- Record: found
- Abstract: found
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Identification and antimicrobial resistance patterns of bacterial enteropathogens from children aged 0–59 months at the University Teaching Hospital, Lusaka, Zambia: a prospective cross sectional study
research-article
Author(s):
Harriet Chiyangi
1
,
2 ,
John B. Muma
3 ,
Sydney Malama
4
,
,
Joel Manyahi
4 ,
Ahmed Abade
2 ,
Geoffrey Kwenda
5 ,
Mecky I. Matee
6
Publication date (Electronic):
2 February 2017
Journal:
BMC Infectious Diseases
Publisher:
BioMed Central
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Abstract
Background
Bacterial diarrhoeal disease is among the most common causes of mortality and morbidity in children 0–59 months at the University Teaching Hospital in Lusaka, Zambia. However, most cases are treated empirically without the knowledge of aetiological agents or antimicrobial susceptibility patterns. The aim of this study was, therefore, to identify bacterial causes of diarrhoea and determine their antimicrobial susceptibility patterns in stool specimens obtained from the children at the hospital.
Methods
This hospital-based cross-sectional study involved children aged 0–59 months presenting with diarrhoea at paediatrics wards at the University Teaching Hospital in Lusaka, Zambia, from January to May 2016. Stool samples were cultured on standard media for enteropathogenic bacteria, and identified further by biochemical tests. Multiplex polymerase chain reaction was used for characterization of diarrhoeagenic Escherichia coli strains. Antimicrobial susceptibility testing was performed on antibiotics that are commonly prescribed at the hospital using the Kirby-Bauer disc diffusion method, which was performed using the Clinical Laboratory Standards International guidelines.
Results
Of the 271 stool samples analysed Vibrio cholerae 01 subtype and Ogawa serotype was the most commonly detected pathogen (40.8%), followed by Salmonella species (25.5%), diarrhoeagenic Escherichia coli (18%), Shigella species (14.4%) and Campylobacter species (3.5%). The majority of the bacterial pathogens were resistant to two or more drugs tested, with ampicillin and co-trimoxazole being the most ineffective drugs. All diarrhoeagenic Escherichia coli isolates were extended spectrum β-lactamase producers.
Conclusion
Five different groups of bacterial pathogens were isolated from the stool specimens, and the majority of these organisms were multidrug resistant. These data calls for urgent revision of the current empiric treatment of diarrhoea in children using ampicillin and co-trimoxazole, and emphasizes the need for continuous antimicrobial surveillance as well as the implementation of prevention programmes for childhood diarrhoea.
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Most cited references29
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Global Causes of Diarrheal Disease Mortality in Children <5 Years of Age: A Systematic Review
Estimation of pathogen-specific causes of child diarrhea deaths is needed to guide vaccine development and other prevention strategies. We did a systematic review of articles published between 1990 and 2011 reporting at least one of 13 pathogens in children <5 years of age hospitalized with diarrhea. We included 2011 rotavirus data from the Rotavirus Surveillance Network coordinated by WHO. We excluded studies conducted during diarrhea outbreaks that did not discriminate between inpatient and outpatient cases, reporting nosocomial infections, those conducted in special populations, not done with adequate methods, and rotavirus studies in countries where the rotavirus vaccine was used. Age-adjusted median proportions for each pathogen were calculated and applied to 712 000 deaths due to diarrhea in children under 5 years for 2011, assuming that those observed among children hospitalized for diarrhea represent those causing child diarrhea deaths. 163 articles and WHO studies done in 31 countries were selected representing 286 inpatient studies. Studies seeking only one pathogen found higher proportions for some pathogens than studies seeking multiple pathogens (e.g. 39% rotavirus in 180 single-pathogen studies vs. 20% in 24 studies with 5–13 pathogens, p<0·0001). The percentage of episodes for which no pathogen could be identified was estimated to be 34%; the total of all age-adjusted percentages for pathogens and no-pathogen cases was 138%. Adjusting all proportions, including unknowns, to add to 100%, we estimated that rotavirus caused 197 000 [Uncertainty range (UR) 110 000–295 000], enteropathogenic E. coli 79 000 (UR 31 000–146 000), calicivirus 71 000 (UR 39 000–113 000), and enterotoxigenic E. coli 42 000 (UR 20 000–76 000) deaths. Rotavirus, calicivirus, enteropathogenic and enterotoxigenic E. coli cause more than half of all diarrheal deaths in children <5 years in the world.
