and IscA to maintain the cluster of IscR, suf induction is a specific response to oxidative stress and iron starvation.

Though both operons are induced in response to oxidative stress and iron starvation, a deletion of the suf operon renders the cell more sensitive to iron starvation than a deletion of the isc operon. This result shows that the isc-encoded proteins cannot function effectively to synthesize Fe–S clusters when iron is limiting. Thus the physiological roles of the suf and isc operons diverge under iron starvation conditions, explaining the requirement for two separate Fe–S cluster assembly operons in E. coli.

Our finding that the E. coli suf mutants show a greater sensitivity to iron starvation than isc mutants is consistent with the reported sensitivity of Erwinia chrysanthemi suf mutants to iron chelators (Nachin et al., 2003). Those results were interpreted to indicate the loss of a suf-dependent reductase in Erwinia, however, we have determined that the suf iron starvation sensitivity in E. coli is not solely caused by loss of the FhuF reductase. Indeed, we found that a decrease in phosphogluconate dehydratase activity is at least partially responsible for the suf growth defects under iron starvation conditions. We suggest that the suf operon plays an essential role in Fe–S assembly during iron starvation and FhuF is one of multiple Fe–S target enzymes disrupted in a suf mutant under iron limiting conditions. Based on assays of gene reporter constructs and the iron-activated antibiotic streptonigrin, suf mutants in E. chrysanthemi were reported to have elevated intracellular iron levels (Nachin et al., 2001). However, as both of these methods assay changes in intracellular iron availability rather than changes in overall iron accumulation, the results may be difficult to interpret. The reduced pathogenesis of E. chrysanthemi suf mutants (Nachin et al., 2001) and the induction of sufA in Shigella upon entry into host cells (Runyen-Janecky and Payne, 2002) are both consistent with a suf operon role in adaptation to the iron starvation faced by pathogens during infection.

The close interplay between iron homeostasis and oxidative stress complicates the interpretation of defects observed in suf mutants exposed to oxidative stress, such as H2O2 or phenazine methosulphate. One possibility is that the suf-encoded complex functions to repair damaged Fe–S clusters. However, in our assay measuring new synthesis of phosphogluconate dehydratase activity after a shift from glucose to gluconate under iron starvation conditions, the suf deletion strain showed decreased induction, consistent with disruption of de novo synthesis of the 4Fe-4S cluster in the enzyme. The suf operon also does not appear to be directly involved in Fe–S cluster repair under oxidative stress conditions (O. Djaman and J. A. Imlay, unpublished results). Another possibility is that the suf operon is required to synthesize Fe–S clusters in a subset of enzymes distinct from those that require the isc operon. This second hypothesis is weakened by the synthetic lethality observed in the isc and suf double mutants, which implies at least some overlap in the Fe–S synthesis capacity of the two systems.

We favour a third hypothesis that the iron available for Fe–S cluster assembly is limiting during oxidative stress and that the suf operon has a critical function under such conditions. Cellular iron pools are thought to consist of two major components; iron bound tightly by iron proteins, including Fe–S enzymes, haeme proteins, and iron storage proteins, and a more labile pool of iron that is available for the synthesis of iron enzymes or control of iron responsive genes (Woodmansee and Imlay, 2002). The distribution of iron between these two components can be altered upon oxidative stress. The global iron regulator Fur is induced by OxyR in response to H2O2 (Zheng et al., 1999), which may lead to the repression of iron uptake genes. The pool of available iron may be further depleted in E. coli as it is sequestered into tightly binding iron proteins such as Dps. Dps (DNA-binding protein from starved cells) is strongly induced by hydrogen peroxide and during stationary phase and binds iron to form a ferritin-like microcrystalline core with 255 ± 42 atoms per dodecamer (Altuvia et al., 1994; Ilari et al., 2002). Restricting iron uptake and sequestering intracellular iron during H2O2 stress protects the cell from the Fenton reaction whereby Fe2+ reacts with H2O2 to form hydroxyl radicals (Imlay et al., 1988). However, the shift of total cellular iron into storage proteins could reduce the amount of iron available for synthesis of essential iron proteins even as some of these proteins, including Fe–S enzymes, are damaged by H2O2 stress. In this way, H2O2 stress may lead to de facto iron starvation and disrupt synthesis of iron enzymes required for basic metabolic functions.

