Investigating the composition and recruitment of the mycobacterial ImuA′–ImuB–DnaE2 mutasome

  1. Sophia Gessner
  2. Zela Alexandria-Mae Martin
  3. Michael A Reiche
  4. Joana A Santos
  5. Ryan Dinkele
  6. Atondaho Ramudzuli
  7. Neeraj Dhar
  8. Timothy J de Wet
  9. Saber Anoosheh
  10. Dirk M Lang
  11. Jesse Aaron
  12. Teng-Leong Chew
  13. Jennifer Herrmann
  14. Rolf Müller
  15. John D McKinney
  16. Roger Woodgate
  17. Valerie Mizrahi
  18. Česlovas Venclovas
  19. Meindert H Lamers
  20. Digby F Warner  Is a corresponding author
  1. SAMRC/NHLS/UCT Molecular Mycobacteriology Research Unit, DSI/NRF Centre of Excellence for Biomedical TB Research, Department of Pathology, University of Cape Town, South Africa
  2. Institute of Infectious Disease and Molecular Medicine, University of Cape Town, South Africa
  3. Laboratory of Microbiology and Microsystems, School of Life Sciences, Swiss Federal Institute of Technology in Lausanne (EPFL), Switzerland
  4. Advanced Imaging Center, Howard Hughes Medical Institute, United States
  5. Department of Cell and Chemical Biology, Leiden University Medical Center, Netherlands
  6. Department of Integrative Biomedical Sciences, University of Cape Town, South Africa
  7. Confocal and Light Microscope Imaging Facility, Department of Human Biology, University of Cape Town, South Africa
  8. Helmholtz Centre for Infection Research, Helmholtz Institute for Pharmaceutical Research Saarland, Germany
  9. German Centre for Infection Research (DZIF), Partner Site Hannover-Braunschweig, Germany
  10. Laboratory of Genomic Integrity, Eunice Kennedy Shriver National Institute of Child Health and Human Development, United States
  11. Wellcome Centre for Infectious Diseases Research in Africa, University of Cape Town, South Africa
  12. Institute of Biotechnology, Vilnius University, Lithuania

Abstract

A DNA damage-inducible mutagenic gene cassette has been implicated in the emergence of drug resistance in Mycobacterium tuberculosis during anti-tuberculosis (TB) chemotherapy. However, the molecular composition and operation of the encoded ‘mycobacterial mutasome’ – minimally comprising DnaE2 polymerase and ImuA′ and ImuB accessory proteins – remain elusive. Following exposure of mycobacteria to DNA damaging agents, we observe that DnaE2 and ImuB co-localize with the DNA polymerase III β subunit (β clamp) in distinct intracellular foci. Notably, genetic inactivation of the mutasome in an imuBAAAAGG mutant containing a disrupted β clamp-binding motif abolishes ImuB–β clamp focus formation, a phenotype recapitulated pharmacologically by treating bacilli with griselimycin and in biochemical assays in which this β clamp-binding antibiotic collapses pre-formed ImuB–β clamp complexes. These observations establish the essentiality of the ImuB–β clamp interaction for mutagenic DNA repair in mycobacteria, identifying the mutasome as target for adjunctive therapeutics designed to protect anti-TB drugs against emerging resistance.

Editor's evaluation

This important study investigates the localization dynamics of the mycobacterial mutasome complex, comprised of ImuA', ImuB, and DnaE2. The mutasome complex has a key role in promoting mutagenic DNA replication during stress to increase the mutation rate and potential for selection of drug resistant mutations. The authors provide compelling evidence that ImuB localizes with the β-clamp upon damage exposure and that the clamp binding motif in ImuB is essential for its localization. These studies lay the ground for future work in this area and will be intriguing to a broad audience interested in bacterial physiology.

https://doi.org/10.7554/eLife.75628.sa0

Introduction

Mycobacterium tuberculosis, the causative agent of tuberculosis (TB), consistently ranks among the leading infectious killers worldwide (World Health Organization, 2021). The heavy burden imposed by TB on global public health is exacerbated by the emergence and spread of drug-resistant (DR) M. tuberculosis strains, with estimates indicating that DR-TB now accounts for approximately one-third of all deaths owing to antimicrobial resistance (Hasan et al., 2018). In the absence of a wholly protective vaccine, a continually replenishing pipeline of novel chemotherapeutics is required (Evans and Mizrahi, 2018) which, given the realities of modern antibiotic development (Nielsen et al., 2019), appears unsustainable. Therefore, alternative approaches must be explored including the identification of effective multidrug combinations (Cokol et al., 2017), the elucidation of ‘resistance-proof’ compounds (Kling et al., 2015), and the identification of so-called ‘anti-evolution’ drugs that might limit the development of drug resistance (Smith and Romesberg, 2007; Ragheb et al., 2019; Merrikh and Kohli, 2020).

Whereas many bacterial pathogens accelerate their evolution by sampling the immediate environment – for example, via fratricide, natural competence, or conjugation (von Wintersdorff et al., 2016; Veening and Blokesch, 2017) – these mechanisms appear inaccessible to M. tuberculosis: the bacillus does not possess plasmids (Gray and Derbyshire, 2018) and there appears to be no role for horizontal gene transfer in the modern evolution of strains of the M. tuberculosis complex (Galagan, 2014; Boritsch and Brosch, 2016). Instead, genetic variation in M. tuberculosis results exclusively from chromosomal rearrangements and mutations, a feature reflecting its ecological isolation (an obligate pathogen, M. tuberculosis has no known host outside humans) and the natural bottlenecks that occur during transmission (Gagneux, 2018). A question which therefore arises is whether a specific molecular mechanism(s) drives M. tuberculosis mutagenesis – perhaps under stressful conditions – and, consequently, if the activity thereof might be inhibited pharmacologically.

Multiple studies have investigated mycobacterial DNA replication and repair function in TB infection models (for recent reviews, Singh, 2017; Minias et al., 2018; Mittal et al., 2020). From these, the C-family DNA polymerase, DnaE2, has emerged as major contributor to mutagenesis under antibiotic treatment (Boshoff et al., 2003). A non-essential homolog of E. coli DNA Polymerase (Pol) IIIα (Timinskas et al., 2014), DnaE2 does not operate alone: the so-called ‘accessory factors’, imuA′ and imuB, are critical for DnaE2-dependent mutagenesis (Warner et al., 2010). Both proteins are of unknown function, however imuA′ and imuB are upregulated together with dnaE2 following exposure of mycobacteria to DNA damaging agents including mitomycin C (MMC). That observation prompted the proposal that the three proteins might represent a ‘mycobacterial mutasome’ – named according to its functional analogy with the E. coli DNA Pol V mutasome comprising UmuD′2C-RecA-ATP (Jiang et al., 2009; Erdem et al., 2014).

Here, we apply live-cell fluorescence and time-lapse microscopy in characterizing a panel of mycobacterial reporter strains expressing fluorescent translational fusions of each of the known mutasome components. The results of these analyses, together with complementary in vitro biochemical assays utilizing purified mycobacterial proteins, support the inference that ImuB serves as a hub protein, interacting with the dnaN-encoded mycobacterial β clamp and ImuA′. They also reinforce the essentiality of the ImuB–β clamp protein–protein interaction for mutasome function. Notably, while a strong ImuA′–ImuB interaction is detected in vitro, our live-cell data indicate the dispensability of either ImuA′ or DnaE2 for ImuB localization – but not mutasome function – in bacilli exposed to genotoxic stress. Finally, using the β clamp-binding antibiotic, griselimycin (GRS) (Kling et al., 2015), we demonstrate in biochemical assays and in live mycobacteria the capacity to inhibit mutasome function through the pharmacological disruption of ImuB–β focus formation. These observations suggest that, through its inhibition of β clamp binding, GRS might naturally limit the capacity for induced mutagenesis. As well as revealing a built-in mechanism protecting against auto-induced mutations to GRS resistance, our results therefore imply the potential utility of ‘anti-evolution’ antibiotics for TB.

Results

ImuB forms distinct subcellular foci under DNA damaging conditions

Our previous genetic evidence (Warner et al., 2010) informed a tentative model in which the presumed catalytically inactive Y family Pol homolog, ImuB, functioned as an adapter protein. According to the model, DnaE2 gains access to the repair site by interacting with ImuB, which similarly interacts with ImuA′ and the dnaN-encoded β clamp subunit. To investigate the subcellular localizations of each of the mutasome proteins in live bacilli, we constructed reporter alleles in which the M. smegmatis mutasome proteins were labeled by N-terminal translational attachment of either Enhanced Green (EGFP) or Venus Fluorescent Protein (VFP) tags. The reporter alleles were introduced into each of three individual M. smegmatis mutasome gene deletion mutants – ΔdnaE2, ΔimuA′, and ΔimuB (Warner et al., 2010) – to yield the fluorescently tagged complemented strains, ΔdnaE2 attB::egfp-dnaE2 (strain designated G-DnaE2, carrying G-dnaE2 allele), ΔimuB attB::egfp-imuB (G-ImuB), and ΔimuA′ attB::vfp-imuA′ (V-ImuA′) (Figure 1—figure supplement 1A).

The mycobacterial DNA damage response was induced by exposing the strains to the natural product antibiotic, MMC, an alkylating agent that causes monofunctional DNA adducts and inter- and intra-strand cross-links (Bargonetti et al., 2010). Following exposure of G-ImuB to MMC for 4 hr, distinct EGFP-ImuB foci were observed (Figure 1A). In contrast, a yellow fluorescence signal was observable throughout V-ImuAʹ cells, suggesting diffuse distribution of the VFP-ImuA′ protein in the mycobacterial cytoplasm (Figure 1B). Although less distinct than G-ImuB, EGFP-DnaE2 produced similar evidence of focus formation in G-DnaE2 cells (Figure 1C). Notably, the significant increase in signal detectable in V-ImuA′, G-ImuB, and G-DnaE2 cells following MMC exposure (Figure 1—figure supplement 1B) confirmed that expression of the respective fluorescence reporter alleles was DNA damage dependent in all three complemented mutants.

Figure 1 with 1 supplement see all
Visualization of the mycobacterial mutasome components.

Representative stills from fluorescence microscopy experiments of M. smegmatis expressing translational reporters of the different mutasome components in their respective knockout backgrounds. Phase-contrast and fluorescence images of M. smegmatis expressing (i) G-imuB, (ii) V-imuA′, and (iii) G-dnaE2 alleles are represented following 4 hr exposure to ultra-violet (UV) and 1× minimun inhibitory concentration (MIC) mitomycin C (MMC). White boxes indicate zoomed-in regions shown in the panels at right. The far right-hand panels indicate the fluorescence intensity determined along the longitudinal axis of a representative cell from each reporter mutant; the specific cell analyzed is outlined in the corresponding image to the left of the graph. Fluorescence microscopy experiments were repeated two to four times. Scale bars, 5 µm. Source data are available in Figure1.zip which can be accessed at http://doi.org/10.5061/dryad.76hdr7szc.

To ascertain if these observations were true for other types of DNA damage, the three reporter mutants were subjected to ultra-violet (UV) light exposure. Equivalent fluorescence phenotypes were observed for each of the three reporter alleles under both DNA damaging treatments (Figure 1). As UV exposure causes cyclobutane pyrimidine dimers or pyrimidine–pyrimidone (6–4) photoproducts (Boshoff et al., 2003), while MMC generates inter-strand DNA cross-links at CpG sites (Tomasz, 1995), these results indicated that expression and localization (recruitment) of the mutasome components might be independent of the nature of the genotoxic stress applied.

N-terminal fluorescent reporters retain wild-type mutagenic function but are deficient in DNA damage tolerance

The addition of bulky fluorescent tags can disrupt the function of DNA replication and repair proteins (Renzette et al., 2005). To determine if any of the tagged mutasome proteins was affected, the functionalities of the egfp-imuB, vfp-imuA′, and egfp-dnaE2 alleles were assessed in two standard assays (Boshoff et al., 2003; Warner et al., 2010): the first investigated DNA damage-induced mutagenesis by measuring the frequency of rifampicin (RIF) resistance following exposure to genotoxic stress, and the second tested DNA damage tolerance by spotting serial dilutions of each strain on media containing a DNA damaging agent. As observed previously (Boshoff et al., 2003; Warner et al., 2010), exposure of the wild-type parental M. smegmatis mc2155 to a sub-lethal dose of UV irradiation increased the frequency of RIF resistance 50- to 100-fold, as determined from enumeration of colony-forming units (CFU) on RIF-containing solid growth medium. In contrast, induced mutagenesis was greatly reduced in the ΔimuA′, ΔimuB, and ΔdnaE2 deletion mutants, with mutation frequencies for these ‘mutasome-deficient’ strains approximately 20-fold lower than wild-type (Figure 2A). Notably, complementation with the cognate fluorescent reporter allele in V-ImuA′, G-ImuB, and G-DnaE2 restored the UV-induced mutation frequencies of the three respective knockout mutants to near wild-type levels, establishing that each of the fluorescence reporter alleles retained function in UV-induced mutagenesis assays. In assays utilizing MMC instead of UV, a similar 20-fold reduction in MMC-induced mutagenesis was observed in each of the three single knockout strains compared to wild-type, and this defect was restored when complemented with the respective fluorescent reporters (Figure 2B). In combination, these results confirmed the preservation of wild-type mutagenic function in the fluorescently tagged fusion proteins, irrespective of DNA damaging agent applied.

Figure 2 with 1 supplement see all
Functional validation of translational reporters.

