Independent Microevolution Mediated by Mobile Genetic Elements of Individual Clostridium difficile Isolates from Clade 4 Revealed by Whole-Genome Sequencing
ABSTRACT
Horizontal gene transfer of mobile genetic elements (MGEs) accounts for the mosaic genome of Clostridium difficile, leading to acquisition of new phenotypes, including drug resistance and reconstruction of the genomes. MGEs were analyzed according to the whole-genome sequences of 37 C. difficile isolates with a variety of sequence types (STs) within clade 4 from China. Great diversity was found in each transposon even within isolates with the same ST. Two novel transposons were identified in isolates ZR9 and ZR18, of which approximately one third to half of the genes showed heterogenous origins compared with the usual intestinal bacterial genes. Most importantly, catD, known to be harbored by Tn4453a/b, was replaced by aac(6′) aph(2′′) in isolates 2, 7, and 28. This phenomenon illustrated the frequent occurrence of gene exchanges between C. difficile and other enterobacteria with individual heterogeneity. Numerous prophages and CRISPR arrays were identified in C. difficile isolates of clade 4. Approximately 20% of spacers were located in prophage-carried CRISPR arrays, providing a new method for typing and tracing the origins of closely related isolates, as well as in-depth studies of the mechanism underlying genome remodeling. The rates of drug resistance were obviously higher than those reported previously around the world, although all isolates retained high sensitivity to vancomycin and metronidazole. The increasing number of C. difficile isolates resistant to all antibiotics tested here suggests the ease with which resistance is acquired in vivo. This study gives insights into the genetic mechanism of microevolution within clade 4.
IMPORTANCE Mobile genetic elements play a key role in the continuing evolution of Clostridium difficile, resulting in the emergence of new phenotypes for individual isolates. On the basis of whole-genome sequencing analysis, we comprehensively explored transposons, CRISPR, prophage, and genetic sites for drug resistance within clade 4 C. difficile isolates with different sequence types. Great diversity in MGEs and a high rate of multidrug resistance were found within this clade, including new transposons, Tn4453a/b with aac(6′) aph(2′′) instead of catD, and a relatively high rate of prophage-carried CRISPR arrays. These findings provide important new insights into the mechanism of genome remodeling within clade 4 and offer a new method for typing and tracing the origins of closely related isolates.
INTRODUCTION
Clostridium difficile, a Gram-positive, anaerobic bacterium, has emerged as the leading cause of antimicrobial and health care-associated diarrhea in humans (1). C. difficile is widespread in the environment and the gastrointestinal tracts of humans and animals (2). CD630, which was the first fully sequenced and annotated closed genome of C. difficile, comprises a large circular chromosome of 4.3 Mb, with low GC content (29.06%) (3, 4). Subsequently, thousands of whole-genome sequences (WGS) of C. difficile were published, with sizes ranging from 4.1 to 4.3 Mb (5–7). The high proportion (about 11% in strain 630) of mobile genetic elements (MGEs) contributes to the remarkable dynamic and mosaic genome of C. difficile (8). The MGEs include transposable and conjugative elements, plasmids, bacteriophages, insertion sequences (IS), clustered regularly interspersed short palindromic repeat (CRISPR) elements, group I introns, and sigK intervening (skin) elements (9, 10). These elements play a role in horizontal gene transfer (HGT) between distinct C. difficile isolates and between C. difficile and other intestinal pathogens, such as Enterococcus faecalis, and Enterococcus faecium (9, 11–13).
According to the multilocus sequence typing (MLST) scheme established by Griffiths et al. (14), five distinct phylogenetic lineages (clades 1 to 5) are widely recognized, and an additional clade, clade C-I, was identified, which was confirmed by WGS studies (15–17). Clade 1 is highly heterogeneous and includes many clinically significant toxigenic and nontoxigenic MLST sequence types (STs) and PCR ribotypes (RTs), such as RTs 014, 002, and 018 and STs 2, 14, and 49 (18, 19). A representative of clade 2 is hypervirulent RT027, which has caused severe outbreaks, especially in Europe and North America (20–23). Within clade 3, RT023 and STs 5 and 22 were isolated from humans in Europe (18, 24). In clade 4, RT017 (ST37) has been associated with outbreaks in Europe (25, 26) and North America (27) and is responsible for most C. difficile infections (CDIs) in Asia (28). In addition, RT017 is often clindamycin and fluoroquinolone resistant. Clade 5 contains not only RT078 but also RTs 033, 045, 066, 126, 127, 237, 280, 281, and 288 from a diverse collection of clinical, animal, and food sources worldwide and also demonstrates great heterogeneity (29, 30).
