How To Deal with Toxic Amino Acids: the Bipartite AzlCD Complex Exports Histidine in Bacillus subtilis

ABSTRACT

The Gram-positive model bacterium Bacillus subtilis can use several amino acids as sources of carbon and nitrogen. However, some amino acids inhibit the growth of this bacterium. This amino acid toxicity is often enhanced in strains lacking the second messenger cyclic dimeric adenosine 3′,5′-monophosphate (c-di-AMP). We observed that the presence of histidine is also toxic for a B. subtilis strain that lacks all three c-di-AMP synthesizing enzymes. However, suppressor mutants emerged, and whole-genome sequencing revealed mutations in the azlB gene that encode the repressor of the azl operon. This operon encodes an exporter and an importer for branched-chain amino acids. The suppressor mutations result in an overexpression of the azl operon. Deletion of the azlCD genes encoding the branched-chain amino acid exporter restored the toxicity of histidine, indicating that this exporter is required for histidine export and for resistance to otherwise toxic levels of the amino acid. The higher abundance of the amino acid exporter AzlCD increased the extracellular concentration of histidine, thus confirming the new function of AzlCD as a histidine exporter. Unexpectedly, the AzlB-mediated repression of the operon remains active even in the presence of amino acids, suggesting that the expression of the azl operon requires the mutational inactivation of AzlB.

IMPORTANCE Amino acids are building blocks for protein biosynthesis in each living cell. However, due to their reactivity and the similarity between several amino acids, they may also be involved in harmful reactions or in noncognate interactions and thus may be toxic. Bacillus subtilis can deal with otherwise toxic histidine by overexpressing the bipartite amino acid exporter AzlCD. Although encoded in an operon that also contains a gene for an amino acid importer, the corresponding genes are not expressed, irrespective of the availability of amino acids in the medium. This suggests that the azl operon is a last resort by which to deal with histidine stress that can be expressed due to the mutational inactivation of the cognate repressor AzlB.

INTRODUCTION

Amino acids are the essential building blocks for protein biosynthesis and for many other cellular components. Cells can acquire amino acids via uptake from the environment, the degradation of external peptides or proteins, or de novo biosynthesis. Many bacteria, such as the model organisms Escherichia coli and Bacillus subtilis, can use all three possibilities for amino acid acquisition. Although amino acids are essential for growth, they can be toxic due to the misloading of tRNAs resulting in misincorporation into proteins and due to their high reactivity. Moreover, many amino acids are chemically similar to each other, and one amino acid that is available in excess may competitively inhibit the biosynthetic pathway(s) of similar amino acids by binding to the corresponding enzymes (1).

We are interested in the identification of the components that allow for the life of a simple minimal cell and in the construction of such cells based on the model bacterium B. subtilis (2, 3). Such minimal organisms not only are important in obtaining a comprehensive understanding of the requirements of cellular life but also are important workhouses in biotechnological and biomedical applications. Indeed, minimal organisms have been proven to be superior to conventionally constructed strains in the production and secretion of difficult proteins and lantibiotics (4–6). For B. subtilis, the pathways for all amino acid biosyntheses have been completely elucidated. In contrast, the knowledge about amino acid transport is far from being complete, as for several amino acids, no transporter has yet been identified. This knowledge is important for the construction of genome-reduced strains that may be designed to grow in a complex or minimal medium and thus require the complete set of uptake or biosynthetic enzymes, respectively. Moreover, some amino acids, such as glutamate, are toxic for B. subtilis, even at the concentrations present in standard complex medium, if the catabolizing enzymes (e.g., glutamate dehydrogenase) are absent (7, 8). Thus, a complete understanding of all components involved in cellular amino acid homeostasis is required for the successful generation of minimal organisms.

Amino acid toxicity not only is relevant for the design of minimal genomes but also is an important tool for the identification of components involved in amino acid metabolism. While some amino acids, such as serine or threonine, are already toxic for wild-type strains (1, 9, 10), others are well-tolerated. In these cases, the corresponding d-amino acids, amino acid analogs, or structurally similar metabolites may act as anti-metabolites that inhibit normal cellular metabolism and thus the growth of the bacteria. The application of toxic amino acids or of similar compounds to a bacterial growth medium will inhibit growth and will also result in the acquisition of suppressor mutations that allow the cells to resolve the issue of amino acid toxicity. Often, such mutations affect uptake systems and prevent the uptake of the toxic amino acid or its analogues. In this way, transporters for threonine, proline, alanine, serine, and glutamate, as well as those for the anti-metabolites 4-hydroxy-l-threonine and glyphosate, have been identified in B. subtilis (9–15). A second way to achieve resistance against toxic amino acids is via the activation of export mechanisms. This has been reported in the cases of 4-azaleucine and glutamate (14, 16). Third, suppressor mutations may facilitate the detoxification of toxic amino acids via the degradation or modification of a nontoxic metabolite, as is observed with glutamate and serine (7, 10, 14, 17). Fourth, the protein target of the toxic metabolite may be modified in such a way that it becomes resistant (18). Finally, the bacteria can escape inactivation via the increased expression of the target protein, as has been reported for serine and the anti-metabolite glyphosate (10, 15).