(A) N-terminally tagged fluorescence reporter mutants of M. smegmatis ImuA′, ImuB, and DnaE2 retain function in DNA damage-induced mutagenesis. Cultures of M. smegmatis deletion mutants and complemented derivatives were exposed to 25 mJ/cm2 of 254 nm ultra-violet (UV) light and allowed to recover for 3 hr before selection of rifampicin (RIF)-resistant mutants on RIF-containing 7H10 solid agar plates. (i) Mutation frequencies were calculated as a fraction of the CFU/ml of each culture prior to exposure to UV irradiation. Complementation with the corresponding fluorescence reporter alleles restored the resistance frequencies of the three mutasome knockout mutants (ΔimuA′, ΔimuB, and ΔdnaE2) to levels observed in wild-type M. smegmatis. (ii) Representative RIF-containing plates with RIF-resistant mutants. (B) The same strains were exposed to 0.5× MIC mitomycin C (MMC) for 6 hr before plating on RIF-containing 7H10 solid plates. (i) Mutation frequencies were calculated as a fraction of the CFU/ml of each culture prior to exposure to MMC. As for the UV-induced mutagenesis assay, the fluorescence reporter alleles restored mutation frequencies to wild-type levels. (ii) Representative images of the RIF-containing plates with RIF-resistant mutants. (C) Serial dilutions of M. smegmatis deletion mutants and complemented strains were spotted on standard 7H10 and MMC-containing 7H10 plates. Results represent a minimum of three replicates for each strain. Source data are available in Figure2.zip which can be accessed at http://doi.org/10.5061/dryad.76hdr7szc.

Surprisingly, the DNA damage tolerance assay – in which CFU-forming ability was tested during continuous exposure to MMC in solid growth media – produced contrasting results (Figure 2C): whereas the damage hypersusceptibility of the dnaE2 knockout was reversed in the G-DnaE2 strain, complementation of either ΔimuA′ or ΔimuB with its corresponding fluorescent reporter allele failed to restore a wild-type phenotype. The reason for these discrepant observations – restoration of both UV- and MMC-induced mutagenesis but not MMC-induced DNA damage tolerance – in the V-ImuA′ and G-imuB strains is not clear. Although mutasome components are expressed in response to genotoxic stress arising from a variety of different sources, it is possible the different types and/or extent of DNA damage induced in the two separate assays used here (induced mutagenesis vs. DNA damage tolerance) might require distinct interactions with a different partner protein(s) and, further, that one/more of these might have been disrupted by the presence of the fluorescent tag(s). It is also plausible that, in the DNA damage survival assay, extended incubation in the presence of MMC (a clastogen with multiple effects on DNA integrity) might exacerbate the suboptimal operation of the mutasome owing to the presence of the bulky fluorophore – which differs significantly from the very brief exposure to the genotoxins in the induced mutagenesis assays. Consistent with the proposed impact of treatment duration on the functionality of the fluorescently tagged mutasome fusions, both V-ImuA′ and G-ImuB mutants phenocopied wild-type in a UV damage sensitivity assay (Figure 2—figure supplement 1); however, these explanations are speculative and require further investigation. Given the inferred functionality of the fluorescence-tagged alleles in DNA damage-induced mutagenesis, we deemed them useful to investigate mutasome recruitment in live mycobacterial cells.

ImuB localizes with the dnaN-encoded β clamp following DNA damage

We previously inferred that a putative interaction between ImuB and the dnaN-encoded β clamp was essential for mutasome function (Warner et al., 2010). To investigate the predicted interaction of ImuB and the β clamp in live bacilli, each of the three mutasome reporter alleles was introduced separately into an M. smegmatis mutant encoding an mCherry-tagged β clamp, mCherry-DnaN (Santi et al., 2013). The mCherry-DnaN reporter was chosen as background strain owing to its previous validation in single-cell, time-lapse fluorescence microscopy analyses of M. smegmatis replisome location (Santi et al., 2013; Santi and McKinney, 2015). For the time-lapse experiments, the resulting M. smegmatis dual reporter strains were grown in standard 7H9/OADC medium for 12 hr, following which the cells were exposed to MMC for 4.5 hr before switching back to 7H9/OADC for post-treatment recovery (Figure 3; Videos 13). At 4 hr post MMC treatment, distinct EGFP-ImuB foci were observed which, when overlaid with the mCherry-DnaN fluorescence signal, showed considerable overlap, suggesting association of the β clamp with ImuB (Figure 3A, D; Video 1). In addition to G-ImuB, the number of mCherry-DnaN foci also increased upon DNA damage (Figure 3; Figure 3—figure supplement 1A). In MMC-treated cells, the EGFP-ImuB signal was mostly detected in very close proximity to mCherry-DnaN foci (>50% of cells contained mCherry-DnaN and G-ImuB located within 0.3 μm of each other); almost the same frequency of association of mCherry-DnaN and G-ImuB foci was observed in bacilli exposed to UV, though the proportion of cells containing mCherry-DnaN foci alone was greater (Figure 3A, D; Figure 3—figure supplement 1B). In combination, these results are consistent with the direct physical interaction of ImuB and the β clamp suggested previously by yeast two-hybrid and site-directed mutagenesis studies (Warner et al., 2010).

Figure 3 with 2 supplements see all
Representative time-lapse series of single cells of M. smegmatis expressing the mutasome reporters in combination with mCherry-DnaN.

(A) G-ImuB (green) and mCherry-DnaN (magenta), (B) V-ImuA′ (green) and mCherry-DnaN (magenta), and (C) G-DnaE2 (green) and mCherry-DnaN (magenta). Overlapping signals are viewed as white. The cells were exposed to 0.5× MIC MMC from time 0 hr until 4.5 hr, after which the medium was switched back to standard 7H9/OADC medium. Up to 80 XY points were imaged at 10-min intervals on fluorescence and phase channels for up to 36 hr. The experiments were repeated two to four times. Numbers indicate hours elapsed; scale bars, 5 μm. 7H9, Middlebrook 7H9 medium; MMC, mitomycin C. (D) Population-scale analysis of cells with both mCherry-DnaN foci and G-ImuB foci showed distinct overlap in location suggesting co-occurrence of the respective proteins. Source data are available in Figure3.zip which can be accessed at http://doi.org/10.5061/dryad.76hdr7szc.

Video 1
Time-lapse microscopy of G-ImuB and mCherry-DnaN dual reporter.

Representative time-lapse movie of the reporter strain expressing G-ImuB and mCherry-DnaN. Bacteria were imaged on fluorescence and phase channels for up to 36 hr at 10-min intervals. Treatment with MMC (100 ng/ml) was at 0–4.5 hr. This experiment was repeated six times. Numbers indicate the hours elapsed in the time-lapse experiment. 7H9, Middlebrook 7H9/OADC; MMC, mitomycin C. Scale bar, 5 μm. G-ImuB, green; mCherry-DnaN, magenta; overlay, white.

Video 2
Time-lapse microscopy of V-ImuA′ and mCherry-DnaN dual reporter.

Representative time-lapse movie of the reporter strain expressing V-ImuA' and mCherry-DnaN. Bacteria were imaged on fluorescence and phase channels for up to 36 hr at 10-min intervals. Treatment with MMC (100 ng/ml) was at 0–4.5 hr. This experiment was repeated three times. Numbers indicate the hours elapsed in the time-lapse experiment. 7H9, Middlebrook 7H9/OADC; MMC, mitomycin C. Scale bar, 5 μm. V-ImuA', green; mCherry-DnaN, magenta; overlay, white.

Video 3
Time-lapse microscopy of G-DnaE2 and mCherry-DnaN dual reporter.

Representative time-lapse movie of the reporter strain expressing G-DnaE2 and mCherry-DnaN. Bacteria were imaged on fluorescence and phase channels for up to 36 hr at 10-min intervals. Treatment with MMC (100 ng/ml) was at 0–4.5 hr. This experiment was repeated three times. Numbers indicate the hours elapsed in the time-lapse experiment. 7H9, Middlebrook 7H9/OADC; MMC, mitomycin C. Scale bar, 5 μm. G-DnaE2, green; mCherry-DnaN, magenta; overlay, white.

For V-ImuA′, a diffuse fluorescence signal was detected throughout the cells (Figure 3B; Video 2), rendering impossible any conclusion about the potential recruitment of ImuA′ to β clamp (mCherry-DnaN) foci. In contrast, the results for DnaE2 were more nuanced: overlap of peak fluorescence signals from EGFP-DnaE2 and mCherry-DnaN proteins was detected (Figure 3C) and was most evident within 1-hr post removal of MMC from the microfluidic chamber (Video 3). Although not as consistent as the ImuB–β clamp phenotype, the co-occurrence of DnaE2 and β clamp signals was reproducibly observed in multiple cells and across different experiments.

ImuA′ and DnaE2 are not required for ImuB focus formation

We showed previously that deletion of imuA′ phenocopied abrogation of either imuB or dnaE2 in the MMC sensitivity assay (Warner et al., 2010) and, consistent with the interpretation that all three components are individually essential for mutasome activity, this phenotype was not exacerbated in a triple ΔimuA′–imuB–ΔdnaE2 knockout strain. Together with yeast two-hybrid data which indicated a direct interaction between ImuB and ImuA′ (Warner et al., 2010), this observation raised the possibility that a deficiency in ImuA′ might impair ImuB protein localization. To test this prediction, the egfp-imuB allele was introduced into the ΔimuA′ deletion mutant, generating a ΔimuA′ attB::egfp-imuB reporter strain. Despite the absence of ImuA′ in this mutant, EGFP-ImuB foci were observed following treatment with MMC (Figure 3—figure supplement 2A). Similarly, the absence of functional DnaE2 had no discernible impact on ImuB focus formation in either the site-directed dnaE2AIA attB::egfp-imuB strain (Figure 3—figure supplement 2B) or the fully DnaE2-deleted ΔdnaE2 attB::egfp-imuB mutant (Figure 3—figure supplement 2C). In combination, these results appear to eliminate a role for either ImuA′ or DnaE2 in ImuB localization, instead implying the critical importance of the ImuB–β clamp interaction for mutasome assembly.

Purified mutasome proteins interact in biochemical assays in vitro

All inference from this and previous work about the composition of the mycobacterial mutasome has been derived from microbiological assays. To address this limitation, we expressed and purified recombinant M. smegmatis mutasome proteins for biochemical analysis. Expression in E. coli of ImuB alone yielded low quantities of soluble protein that was prone to degradation, while attempts to express ImuA′ alone failed to generate soluble protein. In contrast, co-expression of ImuB with ImuA′ yielded both proteins in a soluble form (Figure 4). Subsequently, the ImuA′B complex could be captured via a histidine (His) affinity tag in ImuB. This confirmed that ImuA′ and ImuB interact in vitro, forming a stable complex even at protein concentrations as low as 400 nM (Figure 4—figure supplement 1A), corroborating previous yeast two-hybrid results (Warner et al., 2010). In E. coli, overexpression of DnaE2 resulted in insoluble protein, while DnaE2 overexpression in M. smegmatis appeared to be incompatible with cell viability: following transformation with the expression construct, very few colonies were obtained and could not be expanded in liquid culture (not shown).

Figure 4 with 1 supplement see all
ImuB and ImuA′–ImuB interact with DnaN and these interactions are disrupted by griselimycin (GRS).

(A) Gel filtration profiles of M. smegmatis (i) ImuA′B-DnaN and (ii) ImuB-DnaN complexes in the absence or presence of 15 μM GRS. For these experiments, 5 μM DnaN was added to 10 μM of (i) ImuA′B or (ii) ImuB. The gel filtration profiles of the individual proteins (ImuB and DnaN) or complex (ImuA′–ImuB) are shown for comparative purposes, and all curves were scaled for clarity. (B) Sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS–PAGE) analysis of sequential fractions of the gel filtration runs. Gels are sorted in the same order as the corresponding gel filtration profiles shown in A. Source data are available in Figure4.zip which can be accessed at http://doi.org/10.5061/dryad.76hdr7szc.

Next, we analyzed the interaction of the dnaN-encoded β clamp with ImuB or the ImuA′B complex (Figure 4). Samples of the M. smegmatis β clamp with ImuA′B (Figure 4A, panel i) or ImuB (Figure 4A, panel ii) were injected into an analytical size-exclusion chromatography column and collected fractions subjected to sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS–PAGE) analysis. Alone, the β clamp and ImuB/ImuA′B eluted at 1.47 and 1.54 ml, respectively. Incubation of the β clamp with either ImuB or ImuA′B caused a shift in the retention volume to 1.36 ml, indicative of complex formation. This was confirmed by SDS–PAGE analysis, which indicated co-elution of the β clamp with ImuB and ImuA′B (Figure 4B).

EGFP-ImuB and VFP-ImuA′ form a stable complex

Our microbiological assays had unexpectedly revealed discrepant complementation phenotypes for the induced mutagenesis versus DNA damage tolerance assay (Figure 2), raising the possibility that the fluorescent tags in the bioreporter mutants might disrupt a protein–protein interaction(s) essential for DNA damage tolerance. We therefore investigated the capacity of the fluorescently labeled EGFP-ImuB and VFP-ImuA′ proteins to form a stable complex. To this end, His-EGFP-ImuB was co-expressed with Strep-VFP-ImuA′ in E. coli and the complex analyzed in three consecutive chromatography steps (Figure 4—figure supplement 1B). First, the cell lysate was loaded onto a HisTrap column to capture the VFP-ImuA′:EGFP-ImuB complex via the His-tag present in EGFP-ImuB. Next, the elution fractions containing the complex were loaded on a StrepTrap column to capture the complex via the strep-tag on VFP-ImuA′. Finally, the VFP-ImuA′:EGFP-ImuB complex was injected onto a size-exclusion column.