In this study, the WGS of 37 clinical C. difficile isolates from clade 4 with divergent origins in China were obtained, annotated, and analyzed. The genomic regions encoding toxins A and B (PaLoc) and the binary toxin (CdtLoc), MGEs and molecular typing (MLST and PCR ribotyping) were compared and analyzed systematically based on the WGS data. In addition, antibiotic resistance genes, putative resistance point mutation sites, and the relationship with the results of drug susceptibility tests were evaluated. These results will further our knowledge of the genetic characterization of clade 4 and further elucidate the role of MGEs in the evolution, pathogenicity, and transfer of drug resistance genes and toxin-related genes of C. difficile.
RESULTS
Genomic characteristics and population structure in clade 4.
Details of the genome sizes of these 37 C. difficile isolates (range from 3.9 to 4.3 Mb), and other common features are summarized in Table S1 in the supplemental material. The population structure of clade 4 based on the neighbor-joining (N-J) phylogenetic tree correlated well with groups defined according to MLST, PCR ribotyping, and toxin profile but showed no obvious correlation with geographic distribution (Fig. 1). RT017 had two distinct sublineages, ST37 and ST81 (Fig. 1). Each ST cluster was highly conserved, although within the ST37 cluster, ZR18 displayed obvious distinctions compared with the rest of the ST37 isolates (Fig. 1). In the subsequent analysis, we explored the reasons for these distinctions at the genomic level. ST332 was a novel ST that we identified with a unique positive tcdA gene, which formed a singleton in the N-J tree (Fig. 1). Two nontoxigenic isolates (ST109 and ST39) also spread as singletons (Fig. 1).
FIG 1
Genetic diversity of MGEs in clade 4 (transposons, prophages, CRISPRs, and plasmids).
(i) Transposons and conjugative transposons. Several transposons (Tns) and conjugative transposons (CTns), including CTn5, Tn4453a/b, Tn5397, Tn5398, Tn6194, Tn916, and novel putative transposons containing regions homologous with CTn2 and CTn7 (Table 1), were identified by comparison with reference genomes. A novel putative transposon element was discovered in isolate ZR9. This element was located in scaffold 35 from 2,689 to 3,4943 bp and contained 36 coding sequences (CDS), of which 10 were unique in this element, while the remaining 26 genes demonstrated 83.33% to 100% identity with CTn2 (Fig. 2A). Among the 10 unique genes, G002737, G002739, G002743, G002760, and G002761 were annotated as C. difficile putative conjugative transposon protein Tn1549-like, CTn2-ORF30, CTn2-ORF32, CTn5 ORF2, 18, and 19, respectively. G002741 was annotated as putative RNA methyltransferase of C. difficile E7. G002744, G002747, G002748, and G002759 were defined as hypothetical protein, bacterial regulatory helix (AraC family protein), transcriptional regulator (effector binding domain protein), and transcriptional regulator (AbrB family domain protein), respectively. In isolate ZR18, another putative novel mobile element was found, including 24 coding sequences, of which 10 genes were absent in CTn7, while the other genes were similar to those in CTn7 (Fig. 3A). Large fragment insertions and deletions located in 630_03693 to 630_03688, and 630_03686 to mgtA2 were identified in isolate ZR18 (Fig. 3A). The 10 novel genes (G001824, G001827, G001828, G001967, G001969, G001972, G001974, G001975, G001976, and G001977) in ZR18 were annotated as hypothetical protein, CTn7-ORF 4, 6, and conjugal transfer protein of C. difficile, and also proteins of other gut pathogens such as Firmicutes and Enterococcus faecium. In addition, a CTn7-like element was identified in reference M68, showing >85% identity with 630 CTn7 (Fig. 3A). Two genes (M68GM003529 and M68GM003530) were inserted between 630_03695 and 630_03694. In M68, M68GM003535 replaced 630_03690 at the same site, while 630_03685 was missing (Fig. 3A). It is noteworthy that nearly half of genes in CTn7 were lost in M68, with the addition of another 11 coding genes (Fig. 3A), encoding transcriptional regulator, ABC transporter, antimicrobial resistance gene ermB (M68GM003552), and bcrA (M68GM003544), putative truncated zeta protein, ATPase-associated proteins, and other genes originating from C. difficile and other enteric bacteria (Erysipelotrichaceae, E. faecalis, E. faecium, Campylobacter jejuni, Firmicutes, and Staphylococcus aureus).