Recently, it has been shown that the sensitivity of B. subtilis to glutamate is strongly enhanced if the bacteria are unable to produce the second messenger cyclic dimeric adenosine 3′,5′-monophosphate (c-di-AMP) (14). This second messenger is essential for the growth of B. subtilis on a complex medium, and it is toxic upon intracellular accumulation (19). Both the essentiality and the toxicity are mainly results of the central role of c-di-AMP in potassium homeostasis. The second messenger prevents the intracellular accumulation of potassium by inhibiting potassium import and by stimulating potassium export. Thus, the intracellular potassium concentration is kept within a narrow range (19, 20). The presence of high potassium concentrations in a strain lacking c-di-AMP results in the activation of potassium export via the acquisition of mutations in a sodium/H⁺ antiporter. These mutations change the specificity of the antiporter toward potassium (21). Even though none of the known targets of c-di-AMP are directly involved in glutamate homeostasis, glutamate is as toxic as is potassium to the mutant lacking all of the diadenylate cyclases that would synthesize c-di-AMP. This can be explained by the fact that glutamate activates the low-affinity potassium channel KtrCD by strongly increasing the affinity of this channel. Thus, even the low potassium concentration, which must be present for the growth of this strain, becomes toxic due to the high affinity of KtrCD for potassium in the presence of glutamate (22). Accordingly, the Δdac strain lacking c-di-AMP acquires mutations that reduce potassium uptake if propagated in the presence of glutamate. In addition, the bacteria usually acquire additional mutations that interfere with glutamate homeostasis by reducing uptake, facilitating export, or allowing degradation of the amino acid (14).

In this study, we were interested in the control of histidine homeostasis. Histidine biosynthesis from ribose 5-phosphate requires 10 enzymes (see http://subtiwiki.uni-goettingen.de/v4/category?id=SW.2.3.1.14; [23]). The degradation of histidine to ammonia, glutamate, and formamide involves the specific transporter HutM and four enzymes. The histidine transporter is induced in the presence of histidine, which is a typical feature for high-affinity transport systems (24, 25). Usually, high-affinity transporters are used for catabolic pathways to use an amino acid as a carbon or nitrogen source. In contrast, constitutively expressed, low-affinity transporters are used to import amino acids from complex media for protein biosynthesis. In many cases, both low-affinity and high-affinity amino acid transporters are encoded in the genome of B. subtilis, and they are expressed depending on the physiological conditions. However, no low-affinity histidine transporter has yet been identified. Histidine degradation yields intracellular glutamate, which is toxic for mutants lacking c-di-AMP (14) due to the activation of the potassium channel KtrCD (22). Thus, we expected that the strain lacking c-di-AMP would have a similar sensitivity against histidine as it has against glutamate. We made use of this sensitivity to the degradation product glutamate to get further insights into the components that contribute to histidine homeostasis in B. subtilis. Our study revealed that the mutational activation of an export system is the major mechanism by which to achieve resistance to histidine.

RESULTS

Histidine is toxic to a B. subtilis strain lacking c-di-AMP, and mutations in the azlB gene overcome the toxicity.

Some amino acids, such as serine and threonine, are toxic for B. subtilis. In the case of glutamate, toxicity becomes visible in the absence of a degradation pathway or if the bacteria are unable to form the second messenger c-di-AMP (14). Here, we tested the growth of a wild-type strain (168) of B. subtilis and of an isogenic strain that had all three of the genes encoding diadenylate cyclases deleted (Δdac, GP2222) (21) in the presence of histidine. As shown in Fig. 1, the growth of the B. subtilis wild-type strain 168 is not affected by histidine concentrations up to 10 mM. At a higher concentration of 20 mM, growth was inhibited. In contrast, histidine is highly toxic for B. subtilis GP2222, even at low histidine concentrations (see Fig. 1). We observed that larger colonies rapidly appeared. It is likely that these larger colonies were formed by suppressor mutants that were resistant to histidine in the medium. We hypothesized that mutations could affect uptake systems for histidine, as already observed for glutamate, serine, or threonine (9, 10, 14). Indeed, we were able to identify mutations in two isolates via whole-genome sequencing. However, to our surprise, the mutations did not cover the known or putative amino acid transporters of B. subtilis (23). In contrast, we observed mutations in the azlB gene, which encodes a Lrp-type transcription repressor that controls the expression of a branched-chain amino acid exporter (AzlCD) and a branched-chain amino acid importer (BrnQ) (9, 26). Strain GP3638 carried an amino acid substitution in AzlB (Asn24 Ser). In the second strain, GP3639, we found an eight-base pair insertion (CATTAATG) after the 37^th base pair of the coding sequence that results in a frameshift and thus prevents the expression of a functional AzlB protein. As the azlB gene seemed to be a hot spot of mutations in histidine-resistant suppressor mutants, we determined the sequence of this gene in four additional mutants. In each case, mutations were present in the azlB gene, either as amino acid substitutions (e.g., Asn24Ser as in GP3638, Ile31Met) or as frameshift mutations. Since the frameshift mutations prevent the formation of functional AzlB proteins, it seemed likely that the amino acid substitutions also resulted in inactive proteins. Indeed, both the N24S and the I31M mutations affect the DNA-binding helix-turn-helix motif of AzlB.

FIG 1.

The isolated suppressors are resistant to histidine stress. Growth of B. subtilis suppressor mutants (GP3638, GP3639 and GP3588) in the presence of histidine. All suppressors carry different mutations in the azlB gene (see Table 2). The cells were harvested and washed, and the OD₆₀₀ was adjusted to 1.0. Serial dilutions were added dropwise to MSSM minimal plates with the indicated histidine concentration. The plates were incubated at 42°C for 48 h.