During all purification steps, EGFP-ImuB and VFP-ImuA′ were co-eluted as a complex, as indicated by SDS–PAGE analysis and fluorescent detection of EGFP-ImuB and VFP-ImuA′ in the same elution fractions. In combination, these observations suggest that the fluorescent tags did not disrupt ImuA′–ImuB complex formation in vitro – a result which implies that the absence in live cells of a clear ImuA′ (co-)localization phenotype was not attributable to the presence of N-terminal fluorophores.

Inhibition of ImuB–β clamp-binding eliminates focus formation

Previous work established that the β clamp-binding domain of ImuB was essential for mutasome function: mutant strains carrying either a imuBΔC168 allele (lacking the 168 amino acids in the ImuB C-terminal region) or a imuBAAAAGG allele (in which the wild-type β clamp-binding motif, 352QLPLWG357, is substituted with the non-functional 352AAAAGG357 peptide sequence) phenocopied full imuB deletion (Warner et al., 2010). Therefore, to test the prediction that the recruitment of EGFP-ImuB and mCherry-DnaN into discernible foci was dependent on the ImuB–β clamp protein–protein interaction, we introduced an egfp-imuBAAAAGG allele (G-imuBAAAAGG) into the ΔimuB mutant. In contrast to the wild-type reporter (G-ImuB), the β clamp-binding motif mutant (G-ImuBAAAAGG) exhibited no EGFP foci in any cell imaged following exposure to MMC (Figure 5A). Instead, the fluorescence was detectable throughout the cell as a diffuse signal. This result supports the inferred essentiality of the physical interaction between ImuB and β for ImuB localization and, moreover, establishes that detection of ImuB–β foci provides a reliable visual proxy for functional mutasome formation.

Figure 5 with 1 supplement see all
Disrupting the ImuB–β clamp interaction.

(A) Representative images of G-ImuB exposed to 2× MIC mitomycin C (MMC) for 4 hr (top panel) or 2× MIC MMC plus griselimycin (GRS) for 4 hr (center panel), and the G-ImuBAAAAGG mutant exposed to 2× MIC MMC for 4 hr (bottom panel). Scale bars, 5 μm. (B) Interactions of M. smegmatis β clamp with (i) GRS, (ii) ImuB, or (iii) the replicative DNA PolIIIα subunit, DnaE1. The interaction of the β clamp with GRS is represented by the X-ray structure of the complex (PDB id: 5AGU). Predicted interactions with ImuB (S347-I359) and DnaE1 (M947-G954) are derived from the respective AlphaFold models. Interacting peptides are shown as smoothed traces with side chains. The brown β clamp region indicates residues in contact with corresponding peptides. Molecular contacts were derived from 3D structures using the VoroContacts web server. Detailed contact data are provided separately (Supplementary file 1 – DnaN Contact Data). (C) Cells aligned by mid-cell position, arranged according to cell length and colored (magenta – mCherry-DnaN foci, green – G-ImuB foci) according to fluorescence intensity, showing the presence of G-ImuB foci following MMC treatment and the lack of foci after GRS exposure. G-ImuBAAAAGG shows no foci after MMC treatment, similar to the G-ImuB strain following GRS exposure. (D) Proportions of cells containing either mCherry-DnaN or G-ImuB foci following exposure to GRS alone at 2× MIC or 5× MIC, or in combination with either single-dose ultra-violet (UV) or MMC at 2× MIC or 5× MIC. Additional Source data are available in Figure5.zip, accessible at http://doi.org/10.5061/dryad.76hdr7szc.

GRS blocks ImuB–β clamp binding, preventing focus formation in M. smegmatis

GRS is a natural product antibiotic that binds the mycobacterial β clamp with high affinity, preventing DNA replication by blocking the essential interaction with the PolIIIα subunit, DnaE1 (Kling et al., 2015). Importantly, the region of GRS binding on β overlaps with the region predicted to interact with other β clamp-binding proteins (Bunting et al., 2003; Burnouf et al., 2004; Kling et al., 2015), including DnaE1 and ImuB (Figure 5B). From structural comparisons, this also holds true for M. tuberculosis (Figure 5—figure supplement 1). Therefore, we hypothesized that GRS might disrupt the ImuB–β interaction. Indeed, addition of GRS disrupted the in vitro interaction between the β clamp and pre-formed ImuA′B complex (Figure 4A, panel i) as well as between the β clamp and ImuB (Figure 4A, panel ii), as indicated by a gel filtration profile that is a superposition of the absorbance traces of the sample individual components (β clamp and ImuB or β clamp and ImuA′B). This was confirmed by SDS–PAGE analysis (Figure 4B). To confirm that the disrupting effect of GRS on the complex was the result of the GRS–β clamp binding (Kling et al., 2015), we measured the melting curves of the β clamp in the presence and absence of GRS (Figure 4—figure supplement 1C). Incubation with GRS led to a 3°C increase in the protein melting temperature, consistent with GRS binding to β. In contrast, GRS had no effect on the observed melting temperature of ImuA′B.

Finally, we examined whether these biochemical observations were recapitulated in vivo in live mycobacterial cells. To this end, the dual reporter mutant expressing mCherry-DnaN and G-ImuB was treated with GRS alone or following induction of DNA damage by UV or MMC exposure (Figure 5). Notably, the addition of GRS in combination with UV or MMC markedly reduced G-imuB focus formation (Figure 5C, panels ii, iii), with most cells phenocopying the diffuse fluorescence distribution observed following exposure of the β clamp-binding deficient EGFP-ImuBAAAAGG mutant to UV or MMC (Figure 5A). Population analyses confirmed that GRS blocked ImuB focus formation in both UV- and MMC-exposed cells (Figure 5D), although the effect appeared more pronounced for the UV-damaged cells. The reasons for this difference are not clear. It is possible that the variety of monofunctional DNA adducts, inter- and intra-strand cross-links caused by MMC (Bargonetti et al., 2010) elicits a more profound DNA damage response than UV, which results in photoproducts and dimers; moreover, unlike UV, which is delivered as a transient exposure, MMC is a chemical clastogen which might persist inside mycobacterial cells before eventual elimination. Consistent with this proposal, it was evident that MMC treatment caused an elevated number of G-ImuB foci compared to UV – and GRS seemed more effective at reducing UV-induced mCherry-DnaN and G-ImuB foci than the corresponding MMC-induced foci, perhaps owing to ‘trapping’ of foci by MMC-induced DNA cross-links. Whatever the reason, the ability of GRS to inhibit G-ImuB focus formation in live mycobacterial cells exposed to two different DNA damaging agents suggested the potential for chemical disruption of mutasome function.

Discussion

In E. coli, the DNA damage-induced SOS response triggers overexpression of umuC, umuD, and recA (Maslowska et al., 2019). UmuC is an error prone Y-family DNA polymerase that requires the binding of UmuD'2, RecA, and ATP to reach full activity; this multi-protein ‘mutasome’, collectively referred to as DNA PolV, has been implicated in DNA damage tolerance and induced mutagenesis (Goodman et al., 2016). At the time of initiating the work reported here, genetic evidence from diverse bacteria lacking PolV homologs supported the co-dependent operation of ImuA, ImuB, and DnaE2 in the LexA-regulated SOS response, suggesting these proteins might function in an analogous manner (McHenry, 2011; Ippoliti et al., 2012). In mycobacteria, in which they have been individually implicated in DNA damage tolerance and induced mutagenesis (Boshoff and Mizrahi, 2000; Warner et al., 2010), the ImuA homolog, ImuA′, replaces ImuA. Nevertheless, the inferred universal model for mutasome function in bacteria lacking an E. coli PolV homolog was the same (Timinskas and Venclovas, 2019): the catalytically inactive Y family polymerase, ImuB, functions as hub protein, interacting physically with the β clamp via a defined β clamp-binding motif and with DnaE2 and ImuA′ (or ImuA) via unknown mechanisms which might include the ImuB C-terminal region or subregions thereof, including the RecA-NT motif (Timinskas and Venclovas, 2019). However, the absence of any direct biochemical and/or structural evidence to support the proposed protein interactions meant this assumption was speculative. Moreover, whereas E. coli PolV is known to be subject to multiple forms of regulation – including temporal (Robinson et al., 2015), spatial (Robinson et al., 2015), internal (Erdem et al., 2014), and conformational (Jiang et al., 2009; Gruber et al., 2015; Jaszczur et al., 2019) – the expression dynamics and subcellular localizations of the mycobacterial mutasome proteins were mostly unknown. Certainly, genomic organization alone (imuA′–imuB/dnaE2 constitute a ‘split’ mutagenic cassette; Erill et al., 2006) could not predict the stoichiometry of any inferred protein complexes, nor the subcellular location(s) of individual mutasome components and their interacting partners.

By fluorescently tagging the known mutasome proteins, we have observed in real time the consistent formation of co-occurring ImuB–β clamp foci in mycobacterial cell populations exposed to genotoxic stress. Although less pronounced than ImuB, we also detected the frequent, reproducible co-occurrence of DnaE2 with the β clamp under the same conditions. Notably, recruitment of ImuB into foci occurred in mutants lacking functional DnaE2 or ImuA′ but was prevented when the ImuB–β clamp-binding motif was mutated – apparently identifying the primacy of the ImuB–β clamp interaction in mutasome organization. In contrast, the function(s) and subcellular dynamics of ImuA′ remain enigmatic: VFP-ImuA′ consistently produced diffuse fluorescence in DNA-damaged bacilli, precluding any definitive insights into its potential association with ImuB (or DnaE2) in vivo. ImuAʹ and ImuB are encoded in a two-gene operon; therefore, their differential intracellular profiles (ImuB concentrated in foci, ImuA′ diffusely distributed) were not predicted (nor predictable) based on genomic organization but instead required direct observation of the tagged proteins. Here, we note that these results have been reproduced by others in very recent work published during revision of our manuscript (Ng et al., 2023). The distinct intracellular distributions in vivo contrasted, too, with the biochemical analyses, in which the demonstrated co-elution of ImuA′–ImuB and ImuA′–ImuB–β clamp complexes provided important confirmation of the ImuA′–ImuB interaction inferred previously (Warner et al., 2010). Therefore, while difficult to reconcile with the in vitro data, the absence here of a clear co-localization signal in live cells might indicate the transient association of ImuA′ with its mutasome partners or, possibly, that a posttranslational modification is required in live bacteria – by analogy with the proteolytic cleavage of UmuD to UmuD' in the E. coli SOS response (Goodman et al., 2016). Future work will require single-molecule tracking of ImuA′ to resolve this possibility.

The original identification of the imuA–imuB–dnaE2 cassette noted its close association with LexA across diverse bacteria; that is, genomes containing the cassette invariably encoded a LexA homolog, too (Erill et al., 2006). Recent work in mycobacteria has added unexpected nuance to that regulatory framework, namely that the split imuA–imuB/dnaE2 cassette is subject to transcriptional control by both the ‘classic’ LexA/RecA-regulated SOS response and the PafBC-mediated DNA damage response (Adefisayo et al., 2021). The authors of that work also report that, while the two regulatory mechanisms are partially redundant for genotoxic stresses including UV and MMC exposure, fluoroquinolones appear to be specific inducers of PafBC only. In addition to suggesting that chromosomal mutagenesis is co-dependent on PafBC and SOS, these observations are important in identifying an apparent ‘fail-safe’ mechanism in mycobacteria in which the mutasome components are induced irrespective of DNA damage type – again reinforcing the centrality of these proteins in damage tolerance and, by implication, adaptive mutagenesis. Here, it is important to consider also the potential role of the mycobacterial DinB-type DNA polymerases in genome diversification in M. tuberculosis (Dupuy et al., 2022; Dupuy et al., 2023). Although expression of these Y family polymerases is not induced in M. tuberculosis in response to DNA damage (the M. smegmatis SOS response includes a third DinB homolog, DinB3, but this gene is absent from the M. tuberculosis genome), there is evidence suggesting some functional redundancy with DnaE2. Moreover, the differential capacity of M. tuberculosis DinB1 and DinB2 to bind the β clamp suggests the potential for complex protein interplay at stalled replication forks, the exact details of which remain to be elucidated.

We previously observed that the imuBAAAAGG β clamp-binding motif mutation eliminated UV-induced mutagenesis and MMC damage tolerance in M. smegmatis (Warner et al., 2010), phenocopying deletion of any of the three mutasome components (imuA′, imuB, and dnaE2) alone or in combination. Given the abrogation of ImuB focus formation, it seems reasonable to infer a direct link between ImuB–β clamp focus formation and mutasome function. In turn, this suggests that blockade of ImuB focus formation might offer a tractable read-out for a screen designed to identify mutasome inhibitors – a possibility reinforced by the observed co-elution in biochemical assays of β with ImuB and, separately, of the β clamp with pre-formed ImuA′–ImuB complexes. In this context, it was notable in this study that GRS disrupted the ImuB–β clamp interaction in vitro and prevented ImuB focus formation in mycobacteria treated simultaneously with MMC and GRS.

