A Single-Amino-Acid Substitution in Obg Activates a New Programmed Cell Death Pathway in Escherichia coli

ObgE*-mediated cell death.We previously explored the role of specific amino acid residues of ObgE in sensitivity toward hydroxyurea by screening a mutant obgE overexpression library constructed by error-prone PCR (14). This study additionally yielded a mutant allele encoding ObgE_K268I (ObgE*) that, upon controlled expression, rapidly leads to cell death and is the focus of the current study.

Cell death caused by ObgE* was investigated by determining the number of CFU per milliliter at different time points before and after induction of protein expression with 0.2% arabinose (Fig. 1A). ObgE* causes a decrease in the number of CFU per milliliter of more than 4 orders of magnitude and this in less than 40 min after induction of expression. ObgE*-mediated toxicity was also investigated at different induction levels 2 h after the addition of inducer (Fig. 1B). Significant killing can already be seen at very low induction levels (0.00002% arabinose and higher), highlighting the strong toxicity of this protein. Protein expression levels were verified by Western blot analysis. To confirm that ObgE* causes cell death rather than a switch to a nongrowing state, cells expressing ObgE* were subjected to LIVE/DEAD staining. The result of this staining confirms that ObgE* is bactericidal and leads to eventual loss of membrane integrity (see Fig. S1 in the supplemental material). These experiments were carried out in an E. coli strain that retains the chromosomal obgE gene. We therefore conclude that obgE* is a dominant negative allele which, when expressed, causes rapid cell death in E. coli.

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FIG 1 

Expression of ObgE* triggers a PCD pathway. (A) The number of CFU per milliliter of an exponential-phase culture was monitored. At time zero, expression of ObgE or ObgE* from pBAD/His A was induced by adding 0.2% arabinose. Error bars represent standard errors of the means. (B) ObgE*-mediated toxicity at different induction levels. Error bars represent standard errors of the means. Pairwise comparisons were made between the control condition (no inducer) and induced cultures. #, P ≤ 0.05; ###, P ≤ 0.0001. (C) DiBAC₄(3) staining as a measure of membrane depolarization. Fluorescence intensity was measured by flow cytometry. (D) Microscopic analysis of induced cells stained with the blue DNA dye DAPI and the red membrane-specific dye FM4-64. Bars, 2 µm. (E) DNA fragmentation was determined using the TUNEL assay. Percentages shown represent the fraction of the population with DNA damage. FSC, forward scatter. (F) SEM images showing the formation of blebs upon ObgE* expression. Bars, 1 µm. (G) Staining of externalized phosphatidylserine by FITC-conjugated annexin V. Percentages shown represent the fraction of the population that displays phosphatidylserine on the cell surface. Cells with compromised membrane integrity were omitted from this figure.

ObgE* triggers a PCD mechanism.To demonstrate that the cell death mechanism triggered by ObgE* is programmed, a wide array of PCD markers were investigated. The investigated markers are loss of membrane potential, chromosome condensation, DNA fragmentation, phosphatidylserine exposure, and membrane blebbing.

It was previously shown that bacterial PCD is associated with a loss of membrane potential (7, 8). Similarly, depolarization of the mitochondrial inner membrane is also a key characteristic of the intrinsic pathway of eukaryotic apoptosis (2). By using the potential-sensitive DiBAC₄(3) [bis-(1,3-dibutylbarbituric acid)trimethine oxonol] dye, we found that loss of membrane potential also occurs during ObgE*-mediated cell death (Fig. 1C).

A second marker of PCD, chromosome condensation, was visualized using the blue DNA stain 4′,6-diamidino-2-phenylindole (DAPI). DAPI staining of cells expressing ObgE* shows multiple condensed foci of DNA per cell, whereas the DNA of cells containing the empty vector or cells expressing ObgE occupies the entire cell volume (Fig. 1D). In these experiments, membranes were visualized by concomitant staining with the lipophilic dye FM4-64. In addition to DNA condensation, it is clear that ObgE* also causes a defect in cell division, leading to the formation of cell chains up to 10 cells long.

The third marker of PCD that was detected is fragmentation of DNA, which is also a key characteristic of apoptosis. DNA fragmentation was evaluated using the TUNEL (terminal deoxynucleotidyltransferase dUTP nick end labeling) assay. This assay makes use of a terminal deoxynucleotidyltransferase that couples free 3′ OH groups of DNA to a dUTP residue that is covalently coupled to fluorescein isothiocyanate (FITC), linking fluorescence to DNA damage. Compared to cells carrying the empty vector or cells expressing wild-type ObgE, ObgE* leads to an increase in FITC fluorescence, indicating an increase in DNA damage (Fig. 1E).

Fourth, externalization of phosphatidylserine, a phospholipid that is normally localized in the inner leaflet of the cell membrane, was demonstrated using FITC-conjugated annexin V (Fig. 1G). Annexin V is a protein that specifically binds to phosphatidylserine and is excluded from cells with intact cell membranes. To discriminate between cells exposing phosphatidylserine on the cell surface and cells that have lost membrane integrity, the membrane-impermeant DNA dye propidium iodide (PI) was used. PI-positive cells with compromised membrane integrity are included in Fig. S2 in the supplemental material. An increase in FITC-annexin V binding can be seen upon expression of ObgE*, clearly showing the externalization of phosphatidylserine during the progression of cell death.