As mentioned above, the growth of the wild-type strain 168 was inhibited above 20 mM histidine. Therefore, we tested the growth of our suppressor mutants in the absence and presence of histidine. While all mutants were viable at 5 mM histidine, they were still inhibited at a concentration of 30 mM (see Fig. 1). However, when suppressor mutants originally isolated at 15 mM histidine were transferred to 30 mM histidine, suppressor mutants appeared again. One of these mutants (GP3588) was subjected to whole-genome sequencing. In coherence with our previously isolated mutants, we found a frameshift mutation in azlB, highlighting the importance of azlB inactivation for the adaptation of the B. subtilis strain lacking c-di-AMP to the presence of histidine. Moreover, we found three additional mutations at a histidine concentration of 30 mM. Both the main potassium transporter KimA and the KtrD membrane subunit of the low-affinity potassium channel KtrCD (21, 27) were inactivated due to frameshift mutations. In addition, the high-affinity glutamate transporter GltT (14, 28) carried a substitution of Thr-342 to a Pro residue. It is known that KtrCD is converted to a high-affinity potassium channel in the presence of glutamate (22), suggesting that glutamate, as the product of histidine utilization, causes activation of KtrCD. Moreover, small amounts of glutamate that are exported from the cell may be reimported by GltT, thus, again, contributing to the activation of KtrCD. This activation of KtrCD, as well as the activity of KimA, contributes to potassium toxicity that can only be bypassed via the inactivation of the major potassium uptake systems.

Histidine toxicity in the Δdac mutant GP2222 might be caused by the formation of glutamate that triggers toxic glutamate accumulation, by the toxicity of histidine due to its chemical reactivity, or by a combination of both. The identification of kimA and ktrD mutants in the suppressor isolated at the elevated histidine concentration suggests that potassium toxicity really can become a problem for the bacteria. However, we never identified suppressor mutants that were affected in the histidine degradation pathway, thus avoiding the problem of intracellular glutamate formation. To test the role of histidine degradation for the acquisition of resistance to histidine, we deleted the hutH gene encoding histidase, the first gene of the catabolic pathway in the wild-type and a Δdac mutant strain. The set of four isogenic strains was tested for growth on minimal medium in the absence of histidine and in the presence of 5 mM, 15 mM, 25 mM, and 35 mM histidine. While all strains grew well in the absence of histidine, growth was inhibited at histidine concentrations exceeding 15 mM and 5 mM for the wild-type and the Δdac mutant, respectively. The inactivation of the histidine degradative pathway (hutH mutation) did not affect growth in either of the genetic backgrounds (data not shown). Thus, growth inhibition of the Δdac mutant by histidine seems to result from a combination of (i) its own chemical reactivity, as has also been observed for E. coli (29), and (ii) its conversion to glutamate, which triggers toxic potassium accumulation in this strain.

Taken together, our results demonstrate that the Lrp-type repressor protein AzlB plays a major role in the adaptation of B. subtilis lacking c-di-AMP to high levels of histidine. At even higher concentrations of histidine, the degradation product glutamate induces the uptake of potassium, which is known to be toxic to strains that are unable to produce c-di-AMP (20–22).

Suppressor mutants exhibit increased expression of the azlBCD-brnQ operon.

To test the effect of the azlB mutations on the expression of the azlBCD-brnQ operon, we analyzed the transcripts of the operon via a Northern blot analysis. For this purpose, we cultivated the wild-type strain 168, the Δdac mutant GP2222, and two suppressor mutants GP3638 (AzlB-Asn24 Ser) and GP3639 (frameshift in AzlB) in modified sodium Spizizen minimal medium (MSSM), isolated the RNA, and performed Northern blot experiments using a riboprobe that is complementary to azlC to detect the specific mRNA(s). Based on the known transcript sizes of the B. subtilis glycolytic gapA operon (30), we estimated the sizes of the transcripts of the azl operon. As shown in Fig. 2A, expression of the operon could not be detected in the wild-type strain 168 or in the Δdac mutant GP2222. Signals corresponding to transcripts of about 1,100, 1,500, 3,300 and 5,100 nucleotides were only visible in the two suppressor mutants carrying the azlB mutations. This result indicates that only inactive azlB allowed for the expression of the azl operon and led to high expression levels. The presence of multiple transcripts suggests internal transcription signals and/or mRNA processing events.

FIG 2.

AzlCD is strongly overexpressed in the histidine suppressors. (A) Northern blot analysis to test the expression levels of the azl operon in the suppressor mutants GP3638 and GP3639. The RNA was isolated from MSSM minimal medium during the exponential growth phase. The gapA probe was used together with wild-type RNA as a control to estimate band sizes and strength, as it is strongly expressed under normal conditions. (B) Transcriptional organization of the azl operon. The length of the individual transcripts is indicated by the arrows. The positions of the transcripts that are smaller than the complete operon mRNA are deduced from the determined transcript sizes and the genomic arrangement. The bent arrow and the hairpin correspond to the positions of the transcriptional upshifts and downshifts, respectively.

