The Locus of Heat Resistance Confers Resistance to Chlorineand Other Oxidizing Chemicals in Escherichia coli
Zhiying Wang,a Yuan Fang,a Shuai Zhi,b David J. Simpson,a Alexander Gill,c Lynn M. McMullen,a Norman F. Neumann,b
Michael G. Gänzlea,d
aUniversity of Alberta, Department of Agricultural, Food and Nutritional Science, Edmonton, Alberta, CanadabUniversity of Alberta, School of Public Health, Edmonton, Alberta, CanadacHealth Canada, Bureau of Microbial Hazards, Ottawa, Ontario, CanadadHubei University of Technology, College of Bioengineering and Food Science, Wuhan, Hubei, People’s Republic of China
ABSTRACT Some chlorine-resistant Escherichia coli isolates harbor the locus of heatresistance (LHR), a genomic island conferring heat resistance. In this study, the pro-tective effect of the LHR for cells challenged by chlorine and oxidative stress wasquantified. Cloning of the LHR protected against NaClO (32 mM; 5 min), H2O2
(120 mM; 5 min), and peroxyacetic acid (105 mg/liter; 5 min) but not against 5.8 mMKIO4, 10 mM acrolein, or 75 mg/liter allyl isothiocyanate. The lethality of oxidizingtreatments for LHR-negative strains of E. coli was about 2 log10 CFU/ml higher thanthat for LHR-positive strains of E. coli. The oxidation of cytoplasmic proteins andmembrane lipids was quantified with the fusion probe roGFP2-Orp1 and the fluores-cent probe BODIPY581/591, respectively. The fragment of the LHR coding for heatshock proteins protected cytoplasmic proteins but not membrane lipids against oxi-dation. The middle fragment of the LHR protected against the oxidation of mem-brane lipids but not of cytoplasmic proteins. The addition of H2O2, NaClO, and per-oxyacetic acid also induced green fluorescent protein (GFP) expression in theoxidation-sensitive reporter strain E. coli O104:H4 Δstx2::gfp::amp. Cloning of pLHR re-duced phage induction in E. coli O104:H4 Δstx2::gfp::amp after treatment with oxidiz-ing chemicals. Screening of 160 strains of Shiga toxin-producing E. coli (STEC) re-vealed that none of them harbors the LHR, additionally suggesting that the LHR andStx prophages are mutually exclusive. Taking our findings together, the contributionof the LHR to resistance to chlorine and oxidative stress is based on the protectionof multiple cellular targets by different proteins encoded by the genetic island.
IMPORTANCE Chlorine treatments are used in water and wastewater sanitation; theresistance of Escherichia coli to chlorine is thus of concern to public health. We showthat a genetic island termed the locus of heat resistance (LHR) protects E. coli notonly against heat but also against chlorine and other oxidizing chemicals, adding toour knowledge of the tools used by E. coli to resist stress. Specific detection of theoxidation of different cellular targets in combination with the cloning of fragmentsof the LHR provided insight into mechanisms of protection and demonstrated thatdifferent fragments of the LHR protect different cellular targets. In E. coli, the pres-ence of the LHR virtually always excluded other virulence factors. It is tempting tospeculate that the LHR is maintained by strains of E. coli with an environmental life-style but is excluded by pathogenic strains that adapted to interact with vertebratehosts.
KEYWORDS locus of heat resistance, chlorine resistance, oxidative stress, Shiga toxinprophage, VTEC, EHEC, uropathogenic Escherichia coli, O104, O157
Citation Wang Z, Fang Y, Zhi S, Simpson DJ,Gill A, McMullen LM, Neumann NF, Gänzle MG.2020. The locus of heat resistance confersresistance to chlorine and other oxidizingchemicals in Escherichia coli. Appl EnvironMicrobiol 86:e02123-19. https://doi.org/10.1128/AEM.02123-19.
Escherichia coli is a common member of the microbiota of the gastrointestinal tractsof vertebrate animals. Most strains of E. coli are nonpathogenic, but pathogenic
strains cause enteric and extraintestinal diseases (1, 2). An important source of exposureto pathogenic E. coli is contaminated water used for irrigation, drinking, and theprocessing of fruits and vegetables (3, 4). Water sanitation can be achieved by ozona-tion, UV light, and, most commonly, chlorination (5, 6). Hypochlorous acid (HOCl), theactive compound of water chlorination, reacts with multiple cellular macromolecules,such as proteins, nucleic acids, and lipids (7). However, bacteria develop resistance tochlorination, and recovery of viability by bacterial cells following chlorination has beenobserved (8). In particular, strains of E. coli isolated from wastewater showed highresistance to chlorine (9).
Chlorine-specific resistance in E. coli involves three chlorine-sensitive transcriptionfactors, hypT, rclR, and nemR, which are activated specifically by chlorine ion oxidation(10–12). The chlorine resistance of E. coli is also mediated through the RpoS-regulatedgeneral stress response (13), the oxidative stress regulons oxyR and soxR (14, 15), andheat shock proteins (16). An overview of the mechanisms affecting chlorine resistancein E. coli is shown in Fig. 1.
A high proportion of chlorine-resistant isolates of E. coli recovered from wastewateralso harbor the locus of heat resistance (LHR) (9), a genomic island that mediatesextreme heat resistance in E. coli (17). The genes on the LHR are predicted to encodeproteins associated with responses to heat shock, cell envelope stress, and oxidativestress (18). The putative mechanisms of LHR-mediated heat resistance (18) overlapthe chlorine resistance mechanisms (Fig. 1). The LHR-encoded heat shock proteinssHSP20, ClpKGI, and sHSPGI may prevent chlorine-mediated protein aggregation.The LHR additionally encodes a homologue of the oxidoreductase thioredoxin. Theactivities of the enzymes encoded in the LHR indicate a potential role in resistanceto chlorine and oxidative stresses during water treatment, although this role hasnot been confirmed experimentally. Moreover, it remains unknown to what extentheat resistance and chlorine resistance overlap with the presence of E. coli virulencefactors. Current knowledge is thus insufficient to assess the frequency of patho-genic strains of E. coli that use LHR-mediated resistance to oxidative water treat-ment. Of particular concern are Shiga toxin-producing E. coli (STEC) strains thatcause foodborne and waterborne illness outbreaks (19–21). The definitive virulencefactor of STEC is Shiga toxin (also termed verotoxin), and STEC strains may havehigh infectivity, with a 1 to 10% risk of infection upon exposure to a single cell (1,22). In addition, a growing body of evidence suggests that uropathogenic E. coli(UPEC) appears to differentially survive wastewater treatments and may also betransmitted by contaminated water (23–26). UPEC strains do not share a defined setof virulence factors but are described on the basis of their ability to cause infectionsin the urinary tract and bladder. This study therefore aimed to investigate whetherthe LHR confers resistance to chlorine in E. coli and to determine the frequency ofthe LHR in strains of STEC and UPEC.
RESULTSThe LHR confers resistance to oxidizing chemicals on E. coli. A previous study
found that 59% of 70 E. coli isolates from chlorinated sewage carried the LHR (9). Incontrast, none of the STEC groundwater isolates tested in this study carried the LHR. Totest the hypothesis that the LHR contributes to chlorine resistance, the heat andchlorine resistance of 10 LHR-positive and 10 LHR-negative E. coli strains isolated fromwastewater was determined (Fig. 2). Ten LHR-positive wastewater isolates were ran-domly selected and matched with 10 randomly selected LHR-negative isolates. TwoLHR-positive strains, E. coli AW1.3 and E. coli AW1.7, and the LHR-negative strain E. coliAW1.7ΔpHR1 served as controls. The reductions in the cell counts of E. coli AW1.7 afterheat and chlorine treatments were about 6 and 2 log10 CFU/ml lower, respectively, thanthe reductions in the cell counts of E. coli AW1.7ΔpHR1, a heat-sensitive derivative ofAW1.7. Similarly, the lethality of chlorine treatment against LHR-positive wastewater
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isolates ranged from 1 to 2 log(N0/N) [where log(N0/N) is the log-transformed ratio ofcell counts before treatment to cell counts after treatment], while chlorine lethalityagainst LHR-negative wastewater isolates ranged from 3.5 to 6 log(N0/N) (Fig. 2).