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Emergence of a Globally Dominant IncHI1 Plasmid Type Associated with Multiple Drug Resistant Typhoid
Kathryn E. Holt, Minh Duy Phan, Stephen Baker … (2011)
Introduction Typhoid fever remains a serious public health problem in many developing countries, with highest incidence in parts of Asia (274 per 100,000 person-years) and Africa (50 per 100,000 person-years) [1], [2]. The causative agent is the bacterium Salmonella enterica serovar Typhi (S. Typhi). While vaccines against S. Typhi exist, it is mainly restricted groups such as travellers [3], [4] and individuals enrolled in large vaccine trials [5] who are immunized, and antimicrobial treatment remains central to the control of typhoid fever [3]. However antimicrobial resistant typhoid has been observed for over half a century and is now common in many areas. Chloramphenicol resistant S. Typhi was first reported in 1950, shortly after the drug was introduced for treatment of typhoid [6]. By the early 1970s, S. Typhi resistant to both chloramphenicol and ampicillin had been observed [7] and multidrug resistant (MDR) S. Typhi (defined here as resistance to chloramphenicol, ampicillin and trimethoprim-sulfamethoxazole) emerged soon after [8]. The rate of MDR among S. Typhi can fluctuate over time and geographical space, as can the precise combination of drug resistance genes and phenotypes [9], [10]. However in many typhoid endemic areas, an increasing prevalence of MDR S. Typhi was observed in the late 1990s [11], [12], [13], and MDR typhoid now predominates in many areas [9], [14] including parts of Asia [15], [16], Africa [17] and the Middle East [18], [19], [20], [21]. MDR S. Typhi with reduced susceptibility to fluoroquinolones are increasingly common [9], [15], [16], [22], leaving macrolides or third generation cephalosporins as the only options for therapy [23], [24]. In S. Typhi the MDR phenotype is almost exclusively conferred by self-transmissible plasmids of the HI1 incompatibility type (IncHI1) [8], [11], [25], [26], [27], [28], [29], [30], although other plasmids are occasionally reported [31]. In the laboratory, IncHI1 plasmids can transfer between Enterobacteriaceae and other Gram-negative bacteria [32] and in nature, IncHI1 plasmids have been detected in pathogenic isolates of Salmonella enterica and Escherichia coli [33], [34], [35], [36]. However it remains unclear whether the increase in MDR typhoid is due to the exchange of resistance genes among co-circulating S. Typhi or to the expansion of MDR S. Typhi clones. Efforts have been made to investigate variability within IncHI1 plasmids [29], [33], [37] or their S. Typhi hosts [22], [38], [39], [40], [41] but little progress has been made in linking the two together to answer fundamental questions of how MDR typhoid spreads. We recently developed a plasmid multi-locus sequence typing (PMLST) scheme for IncHI1 plasmids, which identified eight distinct IncHI1 plasmid sequence types (PSTs) among S. Typhi and S. Paratyphi A isolates, including five PSTs found in S. Typhi [37]. This pattern was not consistent with a single acquisition of an IncHI1 plasmid in S. Typhi followed by divergence into multiple plasmid lineages, rather it indicated that divergent IncHI1 plasmids have entered the S. Typhi population on multiple occasions [37]. However the phylogenetic relatedness of the S. Typhi hosts was not determined, thus we were unable to estimate how many times plasmids may have been independently acquired. In this study, we aimed to investigate the relative contribution of plasmid transfer, as opposed to the expansion of plasmid-bearing S. Typhi clones, to the emergence of MDR typhoid. We found evidence for plasmid transfer in older S. Typhi. However the vast majority of recent MDR typhoid was attributable to a single host-plasmid combination (S. Typhi H58-IncHI1 plasmid ST6). We performed further experiments to investigate possible mechanisms for the success of this host-plasmid combination, and identified a transposon in PST6 that confers tolerance to high osmolarity. Materials and Methods Bacterial isolates and DNA extraction The bacterial isolates analyzed by SNP assay are summarized in Table 1 and listed in full in Table S1. DNA was extracted using Wizard Genomic DNA purification kits (Promega) according to manufacturer's instructions. Details of the isolates used for competition experiments are also listed in Table S1. 10.1371/journal.pntd.0001245.t001 Table 1 Summary of 454 S. Typhi isolates analyzed in this study. Region No. countries pre-1970s 1970s–1980s 1990s 2000–2007 Total isolates South & Central America 4 0 6 3 2 11 Central, Southern, East Africa 7 10 3 3 26 42 North Africa 3 11 1 8 5 25 West Africa 11 28 0 6 12 46 East Asia 8 5 8 22 187 222 Indian Subcontinent 3 0 3 1 66 70 Middle East 3 0 0 0 31 31 Europe 5 1 1 2 2 6 Unknown - 1 0 0 0 1 Total 44 56 22 45 331 454 BRD948 is an attenuated Ty2-derived strain (also known as CVD908-htrA), which has deletion mutations in aroC (t0480), aroD (t1231), and htrA (t0210) [42]. The growth of BRD948 on LB agar or in LB broth was enabled by supplementation with aromatic amino acid mix (aro mix) to achieve the final concentration of 50 µM L-phenylalanine, 50 µM L-tryptophan, 1 µM para-aminobenzoic acid and 1 µM 2,3-dihydroxybenzoic acid. Identification and phylogenetic