So far, the inducer for the azlBCD-brnQ operon has not been identified. Since our results indicate that the operon is involved in the control of histidine homeostasis, we wanted to test the activity of the azlB promoter under different conditions. For this purpose, we fused the azlB promoter region to a promoterless lacZ gene encoding β-galactosidase and integrated this azlB-lacZ fusion into the B. subtilis genome. According to a genome-wide transcriptome analysis (31), the promoter of the operon is located in front of the upstream yrdF gene. However, the same study indicated the presence of an mRNA upshift in front of the azlB gene. Similarly, Belitsky et al. identified promoter activity immediately upstream of azlB (26). To clarify this issue, we also constructed and tested a yrdF-lacZ fusion. The strains carrying the azlB-lacZ and yrdF-lacZ fusions were cultivated in C-Glc minimal medium in the absence or presence of different amino acids as potential inducers. As shown in Table 1, both of the upstream regions of yrdF and azlB had only minor promoter activity. As a control, we used the moderately expressed promoter of the pgi gene, which encodes phosphoglucose isomerase. This promoter yielded 10-fold higher β-galactosidase activity compared to the yrdF and azlB promoters. In addition, the activity of the yrdF and azlB promoters was not induced by any of the tested amino acids, including histidine. Therefore, we also tested casein hydrolysate, a mixture of amino acids. Again, no induction was observed for either promoter. However, the deletion of the azlB gene resulted in an approximately 7-fold increase of the activity of the azlB promoter (see GP3614 versus GP3612), whereas the yrdF promoter was not affected. Moreover, GltR, a LysR family transcription factor of so far unknown function, is encoded downstream of the brnQ gene (32). Therefore, we considered the possibility that GltR might play a role in the control of the azl operon. However, the deletion of the gltR gene did not affect the activity of the azlB promoter.

TABLE 1.

Activity of the azlB promoter

		Units of β-galactosidase per μg of protein Addition to C-Glc minimal medium
Strain	Relevant genotype	None	Ile	Pro	His	CAA
GP314	pgi-lacZ	48 ± 3	NDa	ND	ND	ND
GP3612	azlB-lacZ	4 ± 0.6	3 ± 0.7	2 ± 0.2	4 ± 0.6	7 ± 1.5
GP3614	azlB-lacZ ΔazlB	26 ± 5	25 ± 2	31 ± 4	31 ± 4	49 ± 7
GP3617	azlB-lacZ ΔgltR	3 ± 0.3	ND	ND	ND	5 ± 0.1
GP3611	yrdF-lacZ	3 ± 0.7	4 ± 1	2 ± 0.3	5 ± 2	ND
GP3613	yrdF-lacZ ΔazlB	3 ± 0.3	3 ± 0.6	3 ± 0.1	3 ± 0.4	ND

^aND, not determined.

Taken together, our data confirm that AzlB is the transcriptional repressor of the azl operon. The azlB gene is the first gene of the operon (see Fig. 2B). Moreover, our results demonstrate that the transcriptional regulation by AzlB is not affected by any individual amino acid or by a mixture of them, even though the operon encodes exporters and importers for amino acids. Only the loss of a functional AzlB repressor allows for the expression of the azl operon (see Discussion).

Resistance to histidine depends on the AzlCD amino acid exporter.

So far, we have established that the suppressor mutants have mutations in AzlB that increase the expression of the azl operon, which confers resistance to histidine. In addition to the promoter-proximal repressor gene azlB, this operon encodes the AzlC and AzlD subunits of a bipartite amino acid exporter and the branched-chain amino acid transporter BrnQ as well as the YrdK protein of unknown function and the putative 4-oxalocrotonate tautomerase YrdN. Since the overexpression of AzlCD was also responsible for the resistance of B. subtilis to azaleucine (26), it seemed most plausible that this transporter would also be involved in histidine resistance. To test this hypothesis, we constructed two sets of isogenic strains that differed in the azl operon in the background of the wild-type 168 and in the background of the Δdac mutant GP2222. First, we compared growth of the wild-type, the azlB mutant GP3600, and the azlBCD mutant GP3601. As shown in Fig. 3A, the wild-type strain was sensitive to the presence of 15 mM histidine in the medium, whereas the isogenic azlB mutant that exhibits overexpression of AzlCD was resistant. However, the additional deletion of the azlCD genes in GP3601 restored the sensitivity to histidine, indicating that the increased expression of the AzlCD amino acid exporter is responsible for the acquired resistance to histidine. Similar results were obtained for the set of strains that are unable to synthesize c-di-AMP (Δdac) (Fig. 3B). Again, the strain lacking AzlB (GP3607) was resistant to high levels of histidine (20 mM), whereas the strain lacking the amino acid exporter AzlCD in addition to AzlB (GP3606) was as sensitive as the Δdac mutant (GP2222) even at 5 mM histidine. Ectopic expression of the azlCD genes under the control of the constitutive degQ36 promoter (33) in strain GP3642 that lacks the endogenous azlBCD operon partially restored the resistance to histidine up to a concentration of 5 mM. In contrast, the expression of the AzlC component of the bipartite exporter alone had no effect (GP3643) (Fig. 3B). Taken together, these data strongly suggest that the overexpression of the two-component amino acid exporter AzlCD as a result of the inactivation of AzlB is required for the resistance of B. subtilis to histidine.

FIG 3.

The azlB mutation confers resistance to histidine stress. (A) Sensitivity of wild type B. subtilis (168) and the ΔazlB (GP3600) and ΔazlBCD (GP3601) mutants to histidine. The cells were grown in MSSM minimal medium to an OD₆₀₀ of 1.0 and were then diluted 10-fold to create dilutions ranging from 10⁻¹ to 10⁻⁶. The dilution series was dropped onto MSSM plates with and without (15 mM) histidine. The plates were incubated at 37°C for 48 h. (B) Growth of Δdac Δazl (GP3606), Δdac ΔazlB (GP3607), and Δdac Δazl complemented with azlC (GP3643) and azlCD (GP3642), respectively. Δazl indicates a deletion of the azlBCD genes. The cells were grown as described above. The plates were incubated at 42°C for 48 h.