A final physiological characteristic that is associated with both apoptosis and programmed necrosis is the formation of protrusions of the membrane, called membrane blebs (2). Because of the drastic structural changes of the eukaryotic plasma membrane during PCD, the membrane structure of E. coli during ObgE*-mediated cell death was visually inspected by means of scanning electron microscopy (SEM). Images show extensive membrane blebbing when ObgE* is expressed (Fig. 1F).

Combined, the loss of viability upon expression of ObgE* and the concomitant phenotypic changes that we observed (loss of membrane potential, DNA condensation and fragmentation, exposure of phosphatidylserine at the cell surface, and severe membrane deformations resembling apoptotic blebs) provide convincing evidence that the pathway triggered by ObgE* is indeed a PCD pathway.

ObgE*-mediated PCD is distinct from previously described bacterial cell death mechanisms.To unravel the mechanism underlying ObgE*-mediated PCD, we investigated the involvement of key regulators of previously described bacterial PCD mechanisms. Because ObgE*-triggered cell death occurs in E. coli and exhibits the main markers of eukaryotic apoptosis, the most likely candidate pathway is so-called apoptosis-like death (ALD) (6–8). Although the molecular pathway involved in ALD has not been fully uncovered yet, it was previously shown that this pathway is activated by treatment with DNA-damaging agents like nalidixic acid, norfloxacin, or mitomycin C (MMC) (6–8), requiring the multifunctional enzyme RecA and its role in the SOS response (6–8). It was also reported that ALD can be activated by treatment with bactericidal antibiotics like gentamicin and ampicillin. In addition to a possible specific effect of these antibiotics on the activation of RecA, it is thought that they lead to the formation of DNA-damaging hydroxyl radicals that can activate ALD similarly to other DNA-damaging agents (7). In the presence of DNA damage, the SOS response is activated by RecA and leads to the expression of over 40 genes involved in DNA damage repair (8). In the case of overwhelming DNA damage, however, high levels of activation of the SOS response presumably lead to ALD rather than repair, thereby preventing a futile attempt to repair DNA that is beyond salvage (8). However, deletion of the recA gene did not restore viability in the presence of ObgE* (Fig. 2A) and had no effect on any of the PCD markers investigated (see Fig. S3 in the supplemental material). Moreover, a β-galactosidase assay with E. coli MG1655 λsfiA::lacZ (15), in which lacZ expression is controlled by the promoter of the SOS gene sfiA (also known as sulA), showed that expression of ObgE* does not result in activation of the SOS response (Fig. 2B), although this strain is also sensitive to ObgE* expression (data not shown). The absence of an SOS response in the presence of ObgE* might seem surprising since the TUNEL assay revealed that expression of ObgE* leads to DNA damage. However, this can be explained as follows. PCD as described in eukaryotes occurs in a hierarchically ordered manner, with some events preceding others (2). Likewise, DNA fragmentation presumably is one of the later steps of ObgE*-mediated PCD since the fraction of cells showing DNA damage is lower than the fraction of cells that have lost viability, have lost their membrane potential, or expose phosphatidylserine on the cell surface at the same time point. It is therefore likely that cells showing DNA fragmentation are lethally injured and are no longer capable of activating the SOS response to repair this damage. Together, these results suggest that RecA plays no direct role in ObgE*-mediated cell death. This is consistent with previous reports suggesting that RecA and ObgE act complementarily rather than interdependently (16, 17).

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FIG 2 

ObgE*-mediated PCD differs fundamentally from previously described bacterial PCD pathways. (A) ObgE*-mediated toxicity in E. coli WM2949 pJAT8-araE and its ΔrecA derivative. (B) A β-galactosidase assay with E. coli MG1655 λsfiA::lacZ shows no activation of the SOS response when ObgE* is expressed. As a positive control, MMC was used at a concentration of 5 µg/ml. #, P ≤ 0.05 versus empty-vector control. (C) ObgE*-mediated toxicity in E. coli MC4100 and its ΔmazEF derivative. (D) ObgE*-mediated toxicity in E. coli WM2949 pJAT8-araE and its ΔyohJ and ΔyohK derivatives. wt, wild type. Error bars represent standard errors of the means.

Another PCD pathway described in E. coli is mediated by the type II toxin-antitoxin (TA) module MazEF (6). To exclude a possible role for the MazF toxin in the execution of ObgE*-mediated cell death, we tested toxicity in E. coli MC4100 ΔmazEF and the corresponding wild-type strain, E. coli MC4100 (6). Expression of ObgE* in E. coli MC4100 ΔmazEF did not result in altered survival compared to E. coli MC4100 (Fig. 2C). Therefore, the mazEF module does not play a role in the execution of ObgE*-mediated cell death. Moreover, the E. coli strain used throughout this study (E. coli WM2949) is defective in the production of the extracellular death factor (EDF)—a quorum-sensing peptide that is essential for the toxic action of MazF (6)—further indicating that this pathway is not involved in ObgE*-mediated cell death.