The discrepant complementation phenotypes observed for V-ImuA′ and G-ImuB in the DNA damage tolerance (involving growth on MMC-containing solid media) versus induced mutagenesis (transient MMC or UV exposure during liquid culture) assays suggests that addition of the bulky fluorophore might have prevented full function of these mutasome proteins. Whereas UV irradiation predominantly generates cyclobutane dimers and pyrimidine–pyrimidone (6–4) photoproducts (Franklin et al., 1985), MMC induces a variety of different DNA lesions, including inter- and intra-strand cross-links. These are likely to require multiple repair pathways and, potentially, the interaction of mutasome components with additional protein partners – which might be prevented by the bulky fluorescent tags. The DnaE2–EGFP fusion proved the exception; in this context, it might be instructive to consider recent evidence implicating DnaE2 in gap filling following nucleotide excision repair in non-replicating Caulobacter crescentus cells (Joseph et al., 2021). These observations suggest the importance of identifying other potential interacting partners of mycobacterial DnaE2 (and the other mutasome components), work which is currently underway in our laboratory.

The potential for inhibitors of DNA replication to accelerate the development of genetic resistance through the induction of mutagenic repair/tolerance pathways (Cirz et al., 2005; Barrett et al., 2019; Revitt-Mills and Robinson, 2020) is a valid and commonly cited concern that might partially explain the relative under-exploration of DNA metabolism as source of new antibacterial drug targets (Reiche et al., 2017; van Eijk et al., 2017). Our results suggest that GRS could offer an interesting exception: that is, in binding the β clamp at the site of interaction with the DnaE1 replicative DNA polymerase as well as other DNA metabolizing proteins (Kling et al., 2015), including the clamp loader complex and ImuB, GRS appears to possess an intrinsic protective mechanism against induced mutagenesis – blocking both ImuB-dependent mutasome recruitment to stalled replisomes and post-repair fixation of mutations by the replicative polymerase, DnaE1. This ‘resistance-proofing’ capacity, which is supported by the observed restriction of GRS resistance to low-frequency, high-fitness cost amplifications of the dnaN genomic region with very few to no ‘off-target’ single nucleotide polymorphisms (SNPs), might also contribute to the observed bactericidal effect of GRS against mycobacteria (Kling et al., 2015). In addition, it reinforces the β clamp as a vulnerable target for new TB drug development (Bosch et al., 2021). In this context, it is worth noting that inhibition of DnaE1 replicative polymerase function might represent a general solution to the problem of drug-induced (auto)mutagenesis by preventing fixation of repair/tolerance-generated mutations; in support of this inference, another natural product, nargenicin, which inhibits M. tuberculosis DnaE1 via a DNA-dependent mechanism, fails to yield spontaneous resistance mutations in vitro (Chengalroyen et al., 2021). Therefore, while the essentiality of DNA replication proteins such as DnaN and DnaE1 for mycobacterial viability poses a challenge to the design of assays to detect ‘anti-evolution’ compounds targeting these proteins (because inhibition of their essential, replicative function is growth inhibitory), GRS (and nargenicin) appear to provide compelling evidence that inhibition of some DNA replicative and repair functions might ameliorate the perceived risks in targeting this area of mycobacterial metabolism.

DnaE2-dependent DNA damage tolerance and induced mutagenesis were originally discovered using M. smegmatis as model mycobacterial organism, with key additional observations – including the contribution to pathogenicity and evolution of resistance under drug therapy – made in the pathogen, M. tuberculosis (Boshoff et al., 2003). Continuing that trend, the subsequent elucidation of the roles of ImuA′ and ImuB as essential ‘accessory factors’ confirmed that the fundamentals of mutasome function were equivalent in both species (Warner et al., 2010). Our reliance in the current work on M. smegmatis as proxy is therefore justifiable, but does require caution in extrapolating the refined model for mutasome function to all other mycobacteria encoding mutasome proteins, including M. tuberculosis. That said, it is tempting to consider the implications of the results described here to an obligate pathogen whose persistence within its human host depends on the ability to drive successive cycles of infection, disease – in some cases latency followed by reactivation disease – and transmission (Lin and Flynn, 2018). Such cycles are inevitably vulnerable to multiple potential evolutionary culs-de-sac which might arise in consequence of the elimination of the bacillus by the host (clearance) or the demise of the organism within the infected individual (controlled subclinical infection, or host death). Modern M. tuberculosis strains therefore represent the genotypes that have successfully adapted to human colonization (Gagneux, 2018), evolving with their obligate host through changes in lifestyle and nutritional habits (with their associated implications for non-communicable diseases such as diabetes), the near-universal administration of the BCG vaccination, the emergence of the HIV co-pandemic, and the widespread use of frontline combination chemotherapy (Warner et al., 2015). While the emergence and propagation of drug-resistant isolates characterized by a variety of polymorphisms at multiple genomic loci (Warner et al., 2017; Farhat et al., 2019; Payne et al., 2019) provides strongest proof of the capacity for genetic variation in M. tuberculosis, other lines of evidence include the highly subdivided population structure of the M. tuberculosis complex (Riojas et al., 2018), the well-described geographical host–pathogen sympatry (Hershberg et al., 2008; Brynildsrud et al., 2018) and, more recently, the observation of intra-patient bacillary microdiversity (Ley et al., 2019). In combination, these elements support the ongoing evolution of M. tuberculosis, as well as suggest the potential that ‘anti-evolution’ therapeutics might yield much greater benefit in the clinical context than can be inferred from in vitro studies – in which the pressures on an obligate pathogen can only be approximated. That is, in addition to identifying the mutasome as target for adjunctive therapeutics designed to protect anti-TB drugs against emergent resistance, the results presented here support the further exploration of this and related strategies to disarm host-adaptive mechanisms in a major human pathogen and growing contributor to antimicrobial resistance.

Materials and methods

Bacterial strains and culture conditions

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All mycobacterial strains (Supplementary file 2 – Key Reagents) were grown in liquid culture containing Difco Middlebrook 7H9 Broth (BD Biosciences, San Jose, CA) and supplemented with 0.2% (vol/vol) glycerol (Sigma-Aldrich, St. Louis, MO), 0.005% (vol/vol) Tween 80 (Sigma-Aldrich, St. Louis, MO), and 10% (vol/vol) BBL Middlebrook OADC Enrichment (BD Biosciences, San Jose, CA). For M. smegmatis, liquid cultures were incubated at 37°C with orbital shaking at 100 rpm, until the desired growth density was attained – measured by spectrophotometry at a wavelength of 600 nm – before further experimentation. Solid media comprised Difco Middlebrook 7H10 Agar (BD Biosciences, San Jose, CA) supplemented with 0.5% (vol/vol) glycerol (Sigma-Aldrich, St. Louis, MO), and 10% (vol/vol) BBL Middlebrook OADC Enrichment (BD Biosciences, San Jose, CA). Solid media plates were incubated at 37°C for 3–4 days or until colonies had formed.

Mutasome reporter constructs

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The V-imuA′ construct was designed by altering the coding sequence of imuA′ within the complementing vector, pAINT::imuA′ (Warner et al., 2010), so that the coding sequence of VFP (Nagai et al., 2002) was inserted in-frame after the start codon of the imuA′ ORF. Furthermore, an in-frame FLAG tag sequence (Einhauer and Jungbauer, 2001) was inserted between the coding region of vfp and imuA′ to produce a single ORF encoding VFP-FLAG-ImuA′. For ImuB, the construct PSOS(imuA′)-egfp-imuB was designed such that the regulatory elements immediately upstream of imuA′ were inserted immediately upstream of the imuB ORF which was further altered by inserting the sequence encoding EGFP (Cormack et al., 1996) linked to a FLAG tag-encoded sequence immediately after the start codon of imuB to produce a single ORF encoding EGFP-FLAG-ImuB′ which was cloned into pMCAINT::imuB (Warner et al., 2010). For DnaE2, the egfp sequence was inserted in-frame after the start codon of M. smegmatis dnaE2 (Figure 1—figure supplement 1A).

Mutant binding G-imuBAAAAGG construct

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To introduce the 352AAAAGG357 imuB allele (Warner et al., 2010) into the EGFP-ImuB protein, the nucleotide sequence from pMCAINT::imuBAAAAGG was swapped into the corresponding position of PSOS(imuA′)-egfp-imuB to yield pMCAINT::PSOS(imuA′)-egfp-imuBAAAAGG.

M. smegmatis mutasome reporter strains

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M. smegmatis strain V-ImuA′ was generated by introducing the pAINT::vfp-imuA′ plasmid into ΔimuA′ (Warner et al., 2010) by the standard electroporation method. Strains G-ImuB, and G-ImuBAAAAGG were developed by integration of the pMCAINT::PSOS(imuA′)-egfp-imuB, or pMCAINT::PSOS(imuA′)-egfp-imuBAAAAGG plasmid, respectively, into the genome of ΔimuB (Warner et al., 2010). To generate the G-DnaE2 strain, pTweety::egfp-dnaE2 was electroporated into ΔdnaE2 (Warner et al., 2010). The dnaN-mCherry::G-imuB, dnaN-mCherry::V-imuAʹ, and dnaN-mCherry::G-dnaE2 strains were developed by the electroporation of pMCAINT::PSOS(imuA′)-egfp-imuB, pAINT::vfp-imuA′ and pTweety::egfp-dnaE2 into the M. smegmatis dnaN-mCherry background (Santi et al., 2013). Mutasome-deficient strains ΔimuA′, ΔdnaE2, and dnaE2AIA were electroporated with pMCAINT::PSOS(imuA′)-egfp-imuB to produce ΔimuA′::G-imuB, ΔdnaE2::G-imuB, and dnaE2AIA::G-imuB, respectively.

Antibiotic treatments

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MMC (from Streptomyces caespitosus) (Sigma-Aldrich, St. Louis, MO) was dissolved in ddH2O, while GRS was dissolved in dimethyl sulfoxide (DMSO). Cultures of M. smegmatis were grown in 7H9-OADC – supplemented with selection antibiotic where applicable – at 37°C to an optical density (OD600) ~0.2–0.4. Thereafter, cultures were split into separate 5 ml cultures and MMC and/or GRS added to a final concentration dependent on the MIC (Kling et al., 2015).

DNA damage sensitivity and mutagenesis assays

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UV-induced mutagenesis assays were performed as previously described (Boshoff et al., 2003; Warner et al., 2010), with RIF-resistant colonies enumerated on solid media after 5 days of growth. For UV-induced DNA damage survival assays, M. smegmatis strains were grown in liquid culture to OD600 ~ 0.5, following which a 10-fold dilution series was spotted (5 μl/spot) on standard 7H10 medium, allowed to dry, and then exposed to UV at 12.5 or 25 mJ/cm2; plates were imaged after 3 days’ incubation at 37°C. MMC-induced mutagenesis assays were performed by treating log-phase bacteria with 0.5× MIC MMC for 6 hr, following which the bacteria were washed and RIF-resistant colonies were enumerated on solid media as before. Mutation frequencies were calculated by dividing the number of RIF-resistant colonies of each sample by the CFU/ml of untreated sample. For MMC damage sensitivity assays, the cultures were grown to OD600 ~ 0.4, following which a 10-fold dilution series was spotted on standard 7H10 medium and 7H10 medium supplemented with MMC; plates were incubated for 3 days and imaged.

Snapshot microscopy

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Single snapshot micrographs of M. smegmatis cells were captured with a Zeiss Axioskop M, Zeiss Axio.Scope, and Zeiss Axio.Observer Z1. Briefly, 2.0–5.0 μl of liquid culture was placed between a No. 1.5 glass coverslip and microscope slide. A transmitted mercury lamp light was used together with filter cubes to visualize fluorescence using a ×100 1.4 NA plan apochromatic oil immersion objective lens. Samples were located using either transmitted light, differential interference contrast, or epifluorescence. Snapshot images were captured with either a Zeiss 1 MP or Zeiss AxioCam HRm monochrome camera. Images of the same experiment were captured with the same instrument and exposure settings. Green fluorescence of EGFP was detected using the Zeiss Filter Set 38 HE. Red fluorescence of mCherry was detected using the Zeiss Filter Set 43. Images were captured using AxioVision 4.7 or ZEN Blue Microscope and Imaging Software. Images were processed using Fiji (Schindelin et al., 2012); images of the same strain were contrasted to the same maximum and minimum within an experiment.

Quantitative image analysis

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M. smegmatis bacilli were plotted from shortest to longest and aligned according to their midcell position (0 on the y-axis) using the MicrobeJ plugin of ImageJ (Ducret et al., 2016). Along each point of the cell, a dot was generated and colored according to the fluorescence intensity along the medial axis of the bacillus. Therefore, this plot represents the fluorescence intensity along the medial axis of every bacillus imaged under the relevant experimental conditions. R was used for visual representation of the data.

Single-cell time-lapse fluorescence microscopy

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Liquid cultures of M. smegmatis reporter strains were grown to mid-logarithmic phase (OD600 = 0.6), cells were collected by centrifugation at 3900 × g for 5 min and concentrated 10-fold in 7H9 medium. The cells were filtered through a polyvinylidene difluoride syringe filter (Millipore) with a 5-µm pore size to yield a clump-free cell suspension. The single-cell suspension was spread on a semi-permeable membrane and secured between a glass coverslip and the serpentine 2 chip (Delincé et al., 2016) in a custom-made PMMA/Aluminium holder (Dhar and Manina, 2015). Time-lapse microscopy employing a DeltaVision personalDV inverted fluorescence microscope (Applied Precision, WA) with a ×100 oil immersion objective was used to image single cells of M. smegmatis. The bacteria and microfluidic chip were maintained at 37°C in an environmental chamber with a continuous flow of 7H9 medium, with or without 100 ng/ml of MMC, at a constant flow rate of 25 µl/min, as described previously (Wakamoto et al., 2013; Dhar and Manina, 2015). Images were obtained every 10 min on phase-contrast and fluorescence channels (for EGFP, excitation filter 470/40 nm, emission filter 525/50 nm; for mCherry, excitation filter 572/35, emission filter 632/60; for YFP excitation filter 500/20 nm, emission filter 535/30 nm) using a CoolSnap HQ2 camera. Image-based autofocus was performed on each point prior to image acquisition. Experiments were repeated two to four times; a typical experiment collected images from up to 80 XY points at the 10-min intervals. The images were analyzed using Fiji (Schindelin et al., 2012).