Overexpression of AzlCD results in enhanced histidine export.

AzlCD has previously been identified as an exporter for 4-azaleucine and was hypothesized to be an exporter for other branched-chain amino acids (26). Our data suggest that the complex might also export histidine and thus might contribute to histidine resistance upon overexpression. To test this idea, we determined the relative intracellular and extracellular histidine concentrations in the wild-type strain 168 and in the isogenic azlB, azlBCD, and azlCD deletion mutants GP3600, GP3601, and GP3622, respectively, during growth in MSSM minimal medium. In this condition, de novo histidine biosynthesis is active, as MSSM minimal medium does not contain amino acids. Compared to the wild type, the intracellular histidine levels decreased in the azlB mutant GP3600, thus confirming that the higher AzlCD levels in this strain led to histidine export (Fig. 4A). Mutants lacking the amino acid exporter AzlCD had wild-type-like histidine levels (Fig. 4A). In contrast, the extracellular histidine concentration was 3-fold higher in the azlB mutant, whereas the strains lacking AzlCD had extracellular histidine levels that were comparable to those of the wild-type strain (Fig. 4B). These results demonstrate that AzlCD, which is overexpressed as a result of the azlB mutation, is involved in the control of histidine homeostasis. While the loss of AzlCD has no effect, which corresponds to a lack of expression in the wild-type strain, its overexpression results in reduced and increased intracellular and extracellular histidine levels, respectively. This suggests that AzlCD is an active histidine exporter.

FIG 4.

AzlCD is a histidine exporter in B. subtilis. Box and whisker plot of the intracellular (A) and extracellular (B) histidine levels of B. subtilis ΔazlB, ΔazlBCD, and ΔazlCD mutants relative to the wild-type strain 168. The red lines indicate the median values of 12 biological replicates. The upper box edges show the 75^th percentiles, and the lower box edges show the 25^th percentiles. The whiskers indicate the furthest points that are not considered outliers. The red crosses indicate outliers. Differences between the indicated pairs of strains were tested for statistical significance using a Wilcoxon rank sum test at a significance level (α) of 0.05. P values of <0.05 were considered to be indicative of statistically significant results. The asterisks indicate the orders of magnitude of the P values: *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001.

DISCUSSION

The results presented in this study demonstrate that histidine inhibits the growth of B. subtilis, as has already been shown for serine or threonine (10, 34, 35). Amino acid toxicity is often enhanced if B. subtilis is unable to produce the essential second messenger nucleotide c-di-AMP due to the activation of the potassium channel KtrCD by glutamate, the degradation product of many amino acids (14, 22). This work shows that the increased sensitivity of a strain lacking c-di-AMP to amino acids is also valid for histidine.

Typically, suppressor screens using toxic amino acids, amino acid analogs or related anti-metabolites result in the identification of transporters that have been inactivated in the suppressor mutants (9–15). While this is the predominant type of suppressor mutation, resistance to toxic amino acids and related molecules can also be achieved by the activation of degradation pathways (10, 14), by the activation of export mechanisms (14, 26), or by modifying the target protein/pathway in such a way that it becomes insensitive to the presence of the otherwise toxic molecule. This was observed for glyphosate resistance in Salmonella typhimurium, which can be achieved by mutations that render the target enzyme 5-enolpyruvyl-shikimate-3-phosphate (EPSP) synthase insensitive to inhibition (18), as well as for serine toxicity in B. subtilis, which could be overcome via the increased expression of the genes encoding the threonine biosynthetic pathway (10). Studies about histidine toxicity in E. coli revealed that the amino acid enhances oxidative DNA damage (29). Thus, one might also expect suppressor mutations that prevent DNA damage. The exclusive isolation of azlB mutations that activate the expression of the AzlCD amino acid exporter suggests that all other mechanisms of suppression are more difficult to achieve for the B. subtilis cell.

The fact that we were unable to isolate a single suppressor mutant that had lost histidine uptake strongly suggests that B. subtilis possesses multiple histidine transporters. So far, only the HutM histidine transporter has been identified, based on its similarity to known histidine transporters (25). It is tempting to speculate that the genome of B. subtilis encodes one or more low-affinity transporters for histidine. Indeed, B. subtilis encodes several homologs of the Pseudomonas putida histidine transporter HutT (36). These transporters all belong to the amino acid-polyamine-organocation (APC) superfamily of amino acid transporters. Four of them (AapA, AlaP, YbxG, and YdgF) share more than 40% sequence identity with P. putida HutT, suggesting that these proteins have the same biological activity. Thus, the presence of multiple histidine uptake systems would prevent the rapid simultaneous inactivation of all of these systems in suppressor mutants, which explains why no transporter mutants were isolated.