Expression of non-iron alternative enzymes, such as fumarase C, during oxidative stress may circumvent the de facto iron starvation but not all iron-utilizing enzymes have non-iron alternatives. Therefore, the cell must have mechanisms to channel iron to essential iron proteins during oxidative stress, and Fe–S cluster assembly may require specialized enzymes, such as the suf-encoded proteins, to interface with iron trafficking pathways utilized under these conditions. It has been shown that recovery of Fe–S enzyme activity following challenge with peroxynitrite is reduced in mutants that lack the iron storage proteins ferritin (FtnA) or bacterioferritin (Bfn) (Keyer and Imlay, 1997).

Although we have discussed potential roles for the suf gene products in iron acquisition, it is possible that the limiting component for Fe–S cluster assembly under stress conditions is sulphur. Thus the key role of the suf operon during iron starvation or oxidative stress may actually be to provide a shielded pathway for donating sulfane sulphur to Fe–S clusters. Sulphur availability is likely to be limited as the result of direct oxidation of sulfane sulphur by reactive oxygen species or loss of sulfane sulphur from aborted clusters when iron is limited. Recent work has shown that the transfer of sulfane sulphur from SufS to SufE is more resistant to reductant than transfer of sulphur from IscS to IscU (Urbina et al., 2001; Outten et al., 2003), indicating that the sulphur transfer pathway is shielded in the suf-encoded complex.

In summary, we propose that the suf operon can compensate for the loss of iron or sulphur availability for Fe–S cluster biosynthesis that occurs during both iron starvation and H2O2 stress and is adapted to build Fe–S clusters under these conditions. The suf pathway could acquire iron through direct competition with other cellular chelators for the remaining labile iron pool or through specific interactions between Suf proteins and iron storage or trafficking proteins. Alternatively or in addition, the carefully shielded sulphur transfer pathway between SufS and SufE may be necessary to protect sulfane sulphur from reactive oxygen species and from loss if iron is unavailable. The next challenge is to elucidate the biochemical mechanisms by which suf gene products build Fe–S clusters during iron starvation or oxidative stress.

Experimental procedures

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. Supplementary material
  9. References
  10. Supporting Information

Media

Lennox Broth (10 g of tryptone, 5 g of yeast extract, 5 g of NaCl per litre) was used for experiments with rich media. M9 minimal media with 0.2% carbon source was used for iron starvation sensitivity assays. Initial selection for gene disruption mutants was carried out on TYE plates (10 g of yeast extract, 5 g of tryptone, 8 g of NaCl per litre) with either chloramphenicol (30 µg ml−1) or kanamycin (25 µg ml−1). Other antibiotics used were ampicillin (100 µg ml−1) and tetracycline (6 µg ml−1).

Strain construction

Mutagenesis was performed using the E. coliλ Red targeted mutagenesis system (Datsenko and Wanner, 2000; Yu et al., 2000). Briefly, PCR fragments were amplified from either pKD3 or pKD4 plasmid templates encoding chloramphenicol and kanamycin resistance cassettes respectively (Datsenko and Wanner, 2000). The primers used for targeted gene disruptions are listed in Table S1. The 5′ end of each primer contained 30–40 nucleotides of sequence specific for the targeted gene(s) and the 3′ end of the primer contained 20 nucleotides of sequence specific for the template plasmid. Where necessary, primer sequence was designed to fuse optimized ribosomal binding site and start codon in frame with a downstream gene to construct non-polar mutations. Following transformation of PCR products into DY330, kanamycin- or chloramphenicol-resistant colonies were selected. Mutations were confirmed using colony PCR with primers flanking the mutation. Each mutation was then transduced via P1 phage into MG1655. In some cases antibiotic resistance cassettes were later removed by FLP via plasmid pCP20 as described previously (Datsenko and Wanner, 2000). Removal of antibiotic resistance cassettes was confirmed by colony PCR with flanking primers. Previously described mutations in himA (Granston and Nash, 1993), himD (Flamm and Weisberg, 1985), and iscS (Schwartz et al., 2000) were P1 transduced into the MG1655 background. All strains used in this study are listed in Table 2.