analysis of IncHI1 SNPs Plasmid sequences were downloaded from the European Nucleotide Archive (plasmid details and accessions in Table 2). SNPs between finished plasmid sequences were identified using the nucmer and show-snps algorithms within the MUMmer 3.1 package [43], via pairwise comparisons with pAKU_1. To identify SNPs in S. Typhi PST6 IncHI1 plasmids, 36 bp single-ended Illumina/Solexa sequencing reads from S. Typhi isolates E03-9804, ISP-03-07467 and ISP-04-06979 were aligned to the pAKU_1 sequence using Maq [44] and quality filters as described previously [45]. SNPs called in repetitive regions or inserted sequences were excluded from phylogenetic analysis, so that phylogenetic trees were based only on the conserved IncHI1 core regions. This resulted in a total of 347 SNPs, which were analyzed using BEAST [46] to simultaneously infer a phylogenetic tree and divergence dates (using the year of isolation of each plasmid as listed in Table 1, resulting tree in Figure 1). Parameters used were as follows: generalised time reversible model with a Gamma model of site heterogeneity (4 gamma categories); a relaxed molecular clock with uncorrelated exponential rates [46], a coalescent tree prior estimated using a Bayesian skyline model with 10 groups [47], default priors and 20 million iterations. 10.1371/journal.pntd.0001245.g001 Figure 1 Phylogenetic tree for IncHI1 plasmid sequences. Phylogenetic tree based on 347 SNPs identified among 8 publicly available IncHI1 plasmid sequences (Table 2), constructed using BEAST (with 20 million iterations, 4 replicate runs, exponential clock model). Terminal nodes are labelled with the organism of origin (STy = Salmonella enterica serovar Typhi, SCh = Salmonella enterica serovar Choleraesuis, STm = Salmonella enterica serovar Typhimurium, SPa = Salmonella enterica serovar Paratyphi A, Ec = E. coli O111:H-) and date of isolation. Isolation dates were input into the BEAST model in order to estimate divergence dates for internal nodes (open circles, labelled with divergence date estimates; brackets indicate 95% highest posterior density interval). Insertion sites (grey) are based on sequence data and verified (except for pO111_1 and pMAK1) by PCR. Precise insertion sites and PCR primers for verification are given in Tables 3 & 4. Four major plasmid groups, PST1, PST5, PST6, PST7, are coloured as labelled. 10.1371/journal.pntd.0001245.t002 Table 2 IncHI1 plasmid sequences analyzed in this study. Plasmid Host Year of isolation Plasmid type Accession Citation pHCM1 S. Typhi strain CT18 1993 PST1 AL513383 [54] pAKU_1 S. Paratyphi A strain AKU_12601 2003 PST7 AM412236 [33] R27 S. Typhimurium 1961 PST5 AF250878 [65] pMAK1 S. Choleraesuis strain L-2454 2002 PST1 AB366440 - pO111_1 E. coli O111:H- strain 11128 2001 PST1 AP010961 [66] p9804_1 S. Typhi strain E03-9804 2004 PST6 ERA000001 [45] p7467_1 S. Typhi strain ISP-03-07467 2003 PST6 ERA000001 [45] p6979_1 S. Typhi strain ISP-04-06979 2004 PST6 ERA000001 [45] SNP typing analysis The chromosomal haplotype of S. Typhi isolates was determined based on the SNPs present at 1,485 chromosomal loci identified previously from genome-wide surveys [41], [45] and listed in [22], [39]. IncHI1 plasmid haplotypes were determined using 231 SNPs located in the conserved IncHI1 backbone sequence, listed in Table S2 (note these do not include SNPs specific to pMAK1 or pO111_1 which were not available at the time of assay design, nor any SNPs falling within 10 bp of each other as these cannot be accurately targeted via GoldenGate assay; however additional SNPs identified via plasmid MLST [37] were included, see Table S2). Resistance gene sequences were interrogated using additional oligonucleotide probes, listed in [16]. All loci were interrogated using a GoldenGate (Illumina) custom assay according to the manufacturer's standard protocols, as described previously [16], [22], [39]. SNP calls were generated from raw fluorescence signal data by clustering with a modified version of Illuminus [48] as described previously [22]. The percentage of IncHI1 SNP loci yielding positive signals in the GoldenGate assay clearly divided isolates into two groups, indicating presence of an IncHI plasmid (signals for >90% of IncHI1 loci) or absence of such a plasmid (signals for 90% of IncHI1 target loci were detected, taken to imply presence of an IncHI1 plasmid (red), or 99% identical at the nucleotide level) that included the tra1 and tra2 regions encoding conjugal transfer [29], [33], [37], [52]. Subsequently, we identified 347 single nucleotide polymorphisms (SNPs) within these conserved regions, which were used to construct a phylogenetic tree of IncHI1 plasmids and to estimate the divergence dates of internal nodes of this tree based on the known isolation dates for each plasmid [53] (Figure 1). The tree topology is in general agreement with that inferred previously using a plasmid MLST approach [37]. The sequences of the three most recent S. Typhi plasmids (isolated 2003–2004) were very closely related and correspond to a previously defined plasmid sequence type (PST) known as PST6 [37] (Figure 1, red). According to our divergence date estimates, the most recent common ancestor (mrca) shared by these three plasmids existed circa 1999 (Figure 1). The PST6 plasmids were also closely related to the PST7 plasmid pAKU_1 from S. Paratyphi