Our data clearly demonstrate that the bipartite amino acid transporter AzlCD exports not only the leucine analog 4-azaleucine (16) but also histidine. Corresponding bipartite systems that mediate the export of branched-chain amino acids have also been identified in E. coli and Corynebacterium glutamicum (37, 38). These exporters are members of the LIV-E class of transport proteins (38, 39). As in B. subtilis, these systems consist of a large subunit (corresponding to AzlC) and a small subunit (corresponding to AzlD). While proteins homologous to AzlC are abundant in a wide range of bacteria, including most Actinobacteria and Firmicutes as well as many Proteobacteria, AzlD is conserved in only a few bacteria. The other bacteria that possess a homolog of AzlC obviously have alternative small subunits. This is the case in E. coli, where the small YgaH subunit of the YgaZ/YgaH valine exporter is not similar to its counterparts in B. subtilis and C. glutamicum. We have also considered the possibility that the large subunit AzlC might be sufficient for histidine export; however, this is not the case (see Fig. 3B).

It is interesting to note that the AzlCD amino acid exporter is able to export multiple amino acids. Substrate promiscuity is a common feature in amino acid transport. In B. subtilis, the low affinity transporter AimA is the major transporter for glutamate and serine (10, 14). Similarly, the BcaP permease transports branched-chain amino acids, threonine, and serine (9–11), and the GltT protein is involved in the uptake of aspartate, glutamate, and the antimetabolite glyphosate (14, 15, 28). Thus, AzlCD is another example of the weak substrate specificity of amino acid transporters. It is tempting to speculate that AzlCD might even be involved in the export of other amino acids and related metabolites in B. subtilis.

Based on the chemical properties of each amino acid, it may be generally toxic or toxic only under specific conditions. Therefore, cells often have efficient degradation pathways by which to remove toxic compounds. This is the case for glutamate, which is degraded by the glutamate dehydrogenases GudB or RocG (7, 17). However, other amino acids become toxic only at high concentrations or in particular mutant backgrounds. This is the case for histidine, which is toxic only at high concentrations for the B. subtilis wild-type strain, but can render a strain unable to form c-di-AMP at low concentrations. Similarly, the presence of amino acid analogs, such as 4-azaleucine, might be a rather exceptional event in natural environments. Still, B. subtilis is equipped to meet this challenge via the amino acid exporter AzlCD. Based on a global transcriptome analysis, the azl operon is barely expressed under a wide range of conditions, and no conditions that result in the induction of the operon could be detected (26, 31). Similarly, the putative arginine and lysine exporter YisU was not expressed under any of 104 studied conditions (31). The observation that the expression of the azl operon in the presence of toxic concentrations of histidine or 4-azaleucine is obviously not sufficient to provide resistance against these amino acids suggests that none of these compounds acts as a molecular inducer for the azl operon. In agreement with previous results (26), we observed substantial expression of the operon only if the azlB gene encoding the repressor of the operon was deleted or inactivated due to suppressor mutations. Even the presence of a mixture of amino acids derived from casamino acids did not result in the induction of the operon. As the functions of the operon seem to be related to amino acid export (AzlCD) and uptake (BrnQ), regulation via amino acid availability seemed to be the most likely. However, the results from prior global and operon-specific transcription studies and from our data suggest that the activity of AzlB is not controlled by amino acids, even though the protein belongs to the Lrp family of leucine-responsive regulatory proteins (40). It is tempting to speculate that AzlB has lost the ability to interact with amino acid-related effector molecules, but the expression of the operon can rapidly be activated via the acquisition of mutations that inactivate AzlB. Alternatively, AzlB might respond to a yet unknown signal and then allow for the induction of the operon. The mutational inactivation of a normally silent operon has also been described for the cryptic E. coli bgl operon for the utilization of β-glucosides, which requires the insertion of the mobile element IS5 in the promoter region to get expressed (41).

Due to its strongly increased sensitivity to several amino acids, the B. subtilis mutant lacking c-di-AMP is an excellent tool by which to study mechanisms of amino acid homeostasis and to identify uptake and export systems. This endeavor is required, as the details of amino acid transports are one of the few areas which have several gaps of knowledge in the research on B. subtilis (3). We anticipate that the further use of the c-di-AMP-lacking mutant will continue to help fill these remaining gaps.

MATERIALS AND METHODS

Strains, media, and growth conditions.

E. coli DH5α (42) was used for cloning. All B. subtilis strains used in this study are derivatives of the laboratory strain 168. They are listed in Table 2. B. subtilis and E. coli were grown in Luria-Bertani (LB) or in sporulation (SP) medium (42, 43). For the growth assays, B. subtilis was cultivated in MSSM medium (21). MSSM is a modified SM medium in which KH₂PO₄ is replaced by NaH₂PO₄ and KCl is added as indicated (21). The media were supplemented with ampicillin (100 μg/mL), kanamycin (10 μg/mL), chloramphenicol (5 μg/mL), spectinomycin (150 μg/mL), tetracycline (12.5 μg/mL), or erythromycin plus lincomycin (2 and 25 μg/mL, respectively) if required.

TABLE 2.