Protein expression and purification

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N-terminally His-tagged M. smegmatis ImuB was co-expressed with ImuA′ in E. coli BL21(DE3) cells using two expression vectors from the NKI-LIC vector suite (Luna-Vargas et al., 2011): pETNKI-his-3C-LIC-kan for ImuB and pCDFNKI-StrepII3C-LIC-strep for ImuA′ that have different resistance markers, kanamycin and streptomycin; as well as different origins of replication, ColE1 and CloDF13, respectively. Protein production was induced with isopropyl 1-thio-β-d-galactopyranoside at 30°C for 2 hr. The ImuBA′ complex was purified using a Histrap column followed by a Superdex 200 16/60 column. Both N-His6 M. smegmatis ImuB and β clamp were expressed in E. coli BL21(DE3) cells and purified using HisTrap, HiTrap Q, and S200 columns. All proteins were flash frozen in liquid nitrogen and stored at −80°C.

Size-exclusion chromatography analysis

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Samples of individual proteins and the different complexes were injected onto a PC3.2/30 (2.4 ml) Superdex 200 Increase gel filtration column (GE Healthcare) pre-equilibrated in 50 mM Tris pH 8.5 and 300 mM NaCl. Thereafter, 50 μl fractions were collected and analyzed by SDS–PAGE electrophoresis using 4–12% NuPage Bis‐Tris precast gels (Life Technologies). Gels were stained with 0.01% (vol/vol) 2,2,2-trichloroethanol and imaged with UV light.

Thermal unfolding experiments

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Melting curves of the M. smegmatis β clamp (5 μM) in the presence and absence of GRS (15 μM) were measured in UV capillaries using the Tycho NT6 (NanoTemper Technologies) where the protein unfolding is followed by detecting the fluorescence of intrinsic tryptophan and tyrosine residues at both emission wavelengths of 350 and 330 nm.

Structure modeling and analysis

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Structural models for M. segmatis β clamp interaction with ImuB and DnaE1 were generated using ColabFold implementation (Mirdita et al., 2022) of AlphaFold-Multimer v.2 (Jumper et al., 2021; https://doi.org/10.1101/2021.10.04.463034). Structures of M. smegmatis and M. tuberculosis clamp complexes with GRS were obtained from PDB (PDB ids 5AH2 and 5AGU, respectively). Residues at the interaction interfaces were identified using VoroContacts server (Olechnovič and Venclovas, 2021). Structures were visualized using UCSF ChimeraX (Goddard et al., 2018).

Data availability

Source data for all figures contained in the manuscript and SI have been deposited in Dryad (http://doi.org/10.5061/dryad.76hdr7szc).

The following data sets were generated
    1. Warner DF
    (2023) Dryad Digital Repository
    Data from: The mycobacterial ImuA'-ImuB-DnaE2 mutasome: composition and recruitment in live cells.
    https://doi.org/10.5061/dryad.76hdr7szc

References

Decision letter

  1. Christina L Stallings
    Reviewing Editor; Washington University School of Medicine, United States
  2. Dominique Soldati-Favre
    Senior Editor; University of Geneva, Switzerland

Our editorial process produces two outputs: i) public reviews designed to be posted alongside the preprint for the benefit of readers; ii) feedback on the manuscript for the authors, including requests for revisions, shown below. We also include an acceptance summary that explains what the editors found interesting or important about the work.

Decision letter after peer review:

Thank you for submitting your article "The mycobacterial mutasome: composition and recruitment in live cells" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by Gisela Storz as the Senior Editor. The reviewers have opted to remain anonymous.

The reviewers have discussed their reviews with one another, and the Reviewing Editor has drafted this to help you prepare a revised submission.

Essential revisions:

In this manuscript, the authors test their previously proposed model (also presented in Figure 1A) that ImuB interacts with the DnaN DNA polymerase III β clamp to recruit DnaE2. The previously identified mutasome components ImuA', ImuB, and DnaE2 and essential for DNA-damage induced mutagenesis. Although the exact function of ImuA' and ImuB is unknown, ImuB has long before been proposed to interact with DnaN via an interaction domain within ImuB that has already been identified. Since the experiments herein test and validate a well-establish model, the results are somewhat expected. However, all models should be tested experimentally, making this an important confirmation. The manuscript nicely makes use of both in vivo and in vitro approaches and the data is convincing for the most part, although the inability of the fusion proteins used in this study to complement the knockout strains during exposure to the DNA damaging agent MMC does raise an important limitation of the tools used herein and brings into question whether MMC should have been the genotoxic agent used in the studies. In addition, a major concern is the limited new biological insight gained from the study in its current form and a revision should address these limitations. The reviewers make several suggestions to help improve the scope and impact of the manuscript, as detailed here.

1) The section starting at Line 89 describes experiments expressing the fluorescently tagged proteins in M. smegmatis, but does not mention the mCherry-DnaN strand background even though the data shown in Figure 1 is in that background. Then a couple sections later starting on Line 174 the authors mention using the mCherry-DnaN background in a "new" set of experiments, but refer to the same data in Figure 1. This current organization is deceptive in that it makes it sound like these experiments were done twice, once in a background with mCherry-DnaN and once without. Were the experiments done in the absence of mCherry-DnaN? Is this what is in Figure 1 —figure supplement 1? If so, it would be better to include this in Figure 1 and would be beneficial to show that the localization of these proteins is not affected by the mCherry-DnaN tag. As written these sections basically just repeat each other almost word for word in terms of observations and should be combined up front. Regardless, imaging data without mCherry DnaN would be valuable.

2) The studies examining the effect of the fluorescently tagged proteins on mutagenesis and tolerance to DNA damage should be included in the main figure, particularly given the importance of the tagged constructs not complementing the deletions for survival following DNA damage. This is an important finding and highlights that localization of ImuB to foci is not sufficient to promote viability and function. It also looks like the DnaE2 construct did complement during MMC treatment, but the others did not. The authors should also include experiments with untagged versions to determine if it is the tag that is precluding complementation, rather than an expression issue or something with the vector.

3) Since the deletion if ImuA' affects MMC survival, could the fact that its localization remains disperse explain why these strains are not complementing? Since the DnaE2 construct does appear to complement, it could be interesting to test ImuA' localization in this strain.

4) Figure 1B is not referenced in the text. What time point is this? This information should also be included in the Figure Legend.

5) Does the δ imuA' affect DnaE2 localization?

6) The way the manuscript is written, it jumps around a lot, going to figure 4, then back to figure 3. Are there replicate experiments to present with appropriate controls so that this does not need to happen?

7) The ImuA' and ImuB translational fusions are not active in MMC-induced mutagenesis. Given that these translational fusions failed to support function, the contribution of the observed foci following MMC exposure to ImuABC function is unclear. Do these foci represent functional complexes, dead intermediates due to the fluorescent tags, or are they artefacts of inactive proteins? Given that MMC is the primary damage used by the authors, it is necessary to establish that the tags do not perturb lesion bypass (and mutagenesis) under this damage. The presented evidence would suggest that the tags do perturb mutasome function in MMC and that this treatment may not be reliable for extracting information on dynamics. In addition, the translational ImuA' and ImuB fusions supported UV-induced mutagenesis. Thus, UV seems to be a superior DNA damaging agent for the colocalization studies. Results using UV should be included to provide a more accurate picture of Imu-clamp colocalization.

8) 3D microscopy is not well explained for the non-expert. How was colocalization established along the z-axis? A better description for non-experts is needed. In addition, more quantification of the images is necessary. For example, It is also unclear what percentage cells have ImuB and DnaE2 foci, if these percentages vary in a dose-dependent manner, and if they reduce during recovery. In addition, how many cells (%) show co-localization of the different Imu proteins with β clamp? Do they always colocalize, and how often is an Imu protein or a clamp protein focus seen alone? If most foci show colocalization, do the authors conclude that all clamps on DNA are in complex with ImuB and would this be surprising? β-clamp foci denote all loaded replisomes, including the ones that are actively synthesizing DNA with the replicative polymerase. Thus, do the number of localizations observed for clamp vary with and without damage?

Might foci be comprised of multiple proteins (in addition to the different Imus and the clamp), some of which may be active on the DNA and others of which may be in close proximity and ready to be called into action? ImuB also interacts with Pol III (Warner et al., 2010, PNAS). Is Pol III also present in the complexes? Im Figure 1 supp 1B, do the number of localizations observed vary between UV and MMC treatment?

9) Does addition of GRS impact ImuA'BC-dependent mutagenesis? This would be the direct way to test the proposal that therapeutic disruption of ImuB-clamp interactions would inhibit mutagenesis in M. tuberculosis. Mutagenesis assays should be performed in the GRS-MMC treated conditions to support any conclusion on GRS abrogation of mutasome action.

10) The authors were unable to purify ImuC, so its interactions with ImuA' and ImuB, and the effects of these interactions on its polymerase activity are unknown. While this is unfortunate, it is not the fault of the authors. However, in the absence of ImuC, it seems there is more that could be done with ImuA' and ImuB to test and extend further the published model for ImuABC function (Warner et al., 2010, PNAS). For example, does ImuB interact with itself via its C-terminal domain as it did in yeast-two-hybrid? Does ImuA' interact with the β clamp, or influence the affinity of ImuB for the clamp? Does ImuB lack an intrinsic DNA polymerase activity as predicted by its lack of conserved acid active site residues (Warner et al., 2010, PNAS)? Related to this, are both ImuA' and ImuC dispensable for ImuB-clamp colocalization in live cells?

11) The authors seem to make two contradicting arguments – on the one hand, they argue that ImuA'BC acts on multiple DNA damaging agents, but on the other, they argue that different DNA damaging agents may require different accessory proteins for proper Imu function. The authors could reconcile these arguments early in the text.

12) What was the spontaneous M. tuberculosis mutation frequency, and how does it compare to the frequencies of UV-induced mutagenesis for the ∆imuA', ∆imuB, and ∆imuC strains?

13) In general, the temporal dynamics of mutasome association with clamp are a promising part of the manuscript, but the authors do not explore this thoroughly. Quantitative analysis, dose dependency and temporal dynamics during damage and recovery (as interpreted by the authors throughout the manuscript) are lacking characterization.

14) In Line 206- The authors state that their observations suggest that deficiency in ImuA' would affect ImuB localization, but it is not clear why they believe this to be true. ImuB has a clamp binding motif, so it is likely to associate with the clamp irrespective of ImuA'. In order to support ImuA'-related conclusion, the authors would need to quantify the number of localizations observed for ImuB and β clamp in the presence and absence of ImuA'.

15) GRS impact on clamp/ImuB: In Line 295 and Figure 3B. The authors should show the clamp+GRS alone profile as well.

16) Figure 4 and conclusions with regards to impact of GRS as well as ImuB clamp binding mutant. It is possible GRS alone affects clamp stability, irrespective of mutasome function. The authors need to image the clamp after GRS treatment, to assess whether the lack of ImuB localization is because it cannot bind the clamp or because the clamp itself is no longer localized.

17) What is the impact of the clamp-binding-ImuB mutant on clamp localization? Is it similar to GRS treatment?

Other comments:

1. L139 – L143: To make this conclusion, authors need to carry out experiments across a range of doses.

2. Figure 2 – individual fluorescence panels need to be shown independently. Currently it is hard to visualize the localizations with accuracy.

3. L276 in Mtb, the PafBC regulatory system also influences Imu expression. The authors need to rephrase.

4. Please use β-clamp everywhere (and not only B).

5. Figure 3A. The difference in elution profiles between ImuA'-ImuB-clamp an ImuB-clamp would suggest that ImuB-clamp interaction alone might be less stable (in absence of ImuA'). Could the authors comment on the same?

6. Figure 4 legend. Top and bottom panel references need to be updated.

7. L314-315: Details of population analysis are missing in the legends or text.

8. L341. "when" instead of "where".

9. L338. "repair/ tolerance" pathways.

10. L379. Could the authors clarify? Do they envision DnaE2 acting without clamp? In that case, what could the potential mechanism be?

11. L405. It is unclear whether the GRS phenotype is due to its action on the replisome, independent of damage / mutasome effects, unless the impact of GRS on clamp alone is tested.