The contributions of genes encoded by the LHR to resistance to chlorine and otheroxidants were confirmed by cloning fragments of the LHR into E. coli MG1655, followedby determination of the lethality of NaClO, H2O2, peroxyacetic acid (PAA), and KIO4.Acrolein and allyl isothiocyanate (AITC) were additionally used as oxidizing chemicalsthat E. coli may encounter in natural habitats. The LHR or LHR fragments wereintroduced into E. coli MG1655 after cloning into the low-copy-number vector pRK767.E. coli MG1655 transformed with pRK767 served as a vector control (Fig. 3). Cloning ofthe LHR protected against challenge with NaClO, H2O2, and PAA but provided noprotection against KIO4, acrolein, or AITC. Protection against NaClO, H2O2, and PAA wasprovided by all plasmids encoding the full-length LHR or fragment 1 or 2; plasmid pRF3was less effective (against NaClO and H2O2) or ineffective (against PAA). Remarkably,cloning of the LHR or any of its parts into LHR-negative E. coli increased sensitivity toAITC (Fig. 3).
The LHR prevents the oxidation of multiple cellular targets. The oxidation ofcytoplasmic proteins and membrane lipids was assessed in E. coli MG1655 trans-formed with the complete LHR or LHR fragments. Oxidation of cytoplasmic proteinswas assessed with the probe roGFP2_Orp1 and ratiometric fluorescence spectros-
FIG 1 Cytoplasmic determinants of chlorine resistance in E. coli. (A) Chlorine-specific transcription factors. HypT is specifically activated by chlorinethrough methionine oxidation, and Cys4 becomes oxidized and inhibits DNA binding to avoid unnecessary regulation of target genes (10). Proteinsencoded by rclR form a membrane-associated complex responsible for reducing cellular components specifically oxidized by chlorine (11). TheNemR-mediated chlorine response relies on the reversible oxidative modification of the conserved Cys106 (12). (B) Oxidative stress response. Theinsufficiency of NADPH leads to SoxR reduction, and the oxidized Fe�S clusters trigger a conformational change of SoxR (59). Chlorine activates OxyRby the formation of a disulfide bond (14), and then OxyR is regenerated by the glutaredoxin– glutathione– glutathione reductase (Grx/GSH/Gor) systemupon return to nonstress conditions (15). Oxidized OxyR activates the grxA and katG genes, encoding enzymes involved in chlorine resistance, and sufABCfor the repair of damaged Fe–S clusters (60). (C) General stress response. RpoS regulates the transcription of katE (encoding catalase), dps (encoding aDNA-binding protein), tktB (encoding transketolase 2), and grxB (encoding glutaredoxin 2), which act against chlorine resistance (13). (D) Prevention ofprotein aggregation. The reduced Hsp33 in chlorine leads to the simultaneous formation of two intramolecular disulfide bonds and binds to unfoldingsubstrate proteins, whereas DnaK is inactivated because of the low ATP level. When cellular ATP levels are restored, Hsp33 becomes reduced, and boundsubstrates are transferred to the DnaK system for refolding (16). Inorganic polyphosphate (polyP) forms stable complexes with unfolding proteins. Afterthe relief of stress, polyphosphate can be reconverted to ATP, which can be used by DnaK to refold polyphosphate-protected proteins (61). RidA isN-chlorinated by chlorine and then binds to a wide range of unfolded client proteins, preventing their aggregation. Under nonstress condition, thechlorine is removed, leading to the release of the client protein and protein refolding (62).
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copy (27). Oxidation of the probe enhances green fluorescence when excited at488 nm but not at 405 nm. For the nonoxidized probe, the ratio of the fluorescenceintensity at an excitation wavelength of 405 nm to that at an excitation wavelengthof 488 nm was equal to 1. Oxidation of the probe increases the green fluorescenceand decreases the 405/488-nm fluorescence intensity ratio. In comparison to thatfor untreated E. coli MG1655, a decreased fluorescence ratio of roGFP2_Orp1 upontreatment of E. coli MG1655 with NaClO, H2O2, or PAA indicated that the probe wasoxidized (Fig. 4). Insertion of the full LHR sequence reduced the oxidation of theprobe, as indicated by a higher 405/488-nm fluorescence ratio (Fig. 4). pRF1prevented probe oxidation as effectively as the full-length LHR, while pRF2 or pRF3had a fluorescence ratio similar to that of the empty vector, pRK767, indicating thatpRF2 and pRF3 had little or no effect on probe oxidation. The protective effects ofthe LHR against the oxidation of cytoplasmic proteins can thus be attributed to theheat shock proteins encoded by fragment 1 on pRF1.
The oxidation of membrane lipids was determined with the fluorescent probeC11-BODIPY581/591, which is sensitive to lipid peroxides in membranes (28). The pres-ence of the LHR consistently decreased the population of oxidized cells after treatmentof E. coli MG1655 or AW1.7 with oxidizing chemicals; a corresponding increase in thepopulation of unoxidized cells was observed after H2O2 or PAA treatment (Fig. 5; seealso Table S2 in the supplemental material). A consistent effect of fragments of the LHRwas observed only after treatment with H2O2; the presence of LHR fragment pRF3 orpRF1-2 resulted in a high number of unoxidized cells and a correspondingly reducednumber of oxidized cells following treatment with H2O2.
The LHR reduces the peroxide-induced induction of the Stx prophage in E. coliO104:H4. The expression of the late genes in the Shiga toxin prophage in E. coliO104:H4 is induced by oxidative stress (29). Quantification of green fluorescent protein(GFP) fluorescence in the reporter strain E. coli O104:H4 Δstx2::gfp::amp is thus anindirect indication of cytoplasmic oxidative stress (29). The LHR or LHR fragments werecloned into E. coli O104:H4 Δstx2::gfp::amp in order to determine the effect of the LHRon the expression of the Shiga toxin prophage. H2O2, NaClO, and PAA induced GFPexpression in E. coli O104:H4 Δstx2::gfp::amp (Fig. 6). Cloning of pLHR, pRF1, or pRF1-2reduced the expression of the prophage after treatment with any of the three oxidizingchemicals but not after treatment with the positive control mitomycin C (MMC).Cloning of pRF3 reduced prophage induction after treatment with mitomycin C but notafter treatment with NaClO or PAA; a modest reduction in the percentage of GFP-
FIG 2 Lethality of treatments with heat or chlorine to 23 strains of E. coli. Treatment lethality is expressedas the log-transformed ratio of cell counts before treatment (N0) to cell counts after treatment (N). Filledbars represent cells treated at 60°C for 5 min; open bars represent cells treated with 15 mM NaClO for25 min. E. coli AW1.7ΔpHR1 and FUA strains are LHR negative; the other 12 strains are LHR positive. Dataare shown as means � standard deviations for three independent experiments.
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expressing cells was observed after treatment with H2O2 (Fig. 6). The LHR thus reducesthe level of oxidative-stress-induced prophage expression, and this effect is predomi-nantly attributable to fragment 1 on pRF1.
The inhibition of prophage induction by the LHR was confirmed by quantification ofgfp expression by reverse transcription-quantitative PCR (RT-qPCR). The expression ofgfp after induction by mitomycin C or H2O2 relative to that for uninduced controls wasquantified in E. coli O104:H4 Δstx2::gfp::amp(pLHR) and O104:H4 Δstx2::gfp::amp-(pRK767). The LHR reduced the expression of gfp after H2O2 treatment but not aftermitomycin C treatment (Fig. 7A), a finding consistent with the data obtained by flowcytometry (Fig. 6). To determine whether the effect of the LHR on prophage inductionrelates to the RecA-dependent SOS response (29), the expression of recA was alsoquantified. The LHR did not affect the expression of recA after induction with mitomycinC or H2O2 (Fig. 7B).