B. subtilis strains used in this study

Strain	Genotype	Source or Reference
168	trpC2	Laboratory collection
GP314	trpC2 amyE::(Ppgi-lacZ cat)	54
GP983	trpC2 ΔcdaS::ermC	49
GP997	trpC2 ΔcdaA::cat	49
GP2032	trpC2 ΔcdaA::cat ΔcdaS::ermC	GP997 → GP983
GP2142	trpC2 ΔdisA::tet	This study
GP2222	trpC2 ΔcdaA::cat ΔcdaS::ermC ΔdisA::tet	21
GP2715	trpC2 ΔdisA::spc	This study
GP2782	trpC2 ΔdisA::kan	This study
GP3588	trpC2 ΔcdaA::cat ΔcdaS::ermC ΔdisA::tet azlB fs(pos 126 +T) ktrD fs(pos 72 –T) kimA fs(pos 128 +GTCGCAT) gltT (Thr342 Trp)	Suppressor of GP2222 (30 mM His)
GP3600	trpC2 ΔazlB::spec	This study
GP3601	trpC2 ΔazlBCD::kan	This study
GP3604	trpC2 ΔcdaA::cat ΔcdaS::ermC ΔazlBCD::kan	GP3601 → GP2032
GP3605	trpC2 ΔcdaA::cat ΔcdaS::ermC ΔazlB::spec	GP3600 → GP2032
GP3606	trpC2 ΔcdaA::cat ΔcdaS::ermC ΔdisA::spec ΔazlBCD::kan	GP2715 → GP3604
GP3607	trpC2 ΔcdaA::cat ΔcdaS::ermC ΔdisA::kan ΔazlB::spec	GP2782 → GP3605
GP3611	trpC2 amyE::(PyrdF-lacZ aphA3)	pGP3807 → 168
GP3612	trpC2 amyE::(PazlB-lacZ aphA3)	pGP3808 → 168
GP3613	trpC2 ΔazlB::spec amyE::(PyrdF-lacZ aphA3)	pGP3807 → GP3600
GP3614	trpC2 ΔazlB::spec amyE::(PazlB-lacZ aphA3)	pGP3808 → GP3600
GP3615	trpC2 ΔgltR::spec	This study
GP3617	trpC2 ΔgltR amyE::(pazlB-lacZ aphA3)	pGP3808 → GP3615
GP3622	trpC2 ΔazlCD::kan	This study
GP3623	trpC2 ΔazlBCD::spec	This study
GP3625	trpC2 ΔcdaA::cat ΔcdaS::ermC ΔazlBCD::spec	GP3623 → GP2032
GP3626	trpC2 ΔcdaA::cat ΔcdaS::ermC ΔazlBCD::spec ganA::(pdegQ36-azlC aphA3)	pGP3811 → GP3625
GP3627	trpC2 ΔcdaA::cat ΔcdaS::ermC ΔazlBCD::spec ganA::(pdegQ36-azlCD aphA3)	pGP3812 → GP3625
GP3638	trpC2 ΔcdaA::cat ΔcdaS::ermC ΔdisA::tet azlB (Asn24 Ser)	Suppressor of GP2222 (15 mM His)
GP3639	trpC2 ΔcdaA::cat ΔcdaS::ermC ΔdisA::tet azlB fs (pos 37 +CATTAATG)	Suppressor of GP2222 (15 mM His)
GP3642	trpC2 ΔcdaA::cat ΔcdaS::ermC ΔdisA::tet ΔazlBCD::spec ganA::(pdegQ36-azlCD aphA3)	GP2142 → GP3626
GP3643	trpC2 ΔcdaA::cat ΔcdaS::ermC ΔdisA::tet ΔazlBCD::spec ganA::(pdegQ36-azlC aphA3)	GP2142 → GP3627
GP4202	trpC2 ΔhutH::spec	This study
GP4205	trpC2 ΔhutH::spec ΔcdaA::cat ΔcdaS::ermC	GP4202 → GP2032
GP4206	trpC2 ΔhutH::spec ΔcdaA::cat ΔcdaS::ermC ΔdisA::kan	GP2782 → GP4205

DNA manipulation and transformation.

All commercially available restriction enzymes, T4 DNA ligase, and DNA polymerases were used as recommended by the manufacturers. DNA fragments were purified using the QIAquick PCR Purification Kit (Qiagen, Hilden, Germany). The DNA sequences were determined by the dideoxy chain termination method (42). Standard procedures were used to transform E. coli (42), and the transformants were selected on LB plates containing ampicillin (100 μg/mL). B. subtilis was transformed with plasmid or chromosomal DNA, according to the two-step protocol described previously (43). Transformants were selected on SP plates containing chloramphenicol (Cm 5 μg/mL), kanamycin (Km 10 μg/mL), spectinomycin (Spc 150 μg/mL), tetracycline (Tet 12,5 μg/mL), or erythromycin plus lincomycin (2 μg/mL and 25 μg/mL, respectively).

Genome sequencing.

To identify the mutations in the suppressor mutant strains GP3588, GP3638, and GP3639 (see Table 2), the genomic DNA was subjected to whole-genome sequencing. The concentration and purity of the isolated DNA was first checked with a Nanodrop ND-1000 (PeqLab Erlangen, Germany), and the precise concentration was determined using the Qubit dsDNA HS Assay Kit as recommended by the manufacturer (Life Technologies GmbH, Darmstadt, Germany). Illumina shotgun libraries were prepared using the Nextera XT DNA Sample Preparation Kit and were subsequently sequenced on a MiSeq system with the Reagent Kit v3 with 600 cycles (Illumina, San Diego, CA, USA) as recommended by the manufacturer. The reads were mapped on the reference genome of B. subtilis 168 (GenBank accession number: NC_000964) (44). The mapping of the reads was performed using the Geneious software package (Biomatters Ltd., New Zealand) (45). Frequently occurring hitchhiker mutations (46) and silent mutations were omitted from the screen. The resulting genome sequences were compared to that of our in-house wild-type strain. Single nucleotide polymorphisms were considered significant when the total coverage depth exceeded 25 reads with a variant frequency of ≥90%. All identified mutations were verified by PCR amplification and Sanger sequencing.