FIG 2
FIG 3
TABLE 1
| Isolate | Origin | ST | RT | Toxin gene profile | Tna | Potential plasmid sequence | No. of prophagesb |
|---|---|---|---|---|---|---|---|
| 35 | Beijing | 332 | UN | A+B+C+R+E+ cdtA− cdtB− | Tn5398-like | 1 | 7 |
| 2 | Shanghai | 81 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, Tn4453a/b, Tn5397-like, Tn6194-like | 14 | 11 |
| 28 | Shanghai | 81 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, Tn4453a/b, Tn5397-like, Tn5398-like, Tn6194-like | 13 | 7 |
| 7 | Beijing | 81 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, Tn4453a/b, Tn5397-like, Tn5398-like, Tn6194-like | 14 | 8 |
| 15 | Shanghai | 81 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, Tn5398-like | 4 | 9 |
| 10122 | Beijing | 39 | 085 | A−B−C−R−E− cdtA− cdtB− | CTn5-like | 5 | 5 |
| 11032 | Beijing | 109 | UN | A−B−C−R−E− cdtA− cdtB− | 4 | 3 | |
| 16 | Shanghai | 37 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, Tn5398-like, Tn6194-like | 9 | 8 |
| 23 | Xi’an | 37 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, Tn5397-like, Tn5398-like, Tn6194-like | 5 | 7 |
| 29 | Shanghai | 37 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, Tn5397-like, Tn5398-like, Tn6194-like | 10 | 6 |
| 38 | Shanghai | 37 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, Tn5397-like, Tn6194-like | 14 | 8 |
| 4 | Hangzhou | 37 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, Tn5397-like, Tn5398-like, Tn6194-like | 10 | 8 |
| 5 | Beijing | 37 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, Tn5397-like, Tn5398-like, Tn6194-like | 13 | 9 |
| 50 | Shandong | 37 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, Tn5398-like | 1 | 5 |
| 6 | Shanghai | 37 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, Tn5397-like, Tn5398-like, Tn6194-like | 10 | 7 |
| BJ08 | Beijing | 37 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, Tn5397-like, Tn5398-like | 9 | 7 |
| GZ2 | Guangzhou | 37 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, Tn5397-like, Tn5398-like, Tn6194-like | 10 | 7 |
| GZ3 | Guangzhou | 37 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, Tn5397-like, Tn6194-like | 10 | 8 |
| GZ6 | Guangzhou | 37 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, Tn5397-like, Tn5398-like, Tn6194-like | 10 | 8 |
| GZ8 | Guangzhou | 37 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, Tn5397-like | 10 | 9 |
| GZ11 | Guangzhou | 37 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, Tn5397-like, Tn6194-like | 11 | 8 |
| GZ12 | Guangzhou | 37 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, Tn5397-like, Tn6194-like | 10 | 7 |
| GZ13 | Guangzhou | 37 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, Tn5397-like, Tn5398-like, Tn6194-like | 13 | 9 |
| GZ14 | Guangzhou | 37 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, Tn5397-like, Tn5398-like, Tn6194-like | 13 | 11 |
| ZR58 | Beijing | 37 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, Tn5397-like, Tn6194-like | 14 | 11 |
| ZR59 | Beijing | 37 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, Tn5397-like, Tn6194-like | 14 | 13 |
| ZR65 | Beijing | 37 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, Tn5398-like | 4 | 8 |
| ZR66 | Beijing | 37 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, Tn5397-like, Tn5398-like, Tn6194-like | 14 | 13 |
| ZR68 | Beijing | 37 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, Tn5398-like | 5 | 8 |