The LHR does not interfere with the induction of some but not all of the Stxprophages in E. coli. The influence of the LHR on the expression of the Shiga toxin
FIG 3 (A) Schematic representation of the locus of heat resistance and putative functions encoded bygenes located on the genomic island. Genes that are expressed in E. coli MG1655(pLHR) are framed andprinted in boldface. Proteins are color coded based on their predicted functions: red, heat shock proteins;yellow, hypothetical proteins with a possible relationship to envelope stress; blue, proteins related tooxidative stress; orange, DegP, with possible relationships to signaling in the Cpx, EvgA, and �E pathways.Genes carry the subscript “GI” for genomic island if an orthologue of the same gene is present in E. coligenomes. Open reading frames are numbered if there is no known function associated with the genes.Predicted promoters are indicated by arrows; the gray caret on the right indicates a predicted terminator.The three fragments of the LHR that were used to assemble pRF1, pRF2, pRF3, and pRF1-2 are indicatedbelow the diagram. Modified according to reference 18. (B) Lethality of treatment with different oxidantsto cultures of E. coli MG1655 expressing the LHR or specific fragments of the LHR that are carried on pRF1,pRF2, pRF3, or pRF1-2. Treatment lethality is expressed as the log-transformed ratio of cell counts beforetreatment (N0) to cell counts after treatment (N). Cells were treated with 32 mM NaClO, 120 mM H2O2,105 mg/liter peroxyacetic acid (PAA), 5.80 mM KIO4, 10 mM acrolein, or 75 �g/ml allyl isothiocyanate(AITC) for 5 min. Reactions were terminated by adding an equivalent volume of 10% Na2S2O3 as areducing agent. Values for different plasmids within a treatment that do not have a common superscriptare significantly different (P � 0.05). Data are shown as means � standard deviations for three indepen-dent experiments.
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prophage in E. coli O104:H4 implies that the genetic island may interfere with theconversion of the prophage to the lytic cycle. The low prevalence of the LHR observedin STEC may indicate that the presence of the LHR selects against Shiga toxin pro-phages, or vice versa. About 2% of E. coli strains harbor the LHR (17); however, only0.5% of 615 clinical isolates of STEC and/or E. coli O157 have been reported to harborthe LHR (30). Screening of 100 STEC strains revealed that none of them carried the LHRor parts of the LHR and that none of them exhibited a level of heat resistanceequivalent to that of LHR-positive strains (31) (see Table S3 in the supplementalmaterial). The sequence diversity of the late promoter pR= region of Stx prophagesaffects the efficiency of Stx expression in different strains of STEC (32). Therefore, theeffects of the LHR on gene expression from different pR= regions in native STEC and inheterologous hosts were compared. To investigate if the LHR inhibits the expression ofthe same pR= region in different strains, pLHR and pRK767 were introduced into E. coliO157:H7 CO6CE900 (FUA1399), E. coli O157:H7 1935 (FUA1303), and E. coli O45:H205-6545 (FUA1311). These strains were additionally transformed with Pp1302::rfp::chl,
FIG 4 Oxidation of roGFP2-based probes expressed in E. coli MG1655 with different plasmids (pRK767,pLHR, pRF1, pRF2, pRF3, pRF1-2) after exposure to different oxidants. The ratio of the fluorescenceintensity at an excitation wavelength of 405 nm to the fluorescence intensity at an excitation wavelengthof 488 nm was calculated to indicate the oxidation level in the cytoplasm. Cells either were left untreated(control) or were treated for 5 min with 32 mM NaClO, 120 mM H2O2, or 105 mg/liter peroxyacetic acid.Values for different plasmids within a treatment that do not have a common superscript are significantlydifferent (P � 0.05). Data are means � standard deviations for three independent experiments.
FIG 5 Flow cytometric quantification of the oxidation of membrane lipids in E. coli MG1655 with different plasmids(pRK767, pLHR, pRF1, pRF2, pRF3, pRF1-2) by use of C11-BODIPY581/591 after different treatments. Shown are thelevels of stained, unoxidized cells (A) or stained, oxidized cells (B) as percentages of the total cell population. Cellswere treated for 5 min with 32 mM NaClO, 120 mM H2O2, or 105 mg/liter peroxyacetic acid. Values for differentplasmids within a treatment that do not have a common superscript are significantly different (P � 0.05). Data aremeans � standard deviations for 10 independent experiments.
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which carries the rfp gene, coding for red fluorescent protein (RFP), under the controlof the late promoter of the E. coli O104:H4 11-3088 prophage (see Tables 3 and 4) (32).Prophage expression was induced with H2O2, and RFP expression was quantified byflow cytometry. The presence of the LHR resulted in a reduced proportion of cellsexpressing RFP in all three strains, but the extent of inhibition differed among strains(Fig. 8). The effect of the LHR on the expression of the pR= region was further evaluatedwith RFP under the control of promoter pR= regions derived from E. coli FUA1399,FUA1303, and FUA1311. Reporter plasmids were cloned into homologous and heter-ologous hosts. The presence of the LHR decreased the proportion of E. coli FUA1399cells that expressed RFP from promoters p1399-28 and p1399-79, but the proportion ofcells that expressed RFP remained unchanged with p1303-s1 and increased withp1303-2a in E. coli FUA1303 and with p1311 in E. coli FUA1311. These results demon-strate that the induction of the Shiga toxin promoter in the presence of the LHR was
FIG 6 Quantification of stx expression in the reporter strain E. coli O104:H4 Δstx2::gfp::amp (FUA1302)with different plasmids (pRK767, pLHR, pRF1, pRF2, pRF3, pRF1-2) after exposure to different inducers.Exponential-phase E. coli O104:H4 Δstx2::gfp::amp was incubated at 37°C with the addition of eithermitomycin C (MMC; 0.5 mg/liter) for 3 h, NaClO (10 mM) for 1 h, H2O2 (2.5 mM) for 1 h, or peroxyaceticacid (PAA; 50 mg/liter) for 1 h. GFP fluorescence was quantified by flow cytometry. Values for differentplasmids within each treatment that do not have a common superscript are significantly different(P � 0.05). Data are means � standard deviations for at least three independent experiments.
FIG 7 Expression of gfp (A) and recA (B) in E. coli O104:H4 Δstx2::gfp::amp after mitomycin C or H2O2
treatment. Relative gene expression was quantified by RT-qPCR with gapA as the housekeeping gene anduntreated exponential-phase cultures as reference conditions. Exponential-phase cultures were treatedwith LB broth containing mitomycin C (MMC; 0.5 mg/liter) or with H2O2 (2.5 mM) for 40 min. Values fordifferent plasmids within a treatment highlighted with an asterisk are significantly different (P � 0.05).Data are means � standard errors for three independent experiments.
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diverse, due to the sequence diversity of the pR= region and prophage-encodedregulatory proteins in different strains.
The LHR is correlated with the absence of most UPEC virulence factors. SeventyE. coli wastewater isolates were screened for the LHR, a wastewater marker (IS30), andfive virulence factors of UPEC to identify LHR-positive strains of UPEC (Table 1). Amongthe 70 isolates, 22.9% (16/70) were LHR positive, and 40% (28/70) carried virulencefactors that are typical for UPEC. Two of 16 LHR-positive strains carried fyuA, whichencodes the yersiniabactin receptor (33). The other 14 LHR-positive strains excludedUPEC virulence factors. Conversely, 26 of the 28 strains with UPEC virulence factors, andin particular all strains with multiple virulence factors, were LHR negative (Table 1). BothIS30-positive strains carried the LHR, matching prior results (9).
FIG 8 Effect of the LHR on the expression of RFP under the control of pR= promoters derived from Shigatoxin-producing E. coli. Data are the percentages of cells expressing RFP after induction with H2O2. RFPfluorescence was quantified by flow cytometry. The graph compares STEC strains carrying pLHR withSTEC strains carrying pRK767 as a control. Promoter activity was assessed with pR=::rfp::chl constructscloned into three strains of STEC. Plasmid pR=::rfp::chl, containing the pR= promoter derived from E. coliO104:H7 (FUA1302), was cloned into all strains. In addition, each strain was transformed with a plasmidharboring a fusion of pR=::rfp::chl with the pR= promoter(s) present in that strain, i.e., p1399-28 andp1399-79 in E. coli O157:H7 CO6CE900 (FUA1399), p1303-s1 and p1303-2a in E. coli O157:H7 1935(FUA1303), and p1311 in E. coli O45:H2 05-6545 (FUA1311). Significant differences (P � 0.05) in promoteractivity between strains with pLHR and strains with pRK767 are indicated by asterisks. Data are means �standard deviations for three independent experiments.