A (Figure 1, orange), with mrca circa 1992. The plasmids pHCM1, pO111_1 and pMAK1 formed a distinct group corresponding to PST1, with mrca circa 1989 (Figure 1, green). The eighth reference plasmid R27 (PST5) was quite distinct from the others, with an estimated divergence date of 1952 (Figure 1, black). In addition to the conserved IncHI1 core regions, the plasmids each harbour insertions of drug resistance elements. These include transposons Tn10 (encoding tetracycline resistance), Tn9 (encoding chloramphenicol resistance via the cat gene (SPAP0067)), strAB (SPAP0152-SPAP0153, SPAP0230-SPAP0231; encoding streptomycin resistance), sul1 and sul2 (SPAP0132 , SPAP0151; encoding sulfonamide resistance), dfrA7 (SPAP0133; encoding trimethoprim resistance) and bla TEM-1 (SPAP0143; encoding ampicillin resistance) [29], [33], [54]. The insertion sites of these elements, confirmed using PCR (Tables 3 & 4), differed between lineages of the IncHI1 phylogenetic tree (Figure 1, grey). All plasmid sequences included Tn10, however three different insertion sites were evident (Table 4), suggesting the transposon was acquired by IncHI1 plasmids on at least three separate occasions (Figure 1, grey). Tn9 was present in all plasmids other than R27, however the insertion site in PST6 and PST7 plasmids differed from that in PST1, suggesting at least two independent acquisitions. It was previously noted that pHCM1 (PST1) and pAKU_1 (PST7) share identical insertions into Tn9 of a sequence incorporating Tn21 (including sul1, dfrA7), bla TEM-1, sul2, and strAB [33]; here we found this insertion into Tn9 was conserved in all PST1 and PST6 plasmid sequences. Together, this composite set of drug resistance elements encodes MDR (resistance to chloramphenicol, ampicillin and trimethoprim-sulfamethoxazole). 10.1371/journal.pntd.0001245.t004 Table 4 Resistance gene insertion sites in IncHI1 plasmids inferred from a combination of PCR and sequencing. IncHI1 plasmid sequence type PST1 PST5 PST6 PST6 PST7 PST8 Plasmid or isolate pHCM1 pMAK1 pO111_1 R27 p6979 pSTY7 pAKU1 81918 81863, 81424 Bacterial host STy SCh Ec STm STy STy SPa STy STy Tn10 insertion B B B C A A A A A sequence data B B B C A n/a A n/a n/a N (Tn10 - HCM1.247) + n/d n/d - - - - - - O (tetD - SPAP0276) - n/d n/d - + + + + + P (SPAP0261 - Tn10) - n/d n/d - + + + + + Tn9 insertion B B B - A A A - A sequence data B B B - A n/d A n/a n/a J (cat - trhN) - n/d n/d - + + + - + K (mer - trhI) - n/d n/d - + + + - + M (cat - HCM1.203) + n/d n/d - - - - - - L (insA - tetA) + n/d n/d - - - - - - Tn21 into Tn9 + + + - + + + - + sequence data + + + - + n/d + n/a n/a H (tnpA - Tn9) +* n/d n/d - + + + - + I (merR - Tn9) +* n/d n/d - + + + - + bla/sul/str into Tn21 + + + - + + + - + sequence data + + + - + n/d + n/a n/a G (strB – tniAdelta) + + + - + n/d + - + strAB 2nd copy (SPAP0230- SPAP0231) - - - - - - + + + sequence data - - - - - n/d + n/a n/a Q (strB – SPAP0228) - n/d n/d - - - + + + Summaries of five insertion patterns are shown in bold italics; these are inferred from sequence data where available (italics) and PCR using primers shown in Table 3 (labelled G–Q). STy = Salmonella enterica serovar Typhi, SCh = Salmonella enterica serovar Choleraesuis, STm = Salmonella enterica serovar Typhimurium, SPa = Salmonella enterica serovar Paratyphi A, Ec = E. coli O111:H−. + positive PCR result (i.e. successful amplification); - negative PCR result (i.e. no amplification product detected); *distinct amplicon size for PST1; n/d PCR not done; n/a sequence data not available. “strAB 2nd” copy refers to the insertion of streptomycin resistance genes strAB directly into the plasmid backbone (SPAP0230-SPAP0231), not as part of the bla/sul/str element (SPAP0152-SPAP0153). 10.1371/journal.pntd.0001245.t005 Table 5 Chromosome, plasmid and resistance gene details of drug resistant S. Typhi isolated up to 1993*. Isolate Year Country Chr Plas IS1 cat tetA tetC tetD tetR Tn10LR tnpA merAPRT IntI1 sul1 dhfR dfrA7 bla IS26 sul2 strAB betU 76–54 1976 Chile H50 7654 y y y y y y 78–851 1978 Tunisia H9 78851 y y y y y y y y CT18 1993 Vietnam H1 PST1 y y y y y y y y y y y y y y 76–1406 1976 Indonesia H42 PST2 y y y y y y y y y 75–2507 1975 India H55 PST2 y 77–302 1977 India H55 PST2 y y y y y y y y y 77–303 1977 India H55 PST2 y y y y y y y y y 72–1907 1972 Vietnam H68 PST2 y y y y y y y y y 72–1258 1972 Mexico H11 PST3 y y y y y y y y y y 73–1102 1973 Vietnam H87 PST4 y y y y y y y y 81–863 1981 Peru H50 PST8 y y y y y y y y y y y y 81–424 1981 Peru H77 PST8 y y y y y y y y y y y y y 81–918 1981 Peru H77 PST8 y y y y y y y 57Laos 2000* Laos H1 57Laos y y y y y y y y y y y y y y 03–4747 2003* Togo H42 PST2 y y y y y y y y y y 04–6845 2004* Benin H42 PST2 y y y y y y y Chr - S. Typhi chromosomal haplotype; Plas - IncHI1 plasmid sequence type; *- MDR S. Typhi isolated after 1993 that were not of the H58 haplotype or PST6 IncHI1 haplotype; y - gene detected in isolate. Dissecting the emergence of MDR typhoid In order to investigate the contribution of distinct IncHI1 plasmid types over time to the emergence of MDR S. Typhi, we performed high resolution SNP typing of S. Typhi chromosomal and IncHI1 plasmid loci in a global collection of 454 S. Typhi, isolated between 1958–2007 (Table 1, Table S1). These isolates include 19 S. Typhi isolates sequenced previously [45] and 22 S. Typhi isolated