| ZR72 | Beijing | 37 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, Tn5397-like, Tn5398-like | 9 | 8 |
| ZR73 | Beijing | 37 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, Tn5397-like, Tn5398-like | 8 | 10 |
| ZR82 | Beijing | 37 | 017 | A-B+C+R+E+cdtA-cdtB- | CTn5-like, Tn5397-like, Tn5398-like | 8 | 7 |
| ZR29 | Beijing | 37 | 017 | A-B+C+R+E+ cdtA− cdtB- | CTn5-like, Tn5397-like, Tn5398-like, Tn6194-like | 14 | 9 |
| ZR18 | Beijing | 37 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, npe-CTn7, Tn4453a/b, Tn5397-like | 7 | 5 |
| ZR8 | Beijing | 37 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, Tn5397-like, Tn5398-like, Tn6194-like | 10 | 6 |
| ZR9 | Beijing | 37 | 017 | A−B+C+R+E+ cdtA− cdtB− | npe-CTn2, Tn5397-like, Tn6194-like | 9 | 6 |
| HN9 | Henan | 37 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, Tn5397-like, Tn5398-like | 9 | 8 |
| CD630 | 54 | 012 | A+B+C+R+E+ cdtA− cdtB− | ||||
| M68 | Ireland | 37 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like, CTn7-like, Tn5397-like | ||
| CF5 | Belgium | 86 | 017 | A−B+C+R+E+ cdtA− cdtB− | CTn5-like |
a
npe, novel putative element.
b
Total number of incomplete, intact, and questionable prophage.
CTn5-like transposons, belonging to the Tn1549 family, were detected widely in the tested C. difficile isolates of clade 4, with the exception of isolate 35 and 11032 (Fig. 2B). Ten types were divided according to gene composition as follows (Fig. 2B): type 1 (HN9), type 2 (BJ08), type 3 (15), type 4 (GZ8), type 5 (ZR18), type 6 (GZ3), type 7 (29), type 8 (2, 28, 5, 7, ZR8, ZR9), type 9 (23, 38, 4, 50, 6, GZ11, GZ12, GZ2, GZ6, ZR29, ZR58, ZR59, Z566, 10122), and type 10 (16, GZ13, GZ14, ZR65, ZR68, ZR72, ZR73, ZR82). Two common major deletions in the conjugative region were observed in these 10 types, one being a deletion of 630_02052, and the other one located between 630_02066 and 630_02072 (Fig. 2B). Especially for types 3, 5, and 8, other large fragment deletions were also discovered in the conjugative region (Fig. 2B).
Tn4453a/b was found with typical gene organization in isolates ZR18, 2, 7, and 28 (Fig. 3B). The composition and location of genes in ZR18 was identical to that in reference Tn4453a/b, with the exception of a −114-bp deletion in tnpv (Fig. 3B). The key feature of this element was the presence of catD, which mediates resistance to chloramphenicol. Interestingly, catD was absent in isolates 2, 7, and 28 and replaced by five new genes: aminoglycoside acetyltransferase (aac), three transferases (phosphotransferase, nucleotidyltransferase, and acetyltransferase) and an ATPase (Fig. 3B). The aac gene, which confers resistance to aminoglycoside antibiotics, such as gentamicin, tobramycin, and netilmicin has been widely reported in Enterococcus and Enterobacteria, and can be transferred between Gram-positive and Gram-negative bacteria by transposons (Tn5281, Tn4001, and IS256) or plasmids (31). BLAST analysis revealed that the DNA sequences of aac in isolates 2, 7, and 28 showed 100% coverage and identity with aac(6′) aph(2′′) in C. jejuni. The total length of the gene was 1,455 bp, encoding a bifunctional aminoglycoside N-acetyltransferase and aminoglycoside phosphotransferase, which is one of the aminoglycoside-modified enzymes (AMEs), playing critical role in high-level gentamicin-resistant (HLGR) of Enterococcus (32).