TABLE 1 Screening of 70 E. coli wastewater strains for the locus of heat resistance, IS30,and virulence genesa
LHR-positive strains of E. coli have been observed at increased frequency in waste-water after chlorination, pointing to a contribution of the LHR to chlorine resistance (9).This study documented, with multiple and complementary experimental methods (27,28, 34), that the LHR protects against oxidative stress. The use of multiple methods toquantify the survival and oxidation of cellular components not only confirmed theprotective effect of the LHR but also provided information on the mechanisms ofprotection.
The LHR and stress resistance in E. coli. The LHR protected against NaClO, H2O2,and peroxyacetic acid but not against KIO4, acrolein, or AITC. Acrolein is generated invivo by the chemical conversion of �-hydroxypropionaldehyde, a metabolite of intes-tinal bacteria including Salmonella enterica and Lactobacillus reuteri (35, 36). AITC isformed from glucosinolates in plants of the Brassicaceae family upon cellular injury andreacts with sulfhydryl groups and disulfide bonds (37). KIO4 has been suggested tocontribute to electrochemical inactivation of E. coli, but the mechanisms of activityremain unclear (38). The selective protective effect of the LHR against oxidizing organicand inorganic chemicals may relate to differences in the chemical reactivity andpermeance of the compounds for the cytoplasmic targets. The presence of the LHRconsistently increased resistance to chlorine and peroxides, which are a mainstay in thesanitation of water and in food-processing plants; therefore, further experimentsfocused on these chemicals.
The LHR encodes proteins that are involved in protein folding or disaggregation andmay protect against oxidative stress (Fig. 1) (17). The heat shock proteins encoded bythe protein homeostasis module (fragment 1), sHSP20, ClpKGI, and sHSPGI, preventprotein aggregation or disaggregate proteins (18, 39). In agreement with the functionof these proteins, transformation of E. coli with this portion of the LHR provided thegreatest protection of cytoplasmic proteins from oxidation but was less protective ofmembrane lipids. LHR fragment 3 did not protect against chlorine or the oxidation ofcytoplasmic components but decreased the oxidation of membrane lipids (Fig. 3 to 5).The role of the oxidation of membrane lipids in chlorine resistance is poorly docu-mented (Fig. 1). The oxidative stress module (fragment 3) encodes thioredoxin, KefB,and DegP. Thioredoxin protects against oxidative stress through thiol-disulfide ex-change reactions (40). KefB may have a function related to oxidative stress; it is a K�/H�
antiporter that protects against two electrophiles, methylglyoxal and N-ethylmaleimide(41). The serine protease DegP is a periplasmic chaperone that prevents or reduces theiron-induced oxidation of membrane proteins (42). Fragment 2 encodes YfdX1GI andYfdX2, two hypothetical proteins of unknown function, and HdeDGI, a stress proteinwhose relation to oxidative stress is unknown (43). These genes protected againstchlorine and provided limited protection against the oxidation of membrane lipids butdid not prevent the oxidation of cytoplasmic proteins. These findings not only confirmthe previous designation of fragment 2 as an “envelope stress module” (18) but alsosuggest additional, undescribed mechanisms of protection. Taking these findingstogether, the contribution of LHR to chlorine resistance is based on the protection ofmultiple cellular targets by different parts of the LHR. The multifaceted mechanisms ofprotection provided by the LHR also explain its contribution to resistance to multiplestressors, including heat, chlorine, and peroxides (17, 18, 39, 44).
Two distinct LHR variants in E. coli confer equivalent heat resistance (45). LHR2 is a19-kb genomic island that contains five open reading frames (ORFs) that are not carriedby the LHR but lacks two proteins encoded by the LHR (17, 45). Proteins that arepresent in both versions of the genomic island include sHSP20, ClpKGI, sHSPGI, YfdX1GI,YfdX2, KefB, orf15, and DegP; these proteins appear to be essential for the role of thegenomic island in bacterial stress resistance.
The LHR and the virulence of E. coli. The LHR is a mobile genetic element thatoccurs in diverse members of the Gammaproteobacteria and Betaproteobacteria. TheLHR has been identified in one Salmonella strain and in several isolates of Cronobacter
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spp. and Klebsiella spp. (45, 46). The frequency of the LHR in STEC is very low comparedto that in the general population of E. coli (30; this study). Genes coding for Shiga toxinproduction are invariably found in the late regions of prophages. Shiga toxin produc-tion by E. coli has been proposed to protect against predatory protozoa, which aresignificant E. coli predators in the rumen but are less prevalent in other ecosystems thatare relevant for the evolution of E. coli (47). Expression of the Shiga toxin is upregulatedin the lytic cycle of the phage, which is lethal to the host cell, but it has beenhypothesized that Shiga toxin production by engulfed cells will kill the predator,reducing predation on the remainder of the population. Accordingly, STEC strains arehighly associated with diverse ruminant hosts (48). One explanation of the low overlapof the LHR and Shiga toxin production in E. coli is ecological incompatibility; i.e.,ecosystems that select for maintenance of the Shiga toxin select against the LHR, andvice versa. Molecular incompatibility is an alternative explanation; i.e., the LHR inter-feres with the expression of Shiga toxin prophages and hence reduces or abolishes theecological advantage conferred by lysogeny.
The lytic cycle of the Shiga toxin prophage is regulated by the DNA repair proteinRecA and is induced in response to DNA damage and/or oxidative stress. If the LHRprotects against oxidative stress, it also reduces Shiga toxin production in response topredation by protozoa and may reduce the selective pressure to maintain the Shigatoxin prophages in STEC. Reduced expression of GFP as an indicator of Stx expressionwas observed in E. coli O104:H4 but not for all Shiga toxin prophages in other STECstrains. Different Shiga toxin prophages respond differently to inducing agents (32, 49).In this study, it was observed that all the prophage promoters investigated respondedto H2O2, but not all responded to mitomycin C, which is routinely used to inducelambdoid prophages, including Shiga toxin prophages. In summary, the LHR interferedwith the induction of some, but not all, Shiga toxin prophages. Thus, it does not appearthat the LHR is generally incompatible with the contribution of Shiga toxin to theecological fitness of E. coli in response to protozoan predation. The assumption that theLHR increases fitness in ecological niches in which Shiga toxin prophages do notprovide a selective advantage is a more likely explanation for the low cooccurrence ofthese two mobile genetic elements in E. coli.
Naturalized strains of E. coli found in wastewater possess the LHR, are resistant tochlorine, and carry as many as 44 different virulence genes associated with UPEC (9, 50).UPEC strains appear to differentially survive wastewater treatment processes, includingchlorination and UV-C irradiation (24, 25, 51). Interestingly, the proportion of antibiotic-resistant UPEC cells has been reported to be higher in water after treatment withchlorine (52, 53). Collectively, these studies led us to hypothesize that antibioticresistance correlates with the increased likelihood of UPEC surviving wastewater treat-ment (chlorination and oxidation). LHR-positive E. coli and UPEC strains were found tocoexist in wastewater, and the prevalence of the LHR in wastewater isolates of E. coli,23%, is about 10-fold higher than that in the general population of E. coli, 2%.Surprisingly, this study found low cooccurrence of the LHR and UPEC virulence factorsin 70 wastewater isolates. This finding should be confirmed by screening a largernumber of strains or genomes, but it suggests that the LHR may not account for theresiliency of UPEC in surviving wastewater treatment processes and attests to thediverse mechanisms by which microbes evolve resistance to wastewater treatmentprocesses.
In conclusion, the LHR confers resistance to heat and oxidative stress on E. coli. Theprotein homeostasis module provided protection to the contents of the cytoplasm,while the oxidative stress module provided protection to membrane lipids. In addition,the reduction effect of the LHR on Shiga toxin expression was specific to the latepromoter regions and regulatory proteins. The low prevalence of the LHR in STEC andUPEC strains indicates that the selective pressure for maintenance of the LHR in E. coliis different from the selective pressure that maintains Shiga toxin prophages and UPECvirulence factors. This study also shows that water chlorination selects for LHR-positiveE. coli strains that are heat and chlorine resistant.