from Kenya in 2004–2007 [22]. We also typed eight IncHI1 S. Typhi plasmids harboured in E. coli transconjugants [29], [37]. SNP typing was performed using the GoldenGate (Illumina) platform to simultaneously assay chromosomal and plasmid SNP loci. We targeted 231 SNPs from the conserved region of the IncHI1 plasmid (Table S2, [37]; note 116 of the 347 identified SNPs were not able to be included in the GoldenGate assay, see Methods) and 119 from resistance genes and associated transposons (see [16]). Of the 454 S. Typhi that we typed, 193 (43%) harboured IncHI1 plasmids, which clustered into nine distinct haplotypes (Figure 3B). As expected, the majority of IncHI1 plasmids harboured multiple resistance genes or elements including Tn10, Tn9, strAB, sul1, sul2, dfrA7 and bla TEM-1. Transposon insertion sites were confirmed for representative plasmids using PCR (Table 4) and agree with the patterns of insertion sites determined by sequencing (Figure 1 & 3B). Thirteen IncHI1 plasmids were identified among S. Typhi isolated prior to 1994 (Table 5), including seven of the total nine distinct IncHI1 plasmid haplotypes (Figure 3B). 10.1371/journal.pntd.0001245.g003 Figure 3 Phylogenetic trees of S. Typhi chromosome and IncHI1 plasmid. (A) Phylogenetic tree indicating chromosomal haplotypes of 454 S. Typhi isolates determined by SNP typing with the GoldenGate assay. Circles correspond to detected S. Typhi haplotypes; node sizes are scaled to the number of isolates detected with that haplotype and labelled with this number. Unfilled circle indicates tree root; reference isolates used to define the S. Typhi SNPs are labelled with the isolate name. S. Typhi haplotypes in which IncHI1 plasmids were detected (N = 201) are coloured; black circles indicate no IncHI1 plasmids were found among S. Typhi of that haplotype; other colours indicate the presence of specific IncHI1 plasmid haplotypes corresponding to the colours in (B). Note that most of the coloured nodes also contain S. Typhi isolates with no plasmid, and the colours do not represent the proportion of isolates harbouring the various plasmid types. (B) Phylogenetic tree of IncHI1 plasmids determined by SNP typing with the GoldenGate assay (coloured leaf nodes); grey leaf nodes indicate the position of non-S. Typhi plasmids, as determined from plasmid sequence data listed in Table 2. A total of 26 distinct S. Typhi haplotypes were identified by typing of chromosomal SNPs; their phylogenetic relationships are shown in Figure 3A. The PST2 plasmid was detected in three S. Typhi haplotypes isolated in Asia between 1972 and 1977 (Table 5), consistent with repeated introduction of closely related IncHI1 plasmids into distinct S. Typhi hosts. Similarly, PST8 was present in two S. Typhi haplotypes from Peru in 1981 (Table 5) [55], consistent with transfer of the PST8 plasmid among multiple S. Typhi haplotypes co-circulating in Peru at this time. Significantly, from 1995 onwards, nearly all IncHI1 plasmids were type PST6 (180/184 plasmids, 98%). Remarkably, there was an exclusive relationship between PST6 plasmids and S. Typhi haplogroup H58, with all PST6 plasmids found in S. Typhi H58 hosts, and no S. Typhi H58 harbouring non-PST6 plasmids (although 35% of S. Typhi H58 were non-MDR and plasmid-free). This strongly suggests that the apparent rise in MDR typhoid since the mid-1990s [11], [12], [13] is due to the clonal expansion of H58 S. Typhi carrying the MDR PST6 plasmid. This is in contrast to the longer-term situation described above, which showed that in the years following the first emergence of MDR typhoid (1970s–1980s), MDR IncHI1 plasmids had transferred repeatedly into distinct co-circulating S. Typhi haplotypes. The clonal expansion of H58 S. Typhi has been documented previously [22], [41], however the role of the PST6 plasmid has not been investigated. Among our collection, the oldest S. Typhi H58 isolate dates back to 1995 and carries the PST6 plasmid. To ascertain whether the common ancestor of S. Typhi H58 might have carried the PST6 plasmid, the phylogenetic structure among our 293 S. Typhi H58 isolates was resolved using 45 of the assayed SNP loci that differentiate within the H58 haplogroup (Figure 4). These SNPs divided the isolates into 24 distinct H58 haplotypes, with the majority (N = 270) in 13 haplotypes (Figure 4). Most of the H58 haplotypes (N = 14), including the ancestral haplotype A, included isolates harbouring the PST6 plasmid (Figure 4). We have previously sequenced the genomes of 19 S. Typhi, including seven isolates from the H58 haplogroup [45], and observed the insertion of an IS1 transposase between protein coding sequences STY3618 and STY3619 within all sequenced H58 S. Typhi genomes. This transposase was identical at the nucleotide level to the IS1 sequences within Tn9 in IncHI1 plasmids pHCM1 and pAKU_1, and shared a common insertion site in all seven S. Typhi H58 chromosomes sequenced [45]. In the present study, our SNP assays included a probe targeting sequences within the IS1 gene (SPAP0007). Nearly all of the S. Typhi H58 isolates gave positive signals for this IS1 target (Figure 4; coloured or white), with the sole exception of six isolates belonging to the H58 ancestral haplotype A (Figure 4, grey), which also included three isolates that carried the PST6 plasmid and tested positive for IS1 (Figure 4, purple). This suggests that the PST6 plasmid was likely acquired by the most recent common ancestor of S. Typhi H58 (Figure 4, haplotype A), followed by transposition of IS1 into the S. Typhi chromosome prior to divergence into subtypes of H58. Thus the dominance of PST6 over other MDR IncHI1 plasmids (noted here and previously [37]) and the dominance of H58 over other S. Typhi haplotypes (noted here and previously [22], [41]) appears to be the result of a trans-continental clonal expansion of MDR S. Typhi H58 carrying the PST6 plasmid. 