Tn5397, also known as CTn3 in CD630, was recognized in most isolates in this study and was classified into four types as follows: type 1, ZR18 and M68; type 2, ZR73 and 5; type 3, BJ08, GZ8, and HN9; type 4, ZR82, ZR9, 2, 23, 28, 29, 38, 4, 6, 7, GZ11, GZ12, GZ13, GZ14, GZ2, GZ3, GZ6, ZR29, ZR58, ZR59, ZR66, ZR72, and ZR8 (Table 1). The tndX gene, encoding a large serine recombinase, which is essential for insertion/excision of Tn5397, was absent in all types (Fig. 3C). Furthermore, a group II intron within orf14 was also deleted (Fig. 3C). Significantly, in type 2, Tn5397 was missing some of the downstream genes, including an antimicrobial resistance gene tetM, compared with the reference Tn5397 (Fig. 3C). Type 1 displayed the most similar gene composition and structure compared with the reference Tn5397, although two new genes were inserted (Fig. 3C).
Tn5398, first reported in strain 630, contains erm 1B and erm 2B, and can be transferred between C. difficile isolates, or between C. difficile and S. aureus or Bacillus subtilis (33). Two types of Tn5398-like elements were identified here, in which erm 1B and erm 2B genes were absent (Fig. 3D). Type 1 consisted of 35, ZR73, ZR82, 16, 23, 28, 29, 4, 5, 50, 6, 7, BJ08, GZ13, GZ14, GZ2, GZ6, HN9, ZR29, ZR65, ZR66, ZR68, ZR72, and ZR8, while type 2 was represented only by isolate 15 (Table 1).
Tn916 is a major family transposon reported in C. difficile and carries the tetM resistance gene. The Tn916 element of these tested isolates were subdivided into multiple types as follows: type 1, GZ8; type 2, ZR18; type 3, ZR73; type 4, 23, 29, 4, 6, GZ11, GZ12, GZ13, GZ14, GZ2, GZ3, GZ6, ZR72, ZR8, ZR82, and ZR9; type 5, BJ08 and HN9; type 6, 5; type 7, 2, 28, 38, 7, ZR29, ZR58, ZR59, and ZR66 (Table 1). The tetM resistance and int gene (responsible for Tn insertion), which are responsible for insertion, appeared in all types (Fig. 2C). With the exception of type 6 with deletions in three large regions, deletions occurred in a single area of the regulation region in all the other types (Fig. 2C). Intact xis and int genes were retained in types 3, 4, 6, and 7. The conjugative region was highly homologous within most types, except for types 3, 6, and 7, which had large fragment deletions (Fig. 2C). Isolate ZR18 carried no typical xis and int genes of Tn916, while BJ08, GZ8, and HN9 contain int without xis.
The representative gene cluster of Tn6194 carrying ermB of strain CII7 (GenBank accession no. HG475346) was used as a reference in this study. Tn6194 has a conjugation region that is closely related to that of Tn916 but contains an accessory region that is related to Tn5398. M68 demonstrated high-level homology with only the absence of cds22 and cds 23, and indels within cds5 and cds 11, while intact xis and int genes were retained for excision and insertion (Fig. 2D). The following six types were defined here: type 1, 16, 2, 28, 38, 5, 7, GZ13, and GZ14; type 2, 23; type 3, ZR66; type 4, ZR29, ZR58, and ZR59; type 5, 29, 4, 6, GZ11, GZ12, GZ2, GZ6, ZR8, and ZR9; type 6, GZ3 (Table 1).The ermB gene was present in all types except type 2 (Fig. 2D). The xis gene, which is involved in the excision of Tn916 from the donated strain were deleted in all types (Fig. 2D).The int gene, which plays a key role in the integration of Tn916 into the recipient isolate, appeared in types 2, 5, and 6 only (Fig. 2D).