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MATERIALS AND METHODSCollection and screening of E. coli from wastewater and groundwater. Ten LHR-positive strains
and 10 LHR-negative strains of E. coli were selected from a total of 70 strains that had been isolated fromwastewater previously (9). In addition, 70 E. coli isolates collected from wastewater effluents duringroutine monitoring programs were provided by a municipal water treatment plant in Alberta, Canada.These samples were collected from undigested sludge, digested sludge, and biosolids from a lagoonprior to agricultural land application. Strains of E. coli were subcultured from most-probable-numbertubes in EC broth (Oxoid) after 24 h of incubation. The methods used to confirm the identification of E.coli and to detect specific virulence or resistance markers are outlined below. Sixty-five groundwaterisolates positive for stx1, stx2, or both were collected from routine screening of well water samplessubmitted to the Alberta Provincial Laboratory for Public Health. Isolates were screened for the presenceof the LHR with primers targeting three fragments of the LHR in E. coli AW1.7. The primers used in thisstudy are listed in Table 2; a list of isolates used in the study is provided in Table S1 in the supplementalmaterial.
Determination of chlorine resistance. Chlorine resistance was determined by mixing 200 �l ofovernight cultures with 6.5 �l of a 3% (wt/wt) sodium hypochlorite solution (Sigma-Aldrich, St. Louis, MO)to a concentration of 15 mM NaClO, followed by incubation for 25 min at 20°C. The reaction wasterminated by the addition of 6.5 �l of 10% Na2S2O3 (Sigma-Aldrich, St. Louis, MO). Treatment conditionswere selected to achieve a reduction in cell counts ranging from about 1 to 7 log CFU/ml. To benchmarkthe effect of the LHR on chlorine resistance against the previously described contribution of the LHR toheat resistance, the heat resistance of overnight cultures of E. coli was determined as describedpreviously (17). Cell counts of cultures before and after treatment were determined by surface plating onLB agar and incubation at 37°C for 24 h. Results are expressed as log-transformed ratios of cell countsbefore treatment to cell counts after treatment [log(N0/N)].
Effect of the LHR on resistance to oxidizing chemicals. To assess the contributions of differentregions of the LHR to survival under oxidative stress, E. coli MG1655 was transformed with plasmidspRK767, pLHR, pRF1, pRF2, pRF3, and pRF1-2 (17) (Tables 3 and 4). The pRK767 plasmid was used asa vector control for MG1655. Oxidative stress was induced by treatment of the transformants for5 min with 32 mM NaClO, 120 mM H2O2 (30% [wt/wt] in H2O; Sigma-Aldrich, St. Louis, MO),
TABLE 2 PCR primers for amplification used in this study and melting temperatures for multiplex HRM analysis-qPCR detection of targetgenes
Gene target Primer name Primer sequence (5=–3=) TAa (°C) Tm
b (°C) Source or reference
LHR-F1 LHRmF1-F GCCCGGTGTCGAGGAGAAGG 62.5 This studyLHRmF1-R AAGAATGGCCGAGTTCATTGGAGG 59.4 This study
LHR-F2 LHRmF2-F GCGCGATGCCAAGCAGAACG 62.4 This studyLHRmF2-R TGAACGCGCCATTGACCAAGG 60.8 This study
LHR-F3 LHRmF3-F GGAGACGCTGAGCTTTCTGTCCG 61.7 This studyLHRmF3-R CGCAGCAGCCAGTAGGTCG 61.1 This study
LHR-F1 LHRF1-F TCGTCTACAAGCGTGATCC 58 88.03 � 0.27 This studyLHRF1-R GTCACGCAAACGGATGG
LHR-F2 LHRF2-F TCCGAGCTAAGGTGGAATG 58 82.84 � 0.22 This studyLHRF2-R CTGCTTGCCACTTCGTTATC
LHR-F3 LHRF3-F ACCGAGCTGATTGAAGGA 58 86.13 � 0.32 This studyLHRF3-R AGCGACACCACGATGAT
chl/kan Priming site 1 ATCGAAAGCTTGTGTAGGCTGGAGCTG 60 55Priming site 2 TAAGGAGGATATTCATATGM13/pUC-F CCCAGTCACGACGTTGTAAAACG 60 Invitrogen
105 mg/liter peroxyacetic acid (PAA; 32% [wt/wt] in acetic acid; Sigma-Aldrich, St. Louis, MO),5.80 mM KIO4 (Thermo Fisher Scientific, Waltham, MA, USA), 10 mM acrolein (Thermo Fisher Scien-tific, Waltham, MA, USA), or 75 mg/liter allyl isothiocyanate (AITC; Alfa Aesar Co., Inc.). Reactions wereterminated by adding 10% Na2S2O3 to achieve a final concentration of 16 to 63 mM. The cell countswere determined as described above.
Measurement of cytoplasmic oxidation by a roGFP2-based probe. The fusion protein roGFP2-Orp1 was designed to measure H2O2 in biological systems (27). A plasmid encoding roGFP2-Orp1 wastransformed into E. coli MG1655, along with plasmids carrying the whole LHR or part of the LHR (pRK767,pLHR, pRF1, pRF2, pRF3, pRF1-2) (Tables 3 and 4). Ampicillin (100 mg/liter) and tetracycline (15 mg/liter)were added to the cultivation media to maintain both plasmids. Exponential-phase cultures of transfor-mants were incubated with 100 �M IPTG (isopropyl �-D-1-thiogalactopyranoside; Thermo Fisher Scien-tific, USA) at 37°C overnight to induce the expression of reduction-oxidation-sensitive green fluorescentprotein 2 (roGFP2). Cells were washed twice in phosphate-buffered saline (PBS) buffer (pH 7.4) and weretreated with NaClO (32 mM), H2O2 (120 mM), or PAA (105 mg/liter) for 5 min. Nontreated cells served ascontrols. Reactions were terminated by adding 10% Na2S2O3. Cultures (100 �l) were placed in the wellsof a black, clear-bottom 96-well plate (Corning; Corning, NY, USA), and fluorescence was measured atexcitation wavelengths of 405 and 488 nm and at the emission wavelength of 530 nm. The ratio of thefluorescence intensity obtained at the excitation wavelength of 405 nm to that obtained at 488 nm wasused to evaluate the oxidation level of roGFP2 (27).
TABLE 3 Plasmids used in this study and antibiotics used for plasmid maintenance
Plasmid type and name Antibiotic (concn [mg/liter]) used for maintenance Source or reference
E. coli O45:H2 05-6545 (FUA1311) stx1 pRK767::kan, pLHR::kan, and Pp1302::rfp::chl or Pp1311::rfp::chl
E. coli O157:H7 CO6CE900 (FUA1399) stx2a pRK767::kan, pLHR::kan, and Pp1302::rfp::chl, Pp1399-28::rfp::chl, orPp1399-79::rfp::chl
aThe FUA number (the strain number of the Food Microbiology culture collection at the University of Alberta) is used for plasmid nomenclature (i.e., the origin of thepromoter is identified by the FUA number).
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Determination of membrane lipid oxidation by C11-BODIPY581/591. E. coli MG1655 transformedwith different plasmids (pRK767, pLHR, pRF1, pRF2, pRF3, pRF1-2) (Tables 3 and 4) was treated withNaClO (32 mM), H2O2 (120 mM), or PAA (105 mg/liter) for 5 min. Oxidized E. coli served as a nonstainedcontrol. Nonoxidized E. coli prepared with C11-BODIPY581/591 (Thermo Fisher Scientific, Waltham, MA,USA) served as a nontreatment control. In brief, 1.5 ml of each culture was washed with 2 ml ice-cold50 mM Tris-HCl (pH 8.0) containing 20% (wt/vol) sucrose and was resuspended in 2 ml Tris-HCl buffer.The outer membrane was disrupted by the addition of 0.2 ml lysozyme solution (5 mg/ml lysozyme in0.25 M Tris-HCl [pH 8.0]) and 0.4 ml EDTA (0.25 M; pH 8.0) (54). After incubation at 37°C for 30 min, cellpellets were suspended with 10 mM citrate buffer (pH 7.0) with the addition of 10 �M C11-BODIPY581/591,followed by incubation in the dark for 30 min at 37°C and 200 rpm. Flow cytometry was performed usinga BD LSRFortessa X-20 system (BD Biosciences, San Jose, CA, USA) equipped with 488-nm excitation froma blue air laser at 50 mW and 561-nm excitation from a yellow air laser at 50 mW to excite green(530 � 30 nm) and red (586 � 15 nm) fluorescence. Single cells were quantified by forward scatter andside scatter gating on the flow cytometer. In brief, samples were centrifuged, resuspended, and dilutedwith 1 ml of PBS (pH 7.4) to keep the detected cell number per second (e/s) in the range of 300 to �3,000events. Sample injection and acquisition were started simultaneously and continued until 10,000 eventswere recorded. Data were recorded by BD FACSDiva software and were analyzed by FlowJo (both fromBD Biosciences, San Jose, CA, USA). The single-cell population was defined by selecting the cellpopulation located along the diagonal of the “FSC-A; FSC-H” dot plot. The population was divided intofour subpopulations by red and green fluorescence reference lines. The reference lines were determinedfrom untreated samples, where at least 96% of the population was negative for red and greenfluorescence.