Table 2. . Strains used in this study.
StrainRelevant genotypeReference
MG1655wild-typeLaboratory strain
GSO77oxyR::kanRZheng et al. (2001)
GSO82fur::kanRMukhopadhyay et al. (2004)
GSO92sufABCDSEThis work
GSO93sufAThis work
GSO94sufBThis work
GSO95sufBCDThis work
GSO96sufDThis work
GSO97sufSEThis work
GSO98sufEThis work
GSO99fhuF::kanRThis work
GSO100iscRThis work
GSO101iscS::kanRThis work
GSO102iscS::kanRzff208-Tn10This work
GSO103himA::catRThis work
GSO104himD::catRThis work
DY330λcI857Δ(cro-bioA)Yu et al. (2000)

RNA isolation and primer extension assays

Total RNA was isolated from cell cultures during early exponential growth (OD600 = 0.2–0.3) using either Trizol according to the product protocol (Invitrogen) or acid phenol as previously described (Kawano et al., 2002). Cells in early exponential growth were treated for 5 min with varying amounts of H2O2 or with varying amounts of 2,2′-dipyridyl for 1 h.

Primer extension was conducted on total RNA as described previously (Zheng et al., 2001). For quantitative measurements, gels were exposed to a phosphoimager screen and analysed using a Typhoon 9200 (Amersham Pharmacia) and ImageQuant 5.2 software (Molecular Dynamics). The primers used for primer extension are listed in Table S1.

DNase I footprinting assays

To generate a footprinting probe for the top strand of the suf promoter, the 435 bp BamHI–EcoRV fragment of pGSO133 was labelled with 32P at the BamHI site. DNase I footprinting of oxidized OxyR or IHF binding to the suf promoter was carried out in 25 mM Tris-Cl, pH 7.9, 6 mM MgCl2, 10% glycerol, 0.5 mM EDTA, 0.05% Nonidet P40 (Fluka), and 50 mM KCl. Footprinting with Fur alone and Fur together with IHF was carried out in Fur buffer (10 mM Bis-Tris-HCl, pH 7.5, 5 µg ml−1 sonicated salmon sperm DNA, 5% glycerol, 1 mM MgCl2, 40 mM KCl, 100 µg ml−1 BSA, and 100 µM MnCl2). Footprinting with OxyR and combinations of Fur and IHF was carried out in 25 mM Tris-Cl, pH 7.9, 6 mM MgCl2, 10% glycerol, 0.05% Nonidet P40 (Fluka), and 50 mM KCl. Proteins were allowed to incubate with DNA probe for 15 min at 25°C before DNase I treatment. For footprints containing various mixtures of the three proteins, proteins and DNA probe were allowed to incubate 15 min at 25°C before addition of next protein. Samples were run in parallel with Maxam–Gilbert G/A sequencing ladders. Oxidized OxyR was purified as described previously. IHF protein was kindly provided by H. A. Nash. Fur protein was kindly provided by C. E. Outten and T. V. O’Halloran.

Growth assays

Sensitivity to iron starvation was measured as follows: overnight stationary phase cultures grown in M9 minimal media containing 0.2% gluconate were adjusted to the same starting OD600 and diluted 1:100 in 2 ml of M9 minimal media containing 0.2% gluconate and varying amounts of 2,2′-dipyridyl. Media used for assays with the iscS::kanR strain also contained 12.5 µg ml−1 nicotinic acid. Cells were grown with shaking at 37°C for 14 h and OD600 was measured throughout growth. To assay the effects of added FeCl3, equalized stationary phase cultures were diluted 1:100 into M9/gluconate media with 200 µM 2,2′-dipyridyl and varying amounts of FeCl3. All assays were conducted in triplicate.

Synthetic lethality was assessed by measuring co-transduction frequency of the iscS::kanR. mutation with a nearby Tn10 insertion (zff208-Tn10) into various suf mutant backgrounds. Briefly, iscS::kanRzff208-Tn10 was transduced via P1 phage into MG1655 and various suf mutants followed by initial selection on tetracycline at 30°C. TetR colonies were then tested for kanamycin resistance at 30°C. Co-transduction frequency was calculated by dividing the number of KanR colonies by the total number of TetR colonies. A minimum of 50 colonies was screened for each measurement. The plasmid pGSO164, containing the sufABCDSE operon under the control of the pBAD promoter, was used for complementation studies (Outten et al., 2003). Basal expression from the pBAD promoter was sufficient to complement the suf mutant strains without addition of arabinose.