10.1371/journal.pntd.0001245.g004 Figure 4 Phylogenetic tree of the H58 haplogroup of S. Typhi. Dashed line indicates where this tree joins into the larger phylogenetic tree of S. Typhi (shown in Figure 3A). The two major H58 lineages are indicated by colour (blue, lineage I; red, lineage II; purple, common ancestor of both lineages). Nodes are labelled with isolate names (outer nodes representing sequenced isolates; see [45]), haplotype (H followed by number, as defined in [41]) or letters indicating nodes resolved by SNP typing. Node sizes indicate the relative frequency of each haplotype within the study collection of 269 H58 S. Typhi isolates, according to the scale provided. The proportion of isolates in each node carrying the PST6 plasmid and IS1 (solid colour), IS1 only (white) or neither (grey) is indicated by shading. Possible selective advantages of IncHI1 PST6 These results indicate that the recent global spread of MDR typhoid is attributable to the emergence of a single plasmid-host combination (H58-PST6). We were able to transfer the PST6 plasmid pSTY7 from S. Typhi to E. coli [29] and back to S. Typhi (data not shown), confirming that the PST6 plasmid retains the ability to transfer between bacteria via conjugation, yet we found no evidence of PST6 transfer in natural S. Typhi populations (above). This raises the question of why this particular plasmid-host association has been so successful and exclusive. To investigate whether PST6 could confer any selective advantage over other IncHI1 plasmids harbouring similar antimicrobial resistance genes, representative PST6 (pSTY7) and PST1 (pHCM1) IncHI1 plasmids from Vietnamese S. Typhi were introduced into a common S. Typhi BRD948 host, derived from S. Typhi Ty2 (haplotype H10). The PST1 plasmid pHCM1 was chosen for comparison since its complete sequence is available [54] and it was previously observed to be common in MDR S. Typhi in Vietnam in the early 1990s, just prior to the emergence of PST6 in S. Typhi in Vietnam and elsewhere [29]. BRD948 (pHCM1) grew to three times the number of cfu compared to BRD948 (pSTY7) after 4 days of mixed growth in LB broth (Figure 5, black). We therefore hypothesized that the advantage conferred by PST6 plasmids, if any, might be related to specific environmental conditions or to plasmid-host compatibility. To test the latter, we compared the growth of wildtype PST1-bearing S. Typhi H1 and PST6-bearing S. Typhi H58 isolated from typhoid patients in Vietnam and Pakistan and genotyped using the GoldenGate assay (listed in Table S1). The two PST6-bearing S. Typhi H58 isolates tested were both able to out compete the PST1-bearing H1 isolate, so that S. Typhi H1 was barely detectable after four days of competitive growth (Figure 5, red). However plasmid-free S. Typhi H58 isolates were also able to outcompete a plasmid-free S. Typhi H1 isolate (Figure 5, blue), thus we cannot confirm the plasmid plays a role in the competitive advantage of H58-PST6 S. Typhi over and above that of the H58 chromosomal haplotype. 10.1371/journal.pntd.0001245.g005 Figure 5 Competitive growth assays for S. Typhi H58 and H1 with and without IncHI1 plasmids. The dynamics of five competitive growth assays conducted over four days of sequential sub-culture. Black line indicates competition in a common host background (attenuated laboratory strain S. Typhi BRD948; haplotype H10); the proportion of PST1- and PST6-bearing bacteria at each time point was calculated by streaking an aliquot of the sample onto agar plates and testing random colonies using a PCR that differentiates PST1 and PST6. Coloured lines indicate competition between wildtype S. Typhi isolates as specified in the legend (see Table S1 for isolate names); the proportion of H58 and H1 chromosomes at each time point was calculated by quantifying the relative abundance of two alleles at a SNP locus that differs between H58 and H1 S. Typhi using quantitative PCR. For all assays, experiments were replicated at least three times; data points represent the mean proportion of culture corresponding to the isolate underlined in the legend; error bars show the standard deviation of this proportion. To screen for conditions under which PST6 plasmids confer an advantage compared to PST1 plasmids, we used Biolog phenotyping arrays to compare the growth of plasmid-free S. Typhi BRD948 to BRD948 (pHCM1) and BRD948 (pSTY7) under a wide variety of conditions including various pH levels and osmotic/ionic strengths, and a wide variety of antibiotics and chemicals [51]. As expected, both IncHI1 plasmids conferred enhanced growth in the presence of a wide range of antibiotics including amoxicillin, azlocillin, oxacillin, penicillin G, phenethicillin, chloramphenicol, streptomycin, gentamicin, tetracyclines and trimethoprim (Table S3). BRD948 (pHCM1) displayed some minor growth advantages in the presence of additional antimicrobials, however none of these reached clinically relevant levels (Table S3). The only conditions under which BRD948 (pSTY7) grew better than BRD948 and BRD948 (pHCM1) was under high osmotic stress (3-5% NaCl or 6% KCl) (Table S3). We confirmed this phenotype by inoculating each isolate into high salt concentration media (0.8 M NaCl LB broth, approx. 