Flow cytometric determination of GFP fluorescence. E. coli O104:H4 Δstx2::gfp::amp is derivedfrom a STEC strain, with in-frame replacement of the prophage-harbored stx2 gene with a gfp::ampcassette (29). The fusion of GFP in the Stx2 prophage provides a reporter for protein expressionunder the control of the Shiga toxin promoter. E. coli O104:H4 is tetracycline resistant, a character-istic that interferes with the antibiotic resistance of the other plasmids used in this study. Therefore,the tetracycline resistance gene on plasmids pRK767, pLHR, pRF1, pRF2, pRF3, and pRF1-2 wasreplaced with a chloramphenicol resistance gene from pKD3 (55). PCRs were carried out usingPhusion High-Fidelity DNA polymerase (Thermo Scientific) according to the manufacturer’s guide-lines. The gene encoding chloramphenicol resistance was amplified by the Priming site 1 andPriming site 2 primers (Table 2). Plasmids pRK767, pLHR, pRF1, pRF2, pRF3, and pRF1-2 were digestedwith HindIII to remove the tetracycline resistance gene, and the chloramphenicol resistance genewas then ligated into the plasmid as a HindIII/HindIII insert. The direction of the insert was confirmedby amplification with the primers targeting Priming site 1 and M13/pUC-F (Table 2). The recombinantplasmids were electroporated into E. coli O104:H4 Δstx2::gfp::amp (Tables 3 and 4). Exponential-phase cultures of these transformants were treated with mitomycin C (MMC; 0.5 mg/liter) for 3 h orwith NaClO (10 mM), H2O2 (2.5 mM), or PAA (50 mg/liter) for 1 h. Control samples were incubated inthe same manner without stressors. The method used for the detection of the population offluorescent cells was similar to that described in reference 29. Samples were divided into twosubpopulations, and the percentage values of GFP-positive cells were calculated.
Quantification of stx2 prophage expression in LHR-positive and LHR-negative strains in re-sponse to oxidative stress. Exponential-phase cultures of E. coli O104:H4 Δstx2::gfp::amp(pRK767) andE. coli O104:H4 Δstx2::gfp::amp(pLHR) were centrifuged and were resuspended in LB broth containingmitomycin C (0.5 mg/liter) or H2O2 (2.5 mM), followed by incubation at 37°C for 1 h. Correspondingtreatment of cultures without any addition served as a control. After treatment, cells were harvestedfrom samples; RNA was isolated using the RNAprotect Bacteria reagent and the RNeasy minikit(Qiagen) and was reverse transcribed to cDNA with a QuantiTect reverse transcription kit (Qiagen)according to the manufacturer’s protocols. The expression of gfp and recA was quantified by usingSYBR green reagent (Qiagen) and a 7500 Fast PCR system (Applied Biosystems, Foster City, CA, USA).Negative controls included DNase-treated RNA and nontemplate controls. The gene coding forglyceraldehyde-3-phosphate dehydrogenase A (gapA) served as the reference gene. The ratios ofexpression of the gfp and recA genes in E. coli O104:H4 Δstx2::gfp::amp(pRK767) to their expressionin E. coli O104:H4 Δstx2::gfp::amp(pLHR) under induced and control conditions were calculatedaccording to the method of Pfaffl (57). The primers used for the quantification of gene expressionare listed in Table 2.
Determination of the effects of the LHR on prophage induction in different STEC strains. Theconstruction of the pR=::rfp::chl reporter system has been described previously (32). To measure the effectof the LHR on the induction of Shiga toxin prophages with different promoters, the tetracyclineresistance gene in plasmids pLHR and pRK767 was replaced with the kanamycin resistance genederived from pKD4 (55) as described above, and the resulting plasmids were electroporated into E.coli FUA1303, E. coli FUA1311, and E. coli FUA1399. Strains were additionally transformed withpR=::rfp::chl plasmids to obtain transformants where red fluorescent protein (RFP) expression iscontrolled by the native phage promoter that also controls the expression of the chromosomallyencoded prophage, or to obtain RFP expression by alternate phage promoters that are not encodedon the chromosome of the host (Tables 3 and 4). The expression of rfp was measured by flowcytometry as described elsewhere (32).
Detection of the uspC-IS30-flhDC marker and virulence genes in E. coli wastewater isolates anddetection of the LHR in STEC. PCR screening of 102 STEC strains for the three LHR fragments was
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performed using multiplex PCR with the same primers and protocol that were used for the wastewaterand groundwater strains (Table 2).
Seventy E. coli wastewater isolates were screened for the presence of the uspC-IS30-flhDC marker (9,50), as an indicator for a naturalized global lineage of wastewater isolates, and for five virulence genesassociated with UPEC (papC, iroN, fyuA, ibeA, and sfa/foc). The genomic DNA of E. coli strains wasextracted from bacterial cultures by using a DNeasy blood and tissue kit (Qiagen, Hilden, Germany)according to the manufacturer’s instructions. Lists of genes and primers are provided in Table 2.High-resolution melting (HRM) analysis-qPCR was conducted on a Rotor-Gene Q (Qiagen) system usinga Type-it HRM PCR kit (Qiagen) (58) to detect the target genes listed in Table 2 with group-specificprimers according to the manufacturer’s protocols. Based on the primer annealing temperature (TA),three pairs of primers with the same TA but different melting temperatures (Tm) of the amplicons werecombined in the multiplex PCRs. The melting temperatures of PCR products are presented in Table 2. ThePCR was optimized with the following conditions: initial denaturation at 95°C for 5 min, followed by 45cycles of denaturation at 95°C for 10 s, annealing at the TA for 30 s, and extension at 72°C for 30 s. Duringthe HRM analysis stage, the temperature was increased from 65°C to 95°C at the speed of 0.1°C per stepand was held for 2 s at each step. Results are presented as means � standard deviations (SD) for threebiological replicates.
Statistical analysis. Data were obtained in 10 biological replicates (for the oxidation of membranelipids) or 3 biological replicates (for all other assays) and are expressed as means � standard deviations.Data were analyzed by one-way analysis of variance (ANOVA) using SPSS software, version 21.0 (SPSS Inc.,Chicago, IL, USA). The least significant difference (LSD) was used to test the difference among meansusing a P value of �0.05.
SUPPLEMENTAL MATERIALSupplemental material is available online only.SUPPLEMENTAL FILE 1, PDF file, 0.9 MB.
ACKNOWLEDGMENTSWe are grateful to Lars Leichert for providing plasmids with the roGFP2-based
probes.We acknowledge Canada Research Chairs, Alberta Agriculture and Forestry, the
Natural Sciences and Engineering Research Council of Canada, Alberta Innovates, andthe China Scholarship Council for funding.
REFERENCES1. Croxen MA, Law RJ, Scholz R, Keeney KM, Wlodarska M, Finlay BB. 2013.
2. Sarowska J, Futoma-Koloch B, Jama-Kmiecik A, Frej-Madrzak M, Ksiazc-zyk M, Bugla-Ploskonska G, Choroszy-Krol I. 2019. Virulence factors,prevalence and potential transmission of extraintestinal pathogenicEscherichia coli isolated from different sources: recent reports. Gut Pat-hog 11:10. https://doi.org/10.1186/s13099-019-0290-0.
3. Odonkor ST, Ampofo JK. 2013. Escherichia coli as an indicator of bacte-riological quality of water: an overview. Microbiol Res (Pavia) 4:2. https://doi.org/10.4081/mr.2013.e2.