6-Phosphogluconate dehydratase activity assays and immunoblots

To assay activity in stationary phase cultures, overnight cultures of MG1655 and sufABCDSE (OD600 = 1.3) grown in M9 minimal media containing 0.2% gluconate were diluted to OD600 = 0.25 in M9 minimal media containing 12.5 µg ml−1 nicotinic acid and 0.2% gluconate with various amounts of 2,2′-dipyridyl. After 2 h at 37°C with shaking, cells were harvested by centrifugation at 8000 g for 10 min, washed once in 10 ml of M9 salts, and frozen in liquid nitrogen. Cell pellets were stored at −70°C until use. For time-course of gluconate dehydratase induction, cells were grown in M9 minimal media containing 12.5 µg ml−1 nicotinic acid and 0.2% glucose at 37°C to an OD600 = 0.4–0.5. Cells were collected by centrifugation, washed once with M9 salts, and resuspended in M9 minimal media containing 12.5 µg ml−1 nicotinic acid and 0.2% gluconate. Samples were taken at the indicated time points, harvested, washed, and frozen as described above.

Cell pellets were thawed on ice in an anaerobic chamber (Coy), resuspended in 800 µl of 50 mM Tris-Cl, pH 7.65, and lysed by sonication. Cell lysate (240 µl) was mixed with 30 µl of 100 mM MgCl2. Reactions were initiated by addition of 30 µl of 80 mM 6-phosphogluconate and quenched by addition of 500 µl of boiling Tris buffer followed by boiling for 2–3 min. Pyruvate formed from 6-phosphogluconate and 2-dehydro-3-deoxyphosphogluconate aldolase was determined by mixing 50 µl of reaction with 930 µl of 50 mM Tris-Cl, pH 7.65, 10 µl 20 mM NADH, and 10 µl of 5%l-lactic dehydrogenase. The decrease in absorbance at 340 nm was measured and a millimolar extinction coefficient of 6.22 was used for NADH to calculate the amount of pyruvate present (amount of NADH consumed = amount of pyruvate present in original 5 min reaction). Total protein present in the cell extracts was measured. Specific activity is defined as µmol of product formed per min per mg of total protein.

Twenty microgram aliquots of total protein from each of the cell extracts prepared above were separated on a 4–20% Tris-glycine gel and transferred to a nitrocellulose membrane. The membrane was probed with a 1:5000 dilution of polyclonal antibodies prepared from rabbit specific for 6-phosphogluconate dehydratase that were kindly provided by L. O. Ingram. 6-Phosphogluconate protein levels were visualized using a 1:10 000 dilution of an HRP-conjugated secondary antibody specific for rabbit and the ECL Western Blotting Detection Reagents Kit (Amersham Biosciences).

Glutamate synthase activity assays

Twenty microlitre aliquots from the cell extracts prepared above were mixed with 830 µl 100 mM Tris-Cl, pH 7.8, 50 µl of 0.1 M α-ketoglutarate, 50 µl of 0.2 M l-glutamine, and 50 µl of 4 mM NADPH. The decrease in absorbance at 340 nm was measured over time and a millimolar extinction coefficient of 6.22 was used for NADPH to calculate the amount of l-glutamate formed from α-ketoglutarate and l-glutamine. Total protein present in the cell extracts was measured. Specific activity is defined as µmol of product formed per min per mg of total protein.

Acknowledgements

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. Supplementary material
  9. References
  10. Supporting Information

This work was supported by postdoctoral fellowship from the Pharmacology Research Associate Training Program (to F.W.O.) and by the intramural program at the National Institute of Child Health and Human Development. We would like to thank L. O. Ingram, H. A. Nash, T. V. O’Halloran and C. E. Outten for the kind gifts of reagents, J. A. Imlay for insightful discussions, and J. A. Opdyke, C. E. Outten, and M. J. Wood for critical reading of the manuscript.

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  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. Supplementary material
  9. References
  10. Supporting Information
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Supporting Information

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. Supplementary material
  9. References
  10. Supporting Information

Table S1. Sequences of oligonucleotides used in this study.

FilenameFormatSizeDescription
MMI_4025_sm_tableS1.doc25KSupporting info item

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