4.7% NaCl); only the PST6-bearing isolate BRD948 (pSTY7) was able to grow under these conditions (Figure 6, red and grey). 10.1371/journal.pntd.0001245.g006 Figure 6 The effect of Tn6062 on osmotolerance in S. Typhi BRD948. Growth curves for S. Typhi isolates in 0.8 M NaCl LB broth. Error bars indicate range of maximum and minimum values. We hypothesised that the osmotolerant properties of PST6 plasmids may be explained by the presence of two putative transporters encoded within a composite transposon, Tn6062 (SPAP0100, SPAP0105, SPAP0106, SPAP0110; this transposon was referred to as Ins1056 in [37]). Tn6062 was present in all PST6 plasmids, the novel subtype of PST1 (57Laos) and two of the three PST8 plasmids, but absent from all other isolates (detected via two Tn6062-specific probes included in our SNP typing assay). To determine if Tn6062 was responsible for the osmotolerant phenotype of BRD948 (pSTY7), the two putative transporter genes from Tn6062 (SPAP0105 and SPAP0106) were inserted into the plasmid vector pAYCY184 and we assessed their effect on S. Typhi BRD948 in high salt concentration medium (0.8 M NaCl LB broth, approx. 4.7% NaCl). BRD948 (pAYCY184-Tn6062) was able to grow at a slightly lower rate than BRD948 (pSTY7) (Figure 6, blue), while BRD948 carrying the empty pAYCY184 vector was unable to grow (Figure 6, black). Therefore the transposon Tn6062 carried by the PST6 IncHI1 plasmids confers an osmotolerant phenotype on its S. Typhi host. Discussion Our analysis of IncHI1 plasmid sequences indicates that plasmids responsible for the MDR phenotype in S. Typhi are closely related to those associated with MDR in other enteric pathogens including S. Paratyphi A, S. Choleraesuis and enterohaemorrhagic E. coli O111:H- (Figure 1, Table 2). These plasmids share a recent common ancestor approximately six decades old and have evolved into several distinct lineages via accumulation of point mutations, followed by acquisition of resistance elements and further point mutation (Figure 1). Simultaneous SNP typing of plasmid and host enabled us to differentiate between the clonal expansion of MDR S. Typhi, and independent acquisitions of related MDR plasmids by distinct S. Typhi hosts. Evidence for the latter includes the detection of PST2 and PST8 plasmids in co-circulating S. Typhi isolates of distinct haplotypes in the 1970s and 1980s (Table 5). This indicates that the emergence of MDR typhoid during this period was in part due to transfer of IncHI1 plasmids within local S. Typhi populations. One of the PST2-S. Typhi combinations (chromosomal haplotype H42) was later detected among two isolates from Africa in 2003–2004, suggesting that an individual IncHI1 plasmid may be able to persist in a single host haplotype for decades (Table 5). In stark contrast, all 193 PST6 plasmids were observed in S. Typhi of the H58 haplotype and virtually all MDR S. Typhi observed after 1995 belonged to the same PST6-H58 clone, indicative of global spread of MDR typhoid via clonal expansion. Since humans are the only known reservoir for S. Typhi [56], it is likely that trans-continental spread of this clone depends on international travel or migration. If this is the case it will be particularly difficult to control since S. Typhi can be transmitted by asymptomatic carriers [57], [58], who are usually unaware of their status and are difficult to detect [59], [60]. Our data suggest that the PST6 plasmid was acquired by the most recent common ancestor of S. Typhi H58 (Figure 4), implying that the expansion of S. Typhi H58 did not begin until after acquisition of the plasmid. To our knowledge, the oldest confirmed S. Typhi H58 isolate is 9105928K [41], which was isolated in India in 1991 and is MDR (Mia Torpdhal, personal communication). This suggests that the initial expansion of S. Typhi H58 may have been associated with the acquisition of the PST6 plasmid, implying a selective advantage over and above that of MDR, which was also conferred by other IncHI1 plasmid types circulating in S. Typhi in the 1990s. The only growth advantage we detected for PST6 plasmids via our phenotype screen was that of osmotolerance, which we showed to be conferred by the Tn6062 transposon carried by PST6 plasmids. The transposon Tn6062 includes betU (SPAP0106), which encodes a betaine uptake system capable of transporting glycine betaine and proline betaine [61]. It was first described in E. coli isolates causing pyelonephritis (ascending urinary tract infection) and is believed to be an osmoregulator, allowing E. coli to survive the high osmolarity and urea content in urine [61]. However the gene is distributed among E. coli with a range of pathogenic phenotypes, so its osmoprotectant properties may be useful in other environmental contexts [62]. It is possible that enhanced osmotolerance may enhance survival of S. Typhi in specific niches within the human body, including the gut, gall bladder, urinary