4. Beuchat LR. 2002. Ecological factors influencing survival and growth ofhuman pathogens on raw fruits and vegetables. Microbes Infect4:413– 423. https://doi.org/10.1016/s1286-4579(02)01555-1.
5. Ridgway HF, Olson BH. 1982. Chlorine resistance patterns of bacteriafrom two drinking water distribution systems. Appl Environ Microbiol44:972–987.
6. Macauley JJ, Qiang Z, Adams CD, Surampalli R, Mormile MR. 2006.Disinfection of swine wastewater using chlorine, ultraviolet light andozone. Water Res 40:2017–2026. https://doi.org/10.1016/j.watres.2006.03.021.
7. Harrison JE, Schultz J. 1976. Studies on the chlorinating activity ofmyeloperoxidase. J Biol Chem 251:1371–1374.
8. Bommer A, Böhler O, Johannsen E, Dobrindt U, Kuczius T. 2018. Effect ofchlorine on cultivability of Shiga toxin producing Escherichia coli (STEC)and �-lactamase genes carrying E. coli and Pseudomonas aeruginosa. IntJ Med Microbiol 308:1105–1112. https://doi.org/10.1016/j.ijmm.2018.09.004.
9. Zhi S, Banting G, Li Q, Edge TA, Topp E, Sokurenko M, Scott C, Braith-waite S, Ruecker NJ, Yasui Y, McAllister T, Chui L, Neumann NF. 2016.Evidence of naturalized stress-tolerant strains of Escherichia coli in mu-nicipal wastewater treatment plants. Appl Environ Microbiol 82:5505–5518. https://doi.org/10.1128/AEM.00143-16.
10. Drazic A, Tsoutsoulopoulos A, Peschek J, Gundlach J, Krause M, Bach NC,Gebendorfer KM, Winter J. 2013. Role of cysteines in the stability andDNA-binding activity of the hypochlorite-specific transcription factor HypT.PLoS One 8:e75683. https://doi.org/10.1371/journal.pone.0075683.
11. Parker BW, Schwessinger EA, Jakob U, Gray MJ. 2013. RclR is a reactivechlorine-specific transcription factor in Escherichia coli. J Biol Chem288:32574 –32584. https://doi.org/10.1074/jbc.M113.503516.
12. Gray MJ, Wholey W-Y, Parker BW, Kim M, Jakob U. 2013. NemR is ableach-sensing transcription factor. J Biol Chem 288:13789 –13798.https://doi.org/10.1074/jbc.M113.454421.
13. Du Z, Nandakumar R, Nickerson KW, Li X. 2015. Proteomic adaptations tostarvation prepare Escherichia coli for disinfection tolerance. Water Res69:110 –119. https://doi.org/10.1016/j.watres.2014.11.016.
14. Cabiscol E, Tamarit J, Ros J. 2000. Oxidative stress in bacteria and proteindamage by reactive oxygen species. Int Microbiol 3:3– 8.
15. Hillion M, Antelmann H. 2015. Thiol-based redox switches in prokaryotes.Biol Chem 396:415–444. https://doi.org/10.1515/hsz-2015-0102.
16. Winter J, Linke K, Jatzek A, Jakob U. 2005. Severe oxidative stress causesinactivation of DnaK and activation of the redox-regulated chaperoneHsp33. Mol Cell 17:381–392. https://doi.org/10.1016/j.molcel.2004.12.027.
18. Mercer RG, Nguyen O, Ou Q, McMullen L, Gänzle MG. 2017. Functionalanalysis of genes encoded by the locus of heat resistance in Escherichiacoli. Appl Environ Microbiol 83:e1400-17. https://doi.org/10.1128/AEM.01400-17.
19. Heiman KE, Mody RK, Johnson SD, Griffin PM, Gould LH. 2015. Escherichiacoli O157 outbreaks in the United States, 2003–2012. Emerg Infect Dis21:1293–1301. https://doi.org/10.3201/eid2108.141364.
20. Beer KD, Gargano JW, Roberts VA, Hill VR, Garrison LE, Kutty PK, HilbornED, Wade TJ, Fullerton KE, Yoder JS. 2015. Surveillance for waterbornedisease outbreaks associated with drinking water—United States,
Wang et al. Applied and Environmental Microbiology
February 2020 Volume 86 Issue 4 e02123-19 aem.asm.org 14
21. Kintz E, Brainard J, Hooper L, Hunter P. 2017. Transmission pathways forsporadic Shiga-toxin producing E. coli infections: a systematic reviewand meta-analysis. Int J Hyg Environ Health 220:57– 67. https://doi.org/10.1016/j.ijheh.2016.10.011.
22. Teunis PFM, Ogden ID, Strachan N. 2008. Hierarchical dose responseof E. coli O157:H7 from human outbreaks incorporating heterogene-ity in exposure. Epidemiol Infect 136:761–770. https://doi.org/10.1017/S0950268807008771.
23. Foxman B. 2010. The epidemiology of urinary tract infection. Nat RevUrol 7:653– 660. https://doi.org/10.1038/nrurol.2010.190.
24. Adefisoye MA, Okoh AI. 2016. Identification and antimicrobial resistanceprevalence of pathogenic Escherichia coli strains from treated wastewa-ter effluents in Eastern Cape, South Africa. Microbiologyopen 5:143–151.https://doi.org/10.1002/mbo3.319.
25. Anastasi EM, Matthews B, Gundogdu A, Vollmerhausen TL, Ramos NL,Stratton H, Ahmed W, Katouli M. 2010. Prevalence and persistence ofEscherichia coli strains with uropathogenic virulence characteristics insewage treatment plants. Appl Environ Microbiol 76:5882–5886. https://doi.org/10.1128/AEM.00141-10.
26. Boczek LA, Rice EW, Johnston B, Johnson JR. 2007. Occurrence ofantibiotic-resistant uropathogenic Escherichia coli clonal group A inwastewater effluents. Appl Environ Microbiol 73:4180 – 4184. https://doi.org/10.1128/AEM.02225-06.
27. Degrossoli A, Müller A, Xie K, Schneider JF, Bader V, Winklhofer KF, MeyerAJ, Leichert LI. 2018. Neutrophil-generated HOCl leads to non-specificthiol oxidation in phagocytized bacteria. Elife 7:e32288. https://doi.org/10.7554/eLife.32288.
28. Fang Y, McMullen LM, Gänzle MG. 2020. Effect of drying on oxidation ofmembrane lipids and expression of genes encoded by the Shiga toxinprophage in Escherichia coli. Food Microbiol 86:103332. https://doi.org/10.1016/j.fm.2019.103332.
29. Fang Y, Mercer RG, McMullen LM, Gänzle MG. 2017. Induction of Shigatoxin-encoding prophage by abiotic environmental stress in food. ApplEnviron Microbiol 83:e01378-17. https://doi.org/10.1128/AEM.01378-17.
30. Ma A, Chui L. 2017. Identification of heat resistant Escherichia coli byqPCR for the locus of heat resistance. J Microbiol Methods 133:87– 89.https://doi.org/10.1016/j.mimet.2016.12.019.
31. Gill A, Tamber S, Yang X. 2019. Relative response of populations ofEscherichia coli and Salmonella enterica to exposure to thermal, alkalineand acidic treatments. Int J Food Microbiol 293:94 –101. https://doi.org/10.1016/j.ijfoodmicro.2019.01.007.
32. Zhang L, Simpson D, McMullen L, Gänzle M. 2018. Comparative genom-ics and characterization of the late promoter pR= from Shiga toxinprophages in Escherichia coli. Viruses 10:595. https://doi.org/10.3390/v10110595.
33. Spurbeck RR, Dinh PC, Jr, Walk ST, Stapleton AE, Hooton TM, Nolan LK,Kim KS, Johnson JR, Mobley HL. 2012. Escherichia coli isolates that carryvat, fyuA, chuA, and yfcV efficiently colonize the urinary tract. InfectImmun 80:4115– 4122. https://doi.org/10.1128/IAI.00752-12.
34. Naguib Y. 1998. A fluorometric method for measurement of peroxylradical scavenging activities of lipophilic antioxidants. Anal Biochem265:290 –298. https://doi.org/10.1006/abio.1998.2931.