12. It is understandable that the authors use M. smeg as a model system to derive conclusions on action of the mutasome. However, the paragraph starting L410 needs to be toned down as all experiments are performed in smeg and not M.tb. Any extrapolation of their conclusions need to be explicitly stated to the reader.

https://doi.org/10.7554/eLife.75628.sa1

Author response

Essential revisions:

In this manuscript, the authors test their previously proposed model (also presented in Figure 1A) that ImuB interacts with the DnaN DNA polymerase III β clamp to recruit DnaE2. The previously identified mutasome components ImuA', ImuB, and DnaE2 and essential for DNA-damage induced mutagenesis. Although the exact function of ImuA' and ImuB is unknown, ImuB has long before been proposed to interact with DnaN via an interaction domain within ImuB that has already been identified. Since the experiments herein test and validate a well-establish model, the results are somewhat expected. However, all models should be tested experimentally, making this an important confirmation. The manuscript nicely makes use of both in vivo and in vitro approaches and the data is convincing for the most part, although the inability of the fusion proteins used in this study to complement the knockout strains during exposure to the DNA damaging agent MMC does raise an important limitation of the tools used herein and brings into question whether MMC should have been the genotoxic agent used in the studies.

The reviewers justifiably questioned the appropriateness of mitomycin C (MMC) as genotoxic agent given the observed inability of the fluorescent reporter alleles to complement the respective mutasome deletion mutants under extended MMC exposure in the DNA damage survival assay. To address this concern, we have generated new data showing that V-ImuAʹ and G-ImuB fully complement the corresponding deletion mutants in the MMC damage-induced mutagenesis assay (Figure 2). We also show that UV treatment recapitulates the mutagenesis results observed with MMC; that is, the fluorescent reporter alleles complement loss of the wildtype mutasome proteins under UV-induced mutagenesis (Figure 1 —figure supplement 1). In combination, these data appear sufficient to support the notion that the fluorescent reporters are functional in DNA damageinduced mutagenesis.

Our original observation that V-ImuAʹ and G-ImuB fail to complement the corresponding deletion mutants in the MMC DNA damage survival assay holds. Although the precise reason for this difference remains elusive, we think it might be instructive in revealing that the addition of fluorescent tags to these mutasome proteins has possibly disrupted an interaction(s) with another protein(s) that, under prolonged exposure to MMC a clastogen which causes multiple types of DNA lesion, is critical for DNA damage survival. Work is ongoing to resolve this conundrum.

1) The section starting at Line 89 describes experiments expressing the fluorescently tagged proteins in M. smegmatis, but does not mention the mCherry-DnaN strand background even though the data shown in Figure 1 is in that background. Then a couple sections later starting on Line 174 the authors mention using the mCherry-DnaN background in a "new" set of experiments, but refer to the same data in Figure 1. This current organization is deceptive in that it makes it sound like these experiments were done twice, once in a background with mCherry-DnaN and once without. Were the experiments done in the absence of mCherry-DnaN? Is this what is in Figure 1 —figure supplement 1? If so, it would be better to include this in Figure 1 and would be beneficial to show that the localization of these proteins is not affected by the mCherry-DnaN tag. As written these sections basically just repeat each other almost word for word in terms of observations and should be combined up front. Regardless, imaging data without mCherry DnaN would be valuable.

We apologize for the confusing presentation of Results in the original version. The fluorescent reporters were introduced and visualized in both the respective knock-out backgrounds as well as in the mCherry-DnaN background. As requested, Figure 1 has been revised to contain images obtained from the fluorescent reporters in the knock-out backgrounds alone; this is clarified in the revised figure legend. All subsequent experiments were performed in the mCherry-DnaN background; this is made explicit from the section entitled "ImuB localizes with the dnaN-encoded β clamp following DNA damage" and thereafter.

2) The studies examining the effect of the fluorescently tagged proteins on mutagenesis and tolerance to DNA damage should be included in the main figure, particularly given the importance of the tagged constructs not complementing the deletions for survival following DNA damage. This is an important finding and highlights that localization of ImuB to foci is not sufficient to promote viability and function.

We agree with this suggestion and, as noted in response to reviewer comment 1, we have revised the manuscript to incorporate new data, as well as restructuring the Results section to foreground these observations. The results of the complementation assays are now included in a new Figure 2, which contains original data from the UV-induced mutagenesis (Figure 2A) and MMC damage sensitivity (Figure 2C) assays, and new data from a MMC damage-induced mutagenesis assay (Figure 2B).

It also looks like the DnaE2 construct did complement during MMC treatment, but the others did not. The authors should also include experiments with untagged versions to determine if it is the tag that is precluding complementation, rather than an expression issue or something with the vector.

In this work, we utilized the same complementation system as reported previously in Warner et al. (2010; doi:10.1073/pnas.1002614107), which restored full function using untagged ImuAʹ, ImuB, and DnaE2 alleles (see Figure 1B in that paper); all we did in the current study was append the fluorescent tags to the respective mutasome genes. It seems very unlikely, therefore, that issues with the expression system might account for the lack of complementation in the MMC damage sensitivity assays. Consistent with related comments (see response to introductory reviewers’ comment), our interpretation instead is that the presence of the fluorescent tags might have impacted full protein function in cells under prolonged exposure to a lethal genotoxic stress. This conclusion is supported by the functionality of the tagged alleles in both UV- and MMCinduced mutagenesis assays (Figure 2).

3) Since the deletion if ImuA' affects MMC survival, could the fact that its localization remains disperse explain why these strains are not complementing? Since the DnaE2 construct does appear to complement, it could be interesting to test ImuA' localization in this strain.

This is an excellent question. We cannot exclude the possibility that it is the presence of the VFP tag that prevents ImuA’ association with ImuB in vivo in live mycobacterial cells, resulting in the diffuse fluorescence signal. However, the biochemical data presented in Figure 4 —figure supplement 1B indicate that the addition of fluorescent tags to either ImuB or ImuA’ do not disrupt the interaction of these proteins in vitro. Moreover, (see response to introductory reviewers’ comment) the V-ImuA’ allele does complement DNA damage-induced mutagenesis following exposure to either MMC or UV, suggesting functionality of the tagged protein. Work is underway in our laboratory to elucidate the components of the mutasome in live cells without requiring fluorescent labels.

4) Figure 1B is not referenced in the text. What time point is this? This information should also be included in the Figure Legend.

Figure 1B has been removed in preparing the revised manuscript.

5) Does the δ imuA' affect DnaE2 localization?

This is another very good question. As shown in Figure 1, unlike ImuB, which produces distinct foci, the DnaE2 signal is less definitive. Therefore, addressing the potential roles of the other mutasome proteins in DnaE2 localization requires alternative, more sensitive approaches such as single-molecule tracking. This work is ongoing and, as noted below, includes attempts to express purified DnaE2 for analysis in biochemical assays alone and in combination with the other mutasome components.

6) The way the manuscript is written, it jumps around a lot, going to figure 4, then back to figure 3. Are there replicate experiments to present with appropriate controls so that this does not need to happen?

We apologize for the confusion caused by the previous presentation of the Results. As noted in response to reviewers’ comment 1, the revised version has been restructured in accordance with the reviewers’ recommendations to make it clearer and more logical. All results presented in the manuscript reflect multiple biological replicates utilizing appropriate controls.

7) The ImuA' and ImuB translational fusions are not active in MMC-induced mutagenesis. Given that these translational fusions failed to support function, the contribution of the observed foci following MMC exposure to ImuABC function is unclear. Do these foci represent functional complexes, dead intermediates due to the fluorescent tags, or are they artefacts of inactive proteins? Given that MMC is the primary damage used by the authors, it is necessary to establish that the tags do not perturb lesion bypass (and mutagenesis) under this damage. The presented evidence would suggest that the tags do perturb mutasome function in MMC and that this treatment may not be reliable for extracting information on dynamics. In addition, the translational ImuA' and ImuB fusions supported UV-induced mutagenesis. Thus, UV seems to be a superior DNA damaging agent for the colocalization studies. Results using UV should be included to provide a more accurate picture of Imu-clamp colocalization.

This comment reflects a common concern among all reviewers about the validity of making conclusions based on the MMC phenotypes alone. The revised version submitted here contains new data showing that V-ImuAʹ and G-ImuB fully complement the corresponding deletion mutants in the MMC damage-induced mutagenesis assay (Figure 2B) [Note that the first sentence in the reviewers’ comment, namely “The ImuA' and ImuB translational fusions are not active in MMC-induced mutagenesis”, is therefore not correct; the translational fusions are active in the MMC-induced mutagenesis assay, but not the MMC DNA damage survival assay.]

In addition, we present evidence that UV treatment recapitulates the results observed with MMC; that is, the fluorescent reporter alleles complement loss of the respective wildtype mutasome proteins in UV-induced mutagenesis (Figure 2A), confirming that this effect is not limited to a single DNA damaging agent. We also show that ImuB foci are induced following exposure of cells to both UV and MMC (for MMC, timelapse data confirm that this is dynamic and occurs rapidly following treatment; Figure 3 and Video 1). In previous work (Warner et al., 2010; doi:10.1073/pnas.1002614107), we reported that mutations in the ImuB β clamp-binding motif (QLPLWG) inhibited the ImuB-β clamp interaction in yeast two-hybrid assays (Figure 4 in Warner et al., 2010; doi:10.1073/pnas.1002614107), and abrogated UV-induced mutagenesis in live cells (Figure 6 in Warner et al., 2010; doi:10.1073/pnas.1002614107). Now, in this revised submission, we demonstrate that the imuBAAAAGG allele disrupts ImuB-β clamp focus formation (Figure 5A), linking focus formation to induced mutagenesis. In combination, these observations suggest that the formation of ImuB foci is a common feature of the mycobacterial DNA damage response and is essential for mutasome function.

As indicated in earlier comments, the reason(s) for the discrepant results between the induced mutagenesis assays and survival assays remains unclear – and might reflect disruption of an interaction with an additional partner(s) required for surviving constant MMC exposure, or could be a consequence of the tag impairing optimal protein function under sustained, lethal genotoxic pressure during prolonged incubation on solid media. That said, the ability of the fusion proteins to support induced mutagenesis following MMC treatment suggests that the MMC-induced foci, like the UV-induced foci, are mutagenic.

As recommended by the reviewers, we have included UV treatment in the microscopy analyses; these results have added value to this work and are presented in Figure 3D. The nature of UV experiments unfortunately complicates the use of timelapse microscopy to monitor responses in UV-treated cells; for this reason, timelapse videos are only available for MMC-treated mycobacteria. Nevertheless, single timepoint fluorescent microscopy images are presented which indicate that UV exposure elicits an equivalent response to that observed following MMC treatment (Figure 1A).

8) 3D microscopy is not well explained for the non-expert. How was colocalization established along the z-axis? A better description for non-experts is needed. In addition, more quantification of the images is necessary. For example, It is also unclear what percentage cells have ImuB and DnaE2 foci, if these percentages vary in a dose-dependent manner, and if they reduce during recovery. In addition, how many cells (%) show co-localization of the different Imu proteins with β clamp? Do they always colocalize, and how often is an Imu protein or a clamp protein focus seen alone? If most foci show colocalization, do the authors conclude that all clamps on DNA are in complex with ImuB and would this be surprising? β-clamp foci denote all loaded replisomes, including the ones that are actively synthesizing DNA with the replicative polymerase. Thus, do the number of localizations observed for clamp vary with and without damage?

We thank the reviewer for this comment which prompted an overhaul of our presentation and analyses of the microscopy data. The descriptions of the microscopy and the associated analyses have been modified to make them more accessible to the non-expert. In addition, to address the valid request for quantitative analyses, several new figures have been included:

i) Figure 3D shows the position of mCherry-DnaN foci in relation to G-ImuB foci in cells expressing both fluorescent proteins; these results indicate strong ImuB-β clamp correlation in cells exposed to either UV or MMC, and near absent expression of ImuB in undamaged cells.

ii) Figure 3 —figure supplement 1 includes new data showing the proportion of cells with mCherry-DnaN and G-ImuB foci. As noted earlier, the less distinct signal obtained from GDnaE2 precluded similar analyses of potential DnaE2 correlations.

iii) Figure 5C and Figure 5D present data on the effect of griselimycin treatment on b clamp and ImuB focus formation in cells exposed to either MMC or UV, as well as in the absence of DNA damage.

Might foci be comprised of multiple proteins (in addition to the different Imus and the clamp), some of which may be active on the DNA and others of which may be in close proximity and ready to be called into action? ImuB also interacts with Pol III (Warner et al., 2010, PNAS). Is Pol III also present in the complexes? Im Figure 1 supp 1B, do the number of localizations observed vary between UV and MMC treatment?

The reviewers raise several very interesting questions here. The potential that other proteins, including the dnaE1-encoded DNA Polymerase IIIa subunit, might be present in foci has been subject of much discussion among the authors – especially given the importance of understanding how the switch occurs from the replicative polymerase to the translesion synthesis machinery. Studies to address these questions are ongoing in our laboratories, and no solid data are available yet to support informed speculation.

To address the query about whether the number of foci differs as a function of genotoxic stress (MMC or UV), we have included an additional analysis (Figure 3—figure supplement 1) showing the proportion of cells with foci as well as the proportion of cells that present only a single signal (that is, either mCherry-DnaN or G-ImuB) or both in response to the different DNA damaging treatments.

9) Does addition of GRS impact ImuA'BC-dependent mutagenesis? This would be the direct way to test the proposal that therapeutic disruption of ImuB-clamp interactions would inhibit mutagenesis in M. tuberculosis. Mutagenesis assays should be performed in the GRS-MMC treated conditions to support any conclusion on GRS abrogation of mutasome action.