Construction of mutant strains by allelic replacement.

The deletion of the azlB, azlBCD, azlCD, disA, gltR, and hutH genes was achieved via the transformation of B. subtilis 168 with a PCR product constructed using oligonucleotides to amplify the DNA fragments flanking the target genes and an appropriate intervening resistance cassette as described previously (47). The integrity of the regions flanking the integrated resistance cassette was verified by sequencing PCR products of approximately 1,100 bp that were amplified from the chromosomal DNA of the resulting mutant strains. In the case of the azlB, azlCD, and azlBCD deletions, the cassette carrying the resistance gene lacked a transcription terminator to ensure the expression of the downstream genes.

Phenotypic analysis.

In B. subtilis, amylase activity was detected after growth on plates containing 7.5 g/L nutrient broth, 17 g/L Bacto agar (Difco) and 5 g/L hydrolyzed starch (Connaught). Starch degradation was detected by sublimating iodine onto the plates.

Quantitative studies of lacZ expression in B. subtilis were performed as follows. Cells were grown in MSSM medium supplemented with KCl at different concentrations as indicated. The cells were harvested at an OD₆₀₀ of 0.5 to 0.8. β-galactosidase-specific activities were determined with cell extracts obtained via lysozyme treatment as described previously (43). One unit of β-galactosidase is defined as the amount of enzyme that produces 1 nmol of o-nitrophenol per min at 28°C.

To assay the growth of B. subtilis mutants at different histidine concentrations, a drop dilution assay was performed. Briefly, precultures in MSSM medium at the indicated histidine concentration were washed three times and resuspended to an OD₆₀₀ of 1.0 in a MSSM basal salts solution. A dilution series was then pipetted onto MSSM plates containing the desired histidine concentration.

Plasmid constructions.

The plasmid pAC7 (48) was used to construct translational fusions of the potential yrdF and azlB promoter regions to the promoterless lacZ gene. For this purpose, the promoter regions were amplified using oligonucleotides that attached EcoRI and BamHI restriction to the ends of the products. The fragments were cloned between the EcoRI and BamHI sites of pAC7. The resulting plasmids were pGP3807 and pGP3808 for yrdF and azlB, respectively.

To allow for the ectopic expression of the azlC and azlCD genes, we constructed the plasmids pGP3811 and pGP3812, respectively. The corresponding genes were amplified using oligonucleotides that added BamHI and PstI sites to the ends of the fragments and cloned into the integrative expression vector pGP1460 (49) that was linearized with the same enzymes.

Northern blot analysis.

The strains B. subtilis 168 (wild type) and GP2222 (Δdac mutant), as well as the suppressor mutants GP3638 and GP3639, were grown in MSSM minimal medium and were harvested in the late logarithmic phase. The preparation of the total RNA and the Northern blot analysis were carried out as described previously (50, 51). Digoxigenin (DIG) RNA probes were obtained via in vitro transcription with T7 RNA polymerase (Roche Diagnostics) using PCR-generated DNA fragments as the template. The reverse primer contained a T7 RNA polymerase recognition sequence. In vitro RNA labeling, hybridization, and signal detection were carried out according to the instructions of the manufacturer (DIG RNA Labeling Kit and detection chemicals; Roche Diagnostics).

Determination of intracellular and extracellular histidine pools.

For the determination of the histidine levels of B. subtilis, cells were cultivated in MSSM minimal medium until the exponential growth phase (OD₆₀₀ of 0.4). For the extraction of intracellular metabolites, 4 mL of each culture were harvested by filtration (52). Histidine levels were then determined as described previously (53), using ¹³C labeled histidine from an E. coli extract as the internal standard. Briefly, an Agilent 1290 Infinity II UHPLC system (Agilent Technologies) was used for liquid chromatography. The column was an Acquity BEH Amide 30 × 2.1 mm with a particle size of 1.7 μm (Waters GmbH). The temperature of the column oven was 30°C, and the injection volume was 3 μL. The LC solvent A was water with 10 mM ammonium formate and 0.1% formic acid (vol/vol), and the LC solvent B was acetonitrile with 0.1% formic acid (vol/vol). The gradient was 0 min with 90% B, 1.3 min with 40% B, 1.5 min with 40% B, 1.7 min with 90% B, 2 min with 90% B, and 2.75 min with 90% B. The flow rate was 0.4 mL/min. From minute 1 to minute 2, the sample was injected into the MS. An Agilent 6495 triple quadrupole mass spectrometer (Agilent Technologies) was used for mass spectrometry. The source gas temperature was set to 200°C, with 14 l min⁻¹ drying gas and a nebulizer pressure of 24 lb/in². The sheath gas temperature was set to 300°C, and the flow was set to 11 l min⁻¹. The electrospray nozzle and capillary voltages were set to 500 and 2,500 V, respectively. Isotope-ratio mass spectrometry with a ¹³C internal standard was used to obtain relative data. Fully ¹²C-labeled histidine and ¹³C-labeled histidine was measured via multiple reaction monitoring in the positive ionization mode, using a collision energy of 13 eV. The precursor ion masses were 156 Da and 162 Da, and the product ion masses were 110 Da and 115 Da for ¹²C-histidine and ¹³C-histidine, respectively. The ratios between ¹²C-labeled histidine and ¹³C-labeled histidine were normalized to the ODs and to the median ratio of the control strain 168.