(ii) Prophages, plasmids, and CRISPR. Prophages were identified in various amounts in all tested isolates (Table S2). In total, 497 prophages were predicted in this study. Isolates 2 and ZR66 contained the highest number of prophages (19 prophages), while isolate 15 had the fewest (3 prophages) (Table S2 and Fig. 4A). A total of 115 intact prophages was confirmed among in all the isolates, except ZR18, 15, and 11032 (Fig. 4A). Among the intact prophages, the number of ΦCDHM19, ΦCD506, ΦMMP01, ΦMMP03, ΦMMP02, ΦMMP04, and ΦC2 was more than three per isolate (Fig. 4B). ΦCDHM19 and ΦCD506 represented more than half (54.78%) of all intact prophages (Fig. 4B). All of these prophages belonged to the Myoviridae family, and some were induced during C. difficile infection (34). Most prophages identified here were homologous to known phages reported in C. difficile, although a few were similar to other bacterial phages, such as C. jejuni, S. aureus, Enterobacteria, Bacillus, and other unusual bacteria, such as Prochlorococcus, Gordonia, and Sphingomonas (Table S2). Isolates 50 and 35 carried the highest number of intact prophages, 8 and 7, respectively (Table S2 and Fig. 4A). The P-SSM2-like phage of Prochlorococcus found in isolate 50 showed the highest number of proteins similar to those in the region (Table S2), while the ΦC2-like phage in isolate 35 displayed a high number of “hit_genes_count” in bracket (8 out of 9) with ΦC2 (Table S2). The prophages found within the ST 37 and ST 81 isolates were diverse, although ΦCD506 was predominant. No obvious correlation was found between prophage type and STs. Future studies will focus on the role of these elements in the C. difficile evolution and infection.
FIG 4
A total of 464 CRISPR arrays, composed of different copies of direct repeat (DR) sequences, and separated by unique spacers were identified in the tested isolates from clade 4 (Table S3). The number in each isolate varied from 5 to 19, and the length ranged from 60 to 1,608 bp (Table S3). In total, 90 of the CRISPR arrays displayed homology with prophages (Table S3). Isolates 10122, 11032, 29, and GZ8 harbored prophages without homologous CRISPR sequences (Table S3).
Distinctions of PaLoc and CdtLoc regions in clade 4.
Three types of toxin gene profiles in clade 4 were covered in this study: A+B+ (ST332), A−B+ (ST81 and ST37), and A−B− (ST39 and ST109) (Table 1). All isolates in this study were binary toxin negative (Table 1). Based on gene formation, six types of PaLoc were classified in this study (Fig. 5A). Type 1 (ST332) displayed intact gene composition and sequences, while all genes were lost with the exception of 115 bp remaining in type 6 (ST39 and ST109) (Fig. 5A). The other four types lack a complete and continuing tcdA gene, attributed to insertions and single nucleotide polymorphisms (SNPs), which led to termination of tcdA gene translation (Fig. 5A). Compared with CD630, the SNPs of genes located within PaLoc regions are shown for each isolate in Fig. 5B. In general, diversity within PaLoc regions correlated with its ST type, with some exceptions (Fig. 5B). For example, isolate 15 within ST81 and ZR18 within ST37 showed distinct SNP distribution (Fig. 5B). The tcdB gene demonstrated great heterogeneity, followed by tcdA. With the exception of isolate 35, all tested isolates lacked the CdtLoc region (Fig. 5C). Isolate 35 contained a complete cdtR gene and fragments of cdtA and cdtB genes (Fig. 5C).
FIG 5
Antimicrobial resistance gene and related antimicrobial susceptibility.
In this study, 60 antibiotic resistance genes were identified by comparison with the ARDB and CARD databases (35, 36). These antibiotic resistance genes belonged to the following antimicrobial classes: fluoroquinolones, β-lactam antibiotics, macrolides, aminocoumarin, rifampin, glycopeptides, tetracycline, macrolide-lincosamide-streptogramin (MLS), chloramphenicol, trimethoprim, streptomycin, aminoglycoside, bacitracin, and lipopeptide (Fig. 6). Most isolates carried approximately 13 to 16 copies of the macB gene, except for ZR18 carrying 9 copies. The next highest numbers of copies were for the vanRI and bcrA genes (Fig. 6). Remarkably, some isolates carried a unique antibiotic resistance gene cassette, for example, ZR18 and CF5 contained a unique antibiotic resistance gene cluster of approximately 6 kb comprising vanXYG, vanSG, vanTG, and vanG, which were integrated as glycopeptide resistance gene (Fig. 6). Additionally, ZR18 also carried catD (carried by Tn4453a/b), and cata11, which are related to chloramphenicol resistance, and tetA(48) (Fig. 6). No functional vancomycin resistance operon was identified here. Furthermore, isolate 11032 contained unique tetA(P), tetT, tetpb, and tetpa genes (Fig. 6). In addition, ZR9 carried a unique novA gene, and M68 had unique cata1 and catl genes (Fig. 6).