The reviewers raise a valid point about the need for an experiment demonstrating that disruption of the ImuB-b clamp interaction inhibits mutagenesis. As noted in response to reviewers’ comment 7, this has been demonstrated genetically: the imuBAAAAGG allele which disrupts ImuB-β clamp focus formation (Figure 5A) was shown previously to abrogate UV-induced mutagenesis in live cells (Figure 6 in Warner et al., 2010; doi:10.1073/pnas.1002614107). The question, though, is whether this can be demonstrated pharmacologically – as the reviewers’ comment implies. Again, this is something we have discussed at length but have not been able to identify a suitable approach: although the reviewers suggest performing mutagenesis assays in the presence of griselimycin, this is not definitive since griselimycin targets both DNA replication – by preventing the DnaE1-b clamp interaction (Kling et al., 2015; doi:10.1126/science.aaa4690) – and mutagenic DNA repair, by preventing ImuB-b clamp focus formation (this manuscript), as well as preventing b clamp localization through the likely inhibition of the clamp loader complex. Griselimycin-mediated replisome collapse has been described previously (Trojanowski et al., 2019; Antimicrob Agents Chemother. doi:10.1128/AAC.00739-19).

Given that subsequent rounds of mycobacterial replication are required for fixation of induced mutations in the genome, it is not possible to separate the effects of replication inhibition from blocked mutagenesis – a conundrum which highlights the need for an on-target inhibitor of the mutasome (a DnaE2 inhibitor, for example). In the absence of a suitable test system, we present microbiological evidence (Figure 5) indicating that treatment of cells with griselimycin eliminates ImuB-β clamp focus formation under mutagenic DNA damage (MMC exposure) conditions, and back this up with biochemical data (Figure 4) showing disruption of ImuB-DnaN and ImuA’-ImuBDnaN interactions in assays utilizing the purified proteins. Together, these observations support the inference that inhibiting mutasome recruitment and/or function will substantively eliminate DNA damage-induced mutagenesis in mycobacteria.

10) The authors were unable to purify ImuC, so its interactions with ImuA' and ImuB, and the effects of these interactions on its polymerase activity are unknown. While this is unfortunate, it is not the fault of the authors. However, in the absence of ImuC, it seems there is more that could be done with ImuA' and ImuB to test and extend further the published model for ImuABC function (Warner et al., 2010, PNAS). For example, does ImuB interact with itself via its C-terminal domain as it did in yeast-two-hybrid? Does ImuA' interact with the β clamp, or influence the affinity of ImuB for the clamp? Does ImuB lack an intrinsic DNA polymerase activity as predicted by its lack of conserved acid active site residues (Warner et al., 2010, PNAS)? Related to this, are both ImuA' and ImuC dispensable for ImuB-clamp colocalization in live cells?

The reviewers have raised multiple very interesting questions and identified numerous potential lines of experimental inquiry. We agree that many of these are pressing and would enhance our understanding of the mutasome; in fact, we are actively pursuing these and many other questions currently. However, we respectfully submit that all are beyond the scope of the current study. The question about the potential roles of ImuA’ and DnaE2 (ImuC) in ImuB-b clamp localization is partially addressed in this manuscript: Figure 3 —figure supplement 2 shows that ImuB focus formation occurs despite the absence of ImuA’ or DnaE2, while Figure 4Aii presents biochemical data to support the ability of ImuB to interact directly with the b clamp in the absence of ImuA’ or DnaE2. Although these results do not directly interrogate the impact of either protein on ImuB-b clamp localization, the strong phenotype of ImuB-b clamp focus formation in DNA damaged cells suggests that the interaction is likely independent of other proteins.

11) The authors seem to make two contradicting arguments – on the one hand, they argue that ImuA'BC acts on multiple DNA damaging agents, but on the other, they argue that different DNA damaging agents may require different accessory proteins for proper Imu function. The authors could reconcile these arguments early in the text.

We apologize for this apparent contradiction which arose from clumsy sentence construction. To clarify, we have modified the text to read (L160-4):

"Although mutasome components are expressed in response to genotoxic stress arising from a variety of different sources, it is possible the different types and/or extent of DNA damage induced in the two separate assays used here (induced mutagenesis vs. DNA damage tolerance) might require distinct interactions with a different partner protein(s) and, further, that one/more of these might have been disrupted by the presence of the fluorescent tag(s)."

12) What was the spontaneous M. tuberculosis mutation frequency, and how does it compare to the frequencies of UV-induced mutagenesis for the ∆imuA', ∆imuB, and ∆imuC strains?

The M. tuberculosis mutation rate has previously been reported as 2-3 x 10-10 mutations per cell per generation (Boshoff et al., 2003; doi:10.1016/s0092-8674(03)00270-8). The same study reported that the spontaneous mutation rate was unaffected in a dnaE2 deletion mutant; however, UV-induced mutagenesis was effectively eliminated in the dnaE2 knockout. In follow-up work (Warner et al., 2010; doi:10.1073/pnas.1002614107), we demonstrated that deletion of any of the mutasome components (ImuA’, ImuB, or DnaE2), alone or in combination, eliminated UV-induced mutagenesis in both M. smegmatis and M. tuberculosis: that is, the UV-induced mutation frequencies in the mutasome mutants were indistinguishable from non-DNA-damaged strains.

13) In general, the temporal dynamics of mutasome association with clamp are a promising part of the manuscript, but the authors do not explore this thoroughly. Quantitative analysis, dose dependency and temporal dynamics during damage and recovery (as interpreted by the authors throughout the manuscript) are lacking characterization.

The reviewers have again identified a very interesting area of study which demands considerable additional experimental work. The heterogeneity evident in DNA damaged cells (see the different timelapse movies, for example) hints at the complexity inherent in tracking the recovery of individual cells. To tackle this problem, we are currently exploring an alternative microfluidic system that might enable the spatial constriction of mycobacterial cells necessary for extended single-cell tracking and analysis of sub-cellular protein localizations.

14) In Line 206- The authors state that their observations suggest that deficiency in ImuA' would affect ImuB localization, but it is not clear why they believe this to be true. ImuB has a clamp binding motif, so it is likely to associate with the clamp irrespective of ImuA'. In order to support ImuA'-related conclusion, the authors would need to quantify the number of localizations observed for ImuB and β clamp in the presence and absence of ImuA'.

The reviewers are correct in suggesting that the capacity of ImuB to bind the β clamp directly might be expected to obviate any impact of ImuA’ on ImuB localization. That said, the structural homology of ImuA' to RecA, plus the demonstrated essentiality of ImuA’ for mutasome function, meant we couldn’t dismiss a priori the possibility that ImuAʹ might affect ImuB localization, even indirectly. Two observations motivated this thinking: (i) knowledge that, in the E. coli system, RecA activates UmuDC in forming the functional DNA Polymerase V mutasome, and (ii) recent work indicating that ImuA inhibits the recombinase activity of RecA1 in Myxococcus xanthus, facilitating mutagenesis (Sheng et al., 2021; Appl Environ Microbiol. doi:10.1128/AEM.00919-21).

Although we did not expect mycobacterial ImuA’ to fulfil analogous roles, we needed to exclude either possibility. If it were the case that ImuA’ (or DnaE2) influenced ImuB localization, we would expect to observe no or limited ImuB focus formation in the imuAʹ (or dnaE2) deletion mutant, which was not the case (Figure 3 —figure supplement 2). This result was also critical in establishing the centrality of the ImuB-β clamp interaction in mutasome function, in turn suggesting a potential high-throughput assay for novel chemical inhibitors of this essential protein-protein interaction.

15) GRS impact on clamp/ImuB: In Line 295 and Figure 3B. The authors should show the clamp+GRS alone profile as well.

Kling et al. (2015; doi:10.1126/science.aaa4690) demonstrated the binding of griselimycin to the b clamp in biochemical and structural assays; this observation is reiterated in the new modelling data (Figure 5B, Figure 5 —figure supplement 1) which indicate clearly that the region of griselimycin binding on the mycobacterial β clamp subunit overlaps with the region predicted to interact with other β clamp-binding proteins, including DnaE1 and ImuB.

16) Figure 4 and conclusions with regards to impact of GRS as well as ImuB clamp binding mutant. It is possible GRS alone affects clamp stability, irrespective of mutasome function. The authors need to image the clamp after GRS treatment, to assess whether the lack of ImuB localization is because it cannot bind the clamp or because the clamp itself is no longer localized.

The reviewers raise an important point which we realize was not adequately enunciated in our original submission. In their study, Kling et al. (2015; doi:10.1126/science.aaa4690) noted that griselimycin binds the β clamp at the site of multiple protein interactions, likely including the clamp-loader complex, too – an observation reinforced in our own modelling data (Figure 5B, Figure 5 —figure supplement 1) which extends the list of proteins utilizing this site to include ImuB. As the reviewers state, the effect would be to disrupt β clamp localization independent of any effect on ImuB. We agree with this interpretation, evidence for which is provided in Figure 5C where the disruptive impact of griselimycin treatment on mCherry-DnaN localization is clearly apparent. Moreover, griselimycin-mediated replisome collapse has been demonstrated previously (Trojanowski et al., 2019; Antimicrob Agents Chemother. doi:10.1128/AAC.00739-19).

Our intention in the present work is not to claim that griselimycin specifically reduces mutasome function but to provide proof of concept evidence supporting the potential that an on-target inhibitor of the mutasome might offer a valuable addition to current antimycobacterial drugs by decreasing the capacity for DNA damage-induced mutagenesis in M. tuberculosis. We further argue (L439-49) that these observations might alleviate some of the concerns commonly associated with targeting (essential) DNA replication and repair proteins, namely the potential for accelerated/induced (auto)mutagenesis.

17) What is the impact of the clamp-binding-ImuB mutant on clamp localization? Is it similar to GRS treatment?

See response to reviewers’ comment 16, griselimycin binds the β clamp at the site of multiple protein interactions, including that which enables interaction of the β clamp with the clamp-loader complex. In contrast, the β clamp-binding-defective ImuB mutant (imuBAAAAGG) is not expected to affect β clamp loading. Therefore, this specific question was not experimentally addressed. It is worth noting, though, that analysis of the timelapse series featuring G-ImuB and mCherry-DnaN (Figure 3A, 3D) and the corresponding source data strongly suggests recruitment of ImuB to the β clamp foci; as noted earlier (see response to reviewer comment 13), elucidating the temporal dynamics of protein recruitment and localization is a key area of ongoing study.

Other comments:

1. L139 – L143: To make this conclusion, authors need to carry out experiments across a range of doses.

This comment seems to question the inference that expression and recruitment of mutasome components occurs in response to a variety of DNA damaging conditions. As noted earlier (see response to reviewers’ comment 1), the revised manuscript includes experimental data utilizing the fluorescent reporter alleles in the respective knock-out backgrounds as well as in the mCherry-DnaN background, and following exposure to two concentrations of MMC (1×MIC, 0.5×MIC) and to UV irradiation. These observations, together with evidence that imuA’/imuB expression is co-regulated by both PafBC and SOS pathways in mycobacteria and induced following fluoroquinolone-mediated gyrase inhibition (Adefisayo et al., 2021; Nucleic Acids Res. doi:10.1093/nar/gkab1169), suggest the conclusion is reasonable and supported by multiple independent lines of evidence.

2. Figure 2 – individual fluorescence panels need to be shown independently. Currently it is hard to visualize the localizations with accuracy.

As noted (see response to reviewers’ comment 8), we have restructured the Results to improve presentation of the microscopy data, including the addition of quantitative analyses. The original Figure 2 has been incorporated into a revised Figure 3 which includes quantitative analyses of protein localization data (Figure 3D). We note, too, that the timelapse movies are available as Video files, with all original data accessible via the open access Dryad data platform.

3. L276 in Mtb, the PafBC regulatory system also influences Imu expression. The authors need to rephrase.

This is a good point; we apologize for this inaccuracy which has been corrected in the revised version. The sentence (L294-6) now reads:

“Therefore, to test the prediction that the recruitment of EGFP-ImuB and mCherry-DnaN into discernible foci was dependent on the ImuB-β clamp protein-protein interaction, we introduced an egfp-imuBAAAAGG allele (G-imuBAAAAGG) into the DimuB mutant.”

4. Please use β-clamp everywhere (and not only B).

Corrected throughout the revised manuscript.

5. Figure 3A. The difference in elution profiles between ImuA'-ImuB-clamp an ImuB-clamp would suggest that ImuB-clamp interaction alone might be less stable (in absence of ImuA'). Could the authors comment on the same?

Although tempting to speculate a role for ImuA’ in stabilizing the ImuB-β clamp, we have no additional evidence to support any definitive statements about this possibility.

6. Figure 4 legend. Top and bottom panel references need to be updated.

Figure 4 has been replaced with a new Figure 5.

7. L314-315: Details of population analysis are missing in the legends or text.

As noted, the original Figure 4 has been replaced with a new Figure 5, including expanded figure legend.

8. L341. "when" instead of "where".

Revised as requested.

9. L338. "repair/ tolerance" pathways.

Revised as requested.

10. L379. Could the authors clarify? Do they envision DnaE2 acting without clamp? In that case, what could the potential mechanism be?

This is an excellent question. The absence of a clear β clamp-binding motif in DnaE2, plus the inferred lack of a DnaE2-β clamp interaction in yeast two-hybrid assays (Warner et al., 2010; doi:10.1073/pnas.1002614107), suggests that DnaE2 must operate without the β clamp. How this is accomplished is not clear, we can only assume that our understanding of DnaE2 function will be enhanced if/ when expression of the purified recombinant DnaE2 protein is achieved.

11. L405. It is unclear whether the GRS phenotype is due to its action on the replisome, independent of damage / mutasome effects, unless the impact of GRS on clamp alone is tested.