tract or liver. It is also possible that the ability to grow in the presence of high salt concentrations might enable S. Typhi to continue replicating in certain environments outside the host, which may lower the infectious dose or enhance the possibility of transmission by increasing the level of S. Typhi contamination in certain environments. This may have contributed to the selection of PST6 over other IncHI1 plasmids previously circulating among S. Typhi and the initial clonal expansion of H58 S. Typhi, however questions remain as to why the PST6 plasmid has not been detected among non-H58 S. Typhi. The PST6 plasmid appears to have been lost from H58 S. Typhi in some areas where the recommended treatment of typhoid was switched to fluoroquinolones, including Nepal and Vietnam [39], [63], [64], while it has been maintained in areas such as Kenya where chloramphenicol is still commonly used to treat typhoid [17], [22]. This confirms that antimicrobial use exerts a strong selective pressure for maintenance of the IncHI1 plasmid among S. Typhi and indicates that in the absence of such pressure any additional advantages conferred, including the increased osmotolerance described above, is not enough to maintain the PST6 plasmid indefinitely. Supporting Information Table S1 Bacterial isolates analyzed in this study. (XLS) Click here for additional data file. Table S2 IncHI1 SNP loci targeted in this study. (XLS) Click here for additional data file. Table S3 Biolog phenotype array results. (XLS) Click here for additional data file.
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- Abstract: found
- Article: found
The Global Enteric Multicenter Study (GEMS): Impetus, Rationale, and Genesis
Myron Levine, Karen L Kotloff, James Nataro … (2012)
Diarrheal disease remains one of the top 2 causes of young child mortality in the developing world. Whereas improvements in water/sanitation infrastructure and hygiene can diminish transmission of enteric pathogens, vaccines can also hasten the decline of diarrheal disease morbidity and mortality. From 1980 through approximately 2004, various case/control and small cohort studies were undertaken to address the etiology of pediatric diarrhea in developing countries. Many studies had methodological limitations and came to divergent conclusions, making it difficult to prioritize the relative importance of different pathogens. Consequently, in the first years of the millennium there was no consensus on what diarrheal disease vaccines should be developed or implemented; however, there was consensus on the need for a well-designed study to obtain information on the etiology and burden of more severe forms of diarrheal disease to guide global investment and implementation decisions. Accordingly, the Global Enteric Multicenter Study (GEMS) was designed to overcome drawbacks of earlier studies and determine the etiology and population-based burden of pediatric diarrheal disease. GEMS, which includes one of the largest case/control studies of an infectious disease syndrome ever undertaken (target approximately 12 600 analyzable cases and 12 600 controls), was rolled out in 4 sites in sub-Saharan Africa (Gambia, Kenya, Mali, Mozambique) and 3 in South Asia (Bangladesh, India, Pakistan), with each site linked to a population under demographic surveillance (total approximately 467 000 child years of observation among children <5 years of age). GEMS data will guide investment and help prioritize strategies to mitigate the morbidity and mortality of pediatric diarrheal disease.
Author and article information
Contributors
Sydney Malama: sydneymalama1971@gmail.com
Journal
Journal ID (nlm-ta): BMC Infect Dis
Journal ID (iso-abbrev): BMC Infect. Dis
Title:
BMC Infectious Diseases
Publisher:
BioMed Central
(London
)
ISSN
(Electronic):
1471-2334
Publication date
(Electronic):
2
February
2017
Publication date PMC-release: 2
February
2017
Publication date Collection: 2017
Volume: 17
Electronic Location Identifier: 117
Affiliations
[1
]Department of Epidemiology and Biostatistics, School of Public Health and Social Sciences,
Muhimbili University of Health and Allied Science, Dar es Salaam, Tanzania
[2
]GRID grid.415734.0, , Tanzania Field Epidemiology and Laboratory Management Program, Ministry of Health,
; Dar es Salaam, Tanzania
[3
]ISNI 0000 0000 8914 5257, GRID grid.12984.36, Department of Disease Control, , School of Veterinary, University of Zambia, ; Lusaka, Zambia
[4
]ISNI 0000 0000 8914 5257, GRID grid.12984.36, Health Promotions Research Program, Institute of Economic and Social Research, , University of Zambia, ; Lusaka, Zambia
Article
Publisher ID: 2232DOI: 10.1186/s12879-017-2232-0
PMC ID: 5290660
PubMed ID: 28152988
SO-VID: 248495ef-d6ca-4d41-a82a-038acc9b5541
Copyright © © The Author(s). 2017
License:
Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License ( http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver ( http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.
History
Date
received
: 21
November
2016
Date
accepted
: 31
January
2017
Funding
Funded by: Intra-ACP Academic Mobility Scheme of the Commission of the European Union
Award ID: 2012-3166
Award Recipient
:
Harriet Chiyangi
Categories
Subject:
Research Article
Custom metadata
issue-copyright-statement © The Author(s) 2017
ScienceOpen disciplines: Infectious disease & Microbiology