A final bacterial PCD pathway of interest is mediated by (anti-)holin-like activities and has so far been described only in Staphylococcus aureus and Pseudomonas aeruginosa (9, 18). The cidAB and lrgAB loci are the regulators of this PCD mechanism in S. aureus. We therefore investigated a potential role for the homologous locus in E. coli, yohJK. Single-gene knockouts of both yohJ and yohK were tested for their sensitivity toward ObgE*-mediated cell death (Fig. 2D). Deletion of yohJ or yohK did not alter the viability in the presence of ObgE*, excluding any role for these genes in the execution of cell death and also arguing against a protective function toward ObgE* toxicity. The P. aeruginosa alpB gene that presumably functions as a holin contributing to PCD does not have a homologue in E. coli.

Other bacterial PCD pathways have either previously been shown not to be active in E. coli (10), are induced by specific triggers that do not occur during ObgE* expression (11), or are associated with differentiation and formation of specialized structures (12) and can thus be excluded.

Strains, plasmids, and growth conditions.In this work, E. coli WM2949 (23) with plasmid pJAT8-araE (24) was used except when stated otherwise. In this strain, the chromosomal araFGH and araE genes were deleted and araE expression is controlled by a constitutive promoter on the pJAT8 plasmid, thereby allowing homogeneous expression from the P_BAD promoter (23, 24). Single-gene knockouts of yohJ and yohK were constructed in E. coli WM2949 pJAT8-araE by transducing the corresponding knockouts from the Keio collection (25) as described previously (15). Deletion of the correct gene was confirmed by PCR with primers SPI10966 (GCGCTGATTTCTTAATGTGA) and SPI10967 (ATCAATGTCAGCGGTAATGA) for the deletion of yohJ and primers SPI10968 (CGTAGGTCAGAAAGGATCAG) and SPI10969 (CGCGTGAAACATACATCAAA) for the deletion of yohK.

To investigate ObgE*-mediated effects, strains were transformed with pBAD/His A (Invitrogen), pBAD/His A-obgE (pCMPG13901), or pBAD/His A-obgE* (pCMPG13828). The influence of the N-terminal His tag present in these constructs was investigated by additionally expressing ObgE and ObgE* in their native forms from the pBAD33 vector. The presence or absence of the His tag did not affect the results.

For all phenotypic and toxicity tests, overnight cultures were diluted 100 times in lysogeny broth (LB) containing the appropriate antibiotics, incubated for 2 h at 37°C with continuous shaking at 200 rpm, and induced with 0.2% arabinose for 2 h under the same conditions.

Toxicity tests.To characterize ObgE*-mediated toxicity (Fig. 1A), the number of CFU per milliliter was determined at several time points before and after induction by plating out serial dilutions on LB containing the appropriate antibiotics and 1.5% agar. For all other toxicity tests, the number of CFU per milliliter was determined analogously after 2 h of induction of either ObgE or ObgE*. The percent survival was calculated by dividing the number of cells surviving ObgE* expression by the number of cells present after ObgE expression under the same conditions.

Membrane potential.Loss of membrane potential was determined using the membrane potential-sensitive dye DiBAC₄(3) (Life Technologies), as described previously (21). Fluorescence was measured using a BD Influx cell sorter equipped with 488-nm (green) and 561-nm (red) lasers and standard filter sets.

DNA fragmentation.DNA fragmentation was quantified by the TUNEL assay (Apo-Direct kit; BD Biosciences) as described previously (7). Fluorescence was measured by means of flow cytometry.

DNA condensation.For DAPI (Sigma-Aldrich) and FM4-64 (Life Technologies) staining, cultures were washed and dissolved in 10 mM MgSO₄. The dyes were added simultaneously at a concentration of 10 µg/ml, and cultures were incubated for 10 min in the dark at room temperature. Cells were visualized with the Zeiss Axio Imager Z1. Standard DAPI and Texas Red filters were used.

Phosphatidylserine exposure.Exposure of phosphatidylserine at the cell surface was measured by the FITC-annexin V/dead cell apoptosis kit (Molecular Probes) as described previously (7). Fluorescence was measured by means of flow cytometry.

Scanning electron microscopy.E. coli WM2949 ΔrecA samples were prepared as described above. Cultures were fixed overnight at 4°C in 2.5% glutaraldehyde in 0.1 M cacodylate buffer (pH 7.2) and postfixed in 2% OsO₄ in the same buffer. After serial dehydration, samples were dried at critical point and coated with platinum by standard procedures. Observations were made in a Tecnai FEG ESEM Quanta 200 microscope (FEI), and images were processed by SIS iTEM (Olympus) software.

β-Galactosidase assay.To measure activation of the SOS response, cultures were induced with 0.2% arabinose for 3 h prior to sample collection. The β-galactosidase assay and the calculation of Miller units as a measure of expression level were carried out as described previously (26).