This comment echoes the concern raised previously (see reviewers’ comment 16) which arises from the effectively pleiotropic action griselimycin exerts on multiple replication and repair functions owing to the fact that it binds in the hydrophobic cleft between domains II and III of the β clamp (Kling et al., 2015; doi:10.1126/science.aaa4690), the same interaction site utilized by multiple DNA metabolic proteins including ImuB (Figure 5B, Figure 5 —figure supplement 1). Indeed, our own data (see Figure 5C) demonstrating the disruptive impact of griselimycin treatment on mCherry-DnaN localization support the reviewers’ call for a definitive assay of chemically disrupted mutasome function. The problem, as we noted in our response to reviewer comment 9, is that griselimycin targets both DNA replication – by preventing the DnaE1-b clamp interaction (Kling et al., 2015; doi:10.1126/science.aaa4690) – and mutagenic DNA repair, by preventing ImuB-b clamp focus formation (this manuscript), as well as preventing b clamp localization through the likely inhibition of the clamp loader complex. For this reason, it is not possible to separate the effects of replication inhibition from blocked mutagenesis – a conundrum which, we argued, reinforces the need for an on-target inhibitor of the mutasome. In the absence of such a compound, we have presented microbiological evidence (Figure 5) indicating that treatment of cells with griselimycin eliminates ImuB-β clamp focus formation under mutagenic DNA damage (MMC exposure) conditions, and we complemented this observation with biochemical data (Figure 4) showing disruption of ImuB-DnaN and ImuA’-ImuBDnaN interactions in assays utilizing the purified proteins. Based on these results, and previous work showing that genetic disruption of the ImuB-β clamp interaction eliminates UV-induced mutagenesis, we have argued that inhibiting mutasome recruitment and/or function will substantively eliminate DNA damage-induced mutagenesis in mycobacteria.

12. It is understandable that the authors use M. smeg as a model system to derive conclusions on action of the mutasome. However, the paragraph starting L410 needs to be toned down as all experiments are performed in smeg and not M.tb. Any extrapolation of their conclusions need to be explicitly stated to the reader.

The reviewers’ point is well made. We have revised the concluding paragraph (L45082) to reflect the reliance of the current study on M. smegmatis as mycobacterial model, and to encourage caution in extrapolating the results to M. tuberculosis.

https://doi.org/10.7554/eLife.75628.sa2

Article and author information

Author details

  1. Sophia Gessner

    1. SAMRC/NHLS/UCT Molecular Mycobacteriology Research Unit, DSI/NRF Centre of Excellence for Biomedical TB Research, Department of Pathology, University of Cape Town, Cape Town, South Africa
    2. Institute of Infectious Disease and Molecular Medicine, University of Cape Town, Cape Town, South Africa
    Contribution
    Conceptualization, Formal analysis, Investigation, Methodology, Writing – original draft, Writing – review and editing
    Contributed equally with
    Zela Alexandria-Mae Martin and Michael A Reiche
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-0824-0079
  2. Zela Alexandria-Mae Martin

    1. SAMRC/NHLS/UCT Molecular Mycobacteriology Research Unit, DSI/NRF Centre of Excellence for Biomedical TB Research, Department of Pathology, University of Cape Town, Cape Town, South Africa
    2. Institute of Infectious Disease and Molecular Medicine, University of Cape Town, Cape Town, South Africa
    3. Laboratory of Microbiology and Microsystems, School of Life Sciences, Swiss Federal Institute of Technology in Lausanne (EPFL), Lausanne, Switzerland
    Contribution
    Conceptualization, Investigation, Methodology, Writing – original draft, Writing – review and editing
    Contributed equally with
    Sophia Gessner and Michael A Reiche
    Competing interests
    No competing interests declared
  3. Michael A Reiche

    1. SAMRC/NHLS/UCT Molecular Mycobacteriology Research Unit, DSI/NRF Centre of Excellence for Biomedical TB Research, Department of Pathology, University of Cape Town, Cape Town, South Africa
    2. Institute of Infectious Disease and Molecular Medicine, University of Cape Town, Cape Town, South Africa
    3. Advanced Imaging Center, Howard Hughes Medical Institute, Ashburn, United States
    Contribution
    Conceptualization, Formal analysis, Investigation, Methodology, Writing – original draft, Writing – review and editing
    Contributed equally with
    Sophia Gessner and Zela Alexandria-Mae Martin
    Competing interests
    No competing interests declared
  4. Joana A Santos

    Department of Cell and Chemical Biology, Leiden University Medical Center, Leiden, Netherlands
    Present address
    Instituto de Investigação e Inovação em Saúde, Universidade do Porto, Porto, Portugal
    Contribution
    Conceptualization, Investigation, Methodology, Writing – original draft
    Competing interests
    No competing interests declared
  5. Ryan Dinkele

    1. SAMRC/NHLS/UCT Molecular Mycobacteriology Research Unit, DSI/NRF Centre of Excellence for Biomedical TB Research, Department of Pathology, University of Cape Town, Cape Town, South Africa
    2. Institute of Infectious Disease and Molecular Medicine, University of Cape Town, Cape Town, South Africa
    Contribution
    Formal analysis, Visualization
    Competing interests
    No competing interests declared
  6. Atondaho Ramudzuli

    1. SAMRC/NHLS/UCT Molecular Mycobacteriology Research Unit, DSI/NRF Centre of Excellence for Biomedical TB Research, Department of Pathology, University of Cape Town, Cape Town, South Africa
    2. Institute of Infectious Disease and Molecular Medicine, University of Cape Town, Cape Town, South Africa
    Contribution
    Resources
    Competing interests
    No competing interests declared
  7. Neeraj Dhar

    Laboratory of Microbiology and Microsystems, School of Life Sciences, Swiss Federal Institute of Technology in Lausanne (EPFL), Lausanne, Switzerland
    Present address
    Vaccine and Infectious Disease Organization, University of Saskatchewan, Saskatoon, Canada
    Contribution
    Resources, Supervision, Investigation
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-5887-8137
  8. Timothy J de Wet

    1. SAMRC/NHLS/UCT Molecular Mycobacteriology Research Unit, DSI/NRF Centre of Excellence for Biomedical TB Research, Department of Pathology, University of Cape Town, Cape Town, South Africa
    2. Institute of Infectious Disease and Molecular Medicine, University of Cape Town, Cape Town, South Africa
    3. Department of Integrative Biomedical Sciences, University of Cape Town, Cape Town, South Africa
    Contribution
    Visualization
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-3978-5322
  9. Saber Anoosheh

    1. SAMRC/NHLS/UCT Molecular Mycobacteriology Research Unit, DSI/NRF Centre of Excellence for Biomedical TB Research, Department of Pathology, University of Cape Town, Cape Town, South Africa
    2. Institute of Infectious Disease and Molecular Medicine, University of Cape Town, Cape Town, South Africa
    Present address
    Department of Chemistry and Umeå Centre for Microbial Research, Umeå University, Umeå, Sweden
    Contribution
    Resources
    Competing interests
    No competing interests declared
  10. Dirk M Lang

    Confocal and Light Microscope Imaging Facility, Department of Human Biology, University of Cape Town, Cape Town, South Africa
    Contribution
    Resources, Supervision
    Competing interests
    No competing interests declared
  11. Jesse Aaron

    Advanced Imaging Center, Howard Hughes Medical Institute, Ashburn, United States
    Contribution
    Resources, Methodology
    Competing interests
    No competing interests declared
  12. Teng-Leong Chew

    Advanced Imaging Center, Howard Hughes Medical Institute, Ashburn, United States
    Contribution
    Resources, Methodology
    Competing interests
    No competing interests declared
  13. Jennifer Herrmann

    1. Helmholtz Centre for Infection Research, Helmholtz Institute for Pharmaceutical Research Saarland, Saarbrücken, Germany
    2. German Centre for Infection Research (DZIF), Partner Site Hannover-Braunschweig, Braunschweig, Germany
    Contribution
    Resources, Writing – review and editing
    Competing interests
    No competing interests declared
  14. Rolf Müller

    1. Helmholtz Centre for Infection Research, Helmholtz Institute for Pharmaceutical Research Saarland, Saarbrücken, Germany
    2. German Centre for Infection Research (DZIF), Partner Site Hannover-Braunschweig, Braunschweig, Germany
    Contribution
    Resources, Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-1042-5665
  15. John D McKinney

    Laboratory of Microbiology and Microsystems, School of Life Sciences, Swiss Federal Institute of Technology in Lausanne (EPFL), Lausanne, Switzerland
    Contribution
    Resources, Supervision
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-0557-3479
  16. Roger Woodgate

    Laboratory of Genomic Integrity, Eunice Kennedy Shriver National Institute of Child Health and Human Development, Bethesda, United States
    Contribution
    Funding acquisition, Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-5581-4616
  17. Valerie Mizrahi

    1. SAMRC/NHLS/UCT Molecular Mycobacteriology Research Unit, DSI/NRF Centre of Excellence for Biomedical TB Research, Department of Pathology, University of Cape Town, Cape Town, South Africa
    2. Institute of Infectious Disease and Molecular Medicine, University of Cape Town, Cape Town, South Africa
    3. Wellcome Centre for Infectious Diseases Research in Africa, University of Cape Town, Cape Town, South Africa
    Contribution
    Supervision, Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-4824-9115
  18. Česlovas Venclovas

    Institute of Biotechnology, Vilnius University, Vilnius, Lithuania
    Contribution
    Formal analysis, Investigation, Methodology, Writing – review and editing
    Competing interests
    No competing interests declared
  19. Meindert H Lamers

    Department of Cell and Chemical Biology, Leiden University Medical Center, Leiden, Netherlands
    Contribution
    Conceptualization, Resources, Supervision
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-4205-1338
  20. Digby F Warner

    1. SAMRC/NHLS/UCT Molecular Mycobacteriology Research Unit, DSI/NRF Centre of Excellence for Biomedical TB Research, Department of Pathology, University of Cape Town, Cape Town, South Africa
    2. Institute of Infectious Disease and Molecular Medicine, University of Cape Town, Cape Town, South Africa
    3. Wellcome Centre for Infectious Diseases Research in Africa, University of Cape Town, Cape Town, South Africa
    Contribution
    Conceptualization, Supervision, Funding acquisition, Methodology, Writing – review and editing
    For correspondence
    digby.warner@uct.ac.za
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-4146-0930

Funding

Eunice Kennedy Shriver National Institute of Child Health and Human Development (U01HD085531)

  • Roger Woodgate

Norges Forskningsråd (261669)

  • Digby F Warner

South African Medical Research Council (SHIP and Extramural Unit)

  • Valerie Mizrahi

National Research Foundation

  • Valerie Mizrahi

Howard Hughes Medical Institute (Senior International Research Scholars)

  • Valerie Mizrahi

Leids Universitair Medisch Centrum (LUMC Fellowship)

  • Meindert H Lamers

National Research Foundation (104683)

  • Michael A Reiche

David and Elaine Potter Foundation (PhD Fellowship)

  • Zela Alexandria-Mae Martin

The funders had no role in study design, data collection, and interpretation, or the decision to submit the work for publication.

Acknowledgements

This work was supported by the US National Institute of Child Health and Human Development (NICHD) U01HD085531 (to DFW and RW). We acknowledge the funding support of the Research Council of Norway (R&D Project 261669 'Reversing antimicrobial resistance') (to DFW), the South African Medical Research Council (to VM and DFW); the National Research Foundation of South Africa (to DFW and VM); a Senior International Research Scholars grant from the Howard Hughes Medical Institute (to VM); and a LUMC Fellowship (to MHL). In addition, MAR is grateful to the South African National Research Foundation (NRF) for financial assistance during his PhD training (grant no. 104683) as well as the Whitehead Scientific Travel Award. ZAM is grateful to the University of Cape Town, the David and Elaine Potter Foundation Research Fellowship, and the Swiss Government Excellence Research Scholarship for financial assistance during her PhD.

Senior Editor

  1. Dominique Soldati-Favre, University of Geneva, Switzerland

Reviewing Editor

  1. Christina L Stallings, Washington University School of Medicine, United States

Version history

  1. Received: November 16, 2021
  2. Preprint posted: November 17, 2021 (view preprint)
  3. Accepted: August 1, 2023
  4. Accepted Manuscript published: August 2, 2023 (version 1)
  5. Version of Record published: August 11, 2023 (version 2)

Copyright

This is an open-access article, free of all copyright, and may be freely reproduced, distributed, transmitted, modified, built upon, or otherwise used by anyone for any lawful purpose. The work is made available under the Creative Commons CC0 public domain dedication.

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  1. Sophia Gessner
  2. Zela Alexandria-Mae Martin
  3. Michael A Reiche
  4. Joana A Santos
  5. Ryan Dinkele
  6. Atondaho Ramudzuli
  7. Neeraj Dhar
  8. Timothy J de Wet
  9. Saber Anoosheh
  10. Dirk M Lang
  11. Jesse Aaron
  12. Teng-Leong Chew
  13. Jennifer Herrmann
  14. Rolf Müller
  15. John D McKinney
  16. Roger Woodgate
  17. Valerie Mizrahi
  18. Česlovas Venclovas
  19. Meindert H Lamers
  20. Digby F Warner
(2023)
Investigating the composition and recruitment of the mycobacterial ImuA′–ImuB–DnaE2 mutasome
eLife 12:e75628.
https://doi.org/10.7554/eLife.75628

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https://doi.org/10.7554/eLife.75628

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