36. Stevens JF, Maier CS. 2008. Acrolein: sources, metabolism, and biomo-lecular interactions relevant to human health and disease. Mol NutrFood Res 52:7–25. https://doi.org/10.1002/mnfr.200700412.
37. Kojima M. 1971. Studies on the effects of isothiocyanates and theiranalogues on microorganisms. I. Effect of isothiocyanates on the oxygenuptake of yeasts. J Ferment Technol 49:740 –746.
38. Okochi M, Yokokawa H, Lim T-K, Taguchi T, Takahashi H, Yokouchi H,Kaiho T, Sakuma A, Matsunaga T. 2005. Disinfection of microorganismsby use of electrochemically regenerated periodate. Appl Environ Micro-biol 71:6410 – 6413. https://doi.org/10.1128/AEM.71.10.6410-6413.2005.
39. Lee C, Franke KB, Kamal SM, Kim H, Lünsdorf H, Jäger J, Nimtz M, TrcekJ, Jänsch L, Bukau B, Mogk A, Römling U. 2018. Stand-alone ClpGdisaggregase confers superior heat tolerance to bacteria. Proc Natl AcadSci U S A 115:E273–E282. https://doi.org/10.1073/pnas.1712051115.
40. Holmgren A. 1979. Thioredoxin catalyzes the reduction of insulin disulfidesby dithiothreitol and dihydrolipoamide. J Biol Chem 254:9627–9632.
41. Ferguson GP, Munro AW, Douglas RM, McLaggan D, Booth IR. 1993.Activation of potassium channels during metabolite detoxification inEscherichia coli. Mol Microbiol 9:1297–1303. https://doi.org/10.1111/j.1365-2958.1993.tb01259.x.
42. Skorko-Glonek J, Zurawa D, Kuczwara E, Wozniak M, Wypych Z, LipinskaB. 1999. The Escherichia coli heat shock protease HtrA participates indefense against oxidative stress. Mol Gen Genet 262:342–350. https://doi.org/10.1007/s004380051092.
43. Mates AK, Sayed AK, Foster JW. 2007. Products of the Escherichia coli acidfitness island attenuate metabolite stress at extremely low pH andmediate a cell density-dependent acid resistance. J Bacteriol 189:2759 –2768. https://doi.org/10.1128/JB.01490-06.
44. Lee C, Wigren E, Trcek J, Peters V, Kim J, Hasni MS, Nimtz M, Lindqvist Y,Park C, Curth U, Lünsdorf H, Römling U. 2015. A novel protein qualitycontrol mechanism contributes to heat shock resistance of worldwide�distributed Pseudomonas aeruginosa clone C strains. Environ Microbiol17:4511– 4526. https://doi.org/10.1111/1462-2920.12915.
45. Boll EJ, Marti R, Hasman H, Overballe-Petersen S, Stegger M, Ng K,Knøchel S, Krogfelt KA, Hummerjohann J, Struve C. 2017. Turn up theheat—food and clinical Escherichia coli isolates feature two transferrableloci of heat resistance. Front Microbiol 8:579. https://doi.org/10.3389/fmicb.2017.00579.
46. Mercer RG, Walker BD, Yang X, McMullen LM, Gänzle MG. 2017. The locusof heat resistance (LHR) mediates heat resistance in Salmonella enterica,Escherichia coli and Enterobacter cloacae. Food Microbiol 64:96 –103.https://doi.org/10.1016/j.fm.2016.12.018.
47. Los JM, Los M, Wegrzyn A, Wegrzyn G. 2012. Altruism of Shiga toxin-producing Escherichia coli: recent hypothesis versus experimental re-sults. Front Cell Infect Microbiol 2:166. https://doi.org/10.3389/fcimb.2012.00166.
48. Imamovic L, Jofre J, Schmidt H, Serra-Moreno R, Muniesa M. 2009.Phage-mediated Shiga toxin 2 gene transfer in food and water. ApplEnviron Microbiol 75:1764 –1768. https://doi.org/10.1128/AEM.02273-08.
49. Łos JM, Łos M, Wegrzyn G, Wegrzyn A. 2009. Differential efficiency ofinduction of various lambdoid prophages responsible for production ofShiga toxins in response to different induction agents. Microb Pathog47:289 –298. https://doi.org/10.1016/j.micpath.2009.09.006.
50. Zhi S, Banting G, Stothard P, Ashbolt NJ, Checkley S, Meyer K, Otto S,Neumann NF. 2019. Evidence for the evolution, clonal expansion andglobal dissemination of water treatment-resistant naturalized strains ofEscherichia coli in wastewater. Water Res 156:208 –222. https://doi.org/10.1016/j.watres.2019.03.024.
51. Anastasi EM, Wohlsen TD, Stratton HM, Katouli M. 2013. Survival ofEscherichia coli in two sewage treatment plants using UV irradiation andchlorination for disinfection. Water Res 47:6670 – 6679. https://doi.org/10.1016/j.watres.2013.09.008.
52. Xi C, Zhang Y, Marrs CF, Ye W, Simon C, Foxman B, Nriagu J. 2009.Prevalence of antibiotic resistance in drinking water treatment anddistribution systems. Appl Environ Microbiol 75:5714 –5718. https://doi.org/10.1128/AEM.00382-09.
53. Rijavec M, Starcic Erjavec M, Ambrozic Avgustin J, Reissbrodt R, Fruth A,Krizan-Hergouth V, Žgur-Bertok D. 2006. High prevalence of multidrugresistance and random distribution of mobile genetic elements amonguropathogenic Escherichia coli (UPEC) of the four major phylogeneticgroups. Curr Microbiol 53:158 –162. https://doi.org/10.1007/s00284-005-0501-4.
54. Gänzle MG, Hertel C, Hammes WP. 1999. Resistance of Escherichia coliand Salmonella against nisin and curvacin A. Int J Food Microbiol48:37–50. https://doi.org/10.1016/s0168-1605(99)00026-4.
55. Datsenko KA, Wanner BL. 2000. One-step inactivation of chromosomalgenes in Escherichia coli K-12 using PCR products. Proc Natl Acad SciU S A 97:6640 – 6645. https://doi.org/10.1073/pnas.120163297.
56. Reference deleted.57. Pfaffl MW. 2001. A new mathematical model for relative quantification in
real-time RT–PCR. Nucleic Acids Res 29:e45. https://doi.org/10.1093/nar/29.9.e45.
58. Lin XB, Gänzle MG. 2014. Quantitative high-resolution melting PCRanalysis for monitoring of fermentation microbiota in sourdough. Int JFood Microbiol 186:42– 48. https://doi.org/10.1016/j.ijfoodmicro.2014.06.010.
59. Siedler S, Schendzielorz G, Binder S, Eggeling L, Bringer S, Bott M.2014. SoxR as a single-cell biosensor for NADPH-consuming enzymesin Escherichia coli. ACS Synth Biol 3:41– 47. https://doi.org/10.1021/sb400110j.
Chlorine Resistance in Escherichia coli Applied and Environmental Microbiology
February 2020 Volume 86 Issue 4 e02123-19 aem.asm.org 15
60. Cornelis P, Wei Q, Andrews SC, Vinckx T. 2011. Iron homeostasis andmanagement of oxidative stress response in bacteria. Metallomics3:540 –549. https://doi.org/10.1039/c1mt00022e.
61. Gray MJ, Jakob U. 2015. Oxidative stress protection by polyphosphate—new roles for an old player. Curr Opin Microbiol 24:1– 6. https://doi.org/10.1016/j.mib.2014.12.004.
62. Müller A, Langklotz S, Lupilova N, Kuhlmann K, Bandow JE, Leichert L.
2014. Activation of RidA chaperone function by N-chlorination. NatCommun 5:5804. https://doi.org/10.1038/ncomms6804.
63. White AP, Sibley KA, Sibley CD, Wasmuth JD, Schaefer R, Surette MG,Edge TA, Neumann NF. 2011. Intergenic sequence comparison ofEscherichia coli isolates reveals lifestyle adaptations but not hostspecificity. Appl Environ Microbiol 77:7620 –7632. https://doi.org/10.1128/AEM.05909-11.
Wang et al. Applied and Environmental Microbiology
February 2020 Volume 86 Issue 4 e02123-19 aem.asm.org 16