FIG 6
According to the Etest results, all isolates except for 10122 and 35 were MDR strains, maintaining a resistance rate as high as 97.30%. Details are summarized in Table 2. Almost all MDR strains displayed high levels of drug resistance (Table 2) with quinolone exhibiting 100% resistance to CIF, followed by LVX (97.30%) and MXF (56.76%) (drug abbreviations in “Antimicrobial susceptibility tests” in Materials and Methods) (Table 2). The quinolone resistance is due to a point mutation and substitution within gyrA and/or gyrB. Clade 4 C. difficile retained high-level resistance (91.89%) to ERY compared with the other isolates of other clades reported in Table 2. The ermB gene, encoding an antibiotic target-modifying enzyme, is related to MLS resistance. The ermB gene was detected in 25 of 34 ERY-resistant isolates (73.53%). ERY resistance is attributed to methylation of position 2058 in 23S rRNA; however, the exact mutation in ERY-resistant C. difficile isolates remains to be identified in the future. In addition, high copy numbers of the macrolide resistance macB gene, encoding a macrolide ABC transporter protein, were identified in all the tested isolates (Fig. 6). The rate of resistance to CLI was 89.19%, with a high level of resistance detected among the isolates. The rate of resistance to CHL was lower (29.73%) and related to cata11 and cata1, which were identified only in isolates ZR18 and M68, respectively, indicating the existence of other mechanisms of CHL resistance. It is worth mentioning that catD, normally carried by Tn4453a/b and also responsible for CHL resistance (Fig. 3), was present in isolate ZR18, which showed an intermediate level of CHL resistance (MIC = 64 μg/ml). However, catD was replaced by another five genes in isolates 7, 2, and 28, which were CHL susceptible. The five new replacement genes included aac(6′) aph(2′), which is correlated significantly with HLGR and is responsible for aminoglycoside resistance, leading to gentamicin and amikacin resistance in isolates 7, 2, and 28. The rates of MEM (51.35%) and RIF (48.65%) resistance were similar among the clade 4 C. difficile isolates. All isolates resistant to CHL and RIF were ST37 (Table 2). The rpoB and rphB genes with several mutations related to RIF resistance were detected in almost all isolates. The mechanism underlying this resistance involves substitution of several nucleotides in the rpoB gene. None of the clade 4 isolates were resistant to VAN or MET, although VAN-related genes or gene cassettes were identified.
TABLE 2
| Drug (concn range [μg/ml]) | No. of resistant isolates (%)a | STb | No. of genes identifiedb | MIC (μg/ml)b |
|---|---|---|---|---|
| MXF (0.002–32) | 21 (56.76) | ST81 (4), ST109 (1), ST37 (16) | E. coli gyrB (37), M. tuberculosis gyrA (18), C. jejuni gyrA (20) | ≥32 (21) |
| LVX (0.002–32) | 36 (97.30) | ST332 (1), ST39 (1), ST109 (1), ST81 (4), ST37 (29) | As above | ≥32 (35) |
| ≥8 < 32 (1) | ||||
| CIF (0.002-32) | 37 (100) | ST332 (1), ST39 (1), ST109 (1), ST81 (4), ST37 (30) | As above | ≥32 (36) |
| ≥8 < 32 (1) | ||||
| CLI (0.016–256) | 33 (89.19) | ST81 (4), ST109 (1), ST37 (28) | ermB (25), cfrC (4), clbA (11) | ≥256 (32) |
| ≥8 < 256 (1) | ||||
| TET (0.016–256) | 13 (35.14) | ST81 (2), ST37 (11) | tetA(P), tetT, tetpb, tetpa were only in isolate 11032; tetA(48)c was only in ZR18; tetM (31), tet44 (30) | ≥16 < 32 (13) |
| ERY (0.016–256) | 34 (91.89) | ST81 (4), ST39 (1), ST109 (1), ST37 (28) | ermB (25), macB (37), cfrC (4) | ≥256 (32) |
| ≥128 < 256 (2) | ||||
| ≥8 < 128 (1) | ||||
| CHL (0.016–256) | 11 (29.73) | ST37 (11) | cata11 was only in ZR18; cata1 was only in M68 | ≥256 (8) |
| ≥128 < 256 (1) | ||||
| ≥32 < 128 (2) | ||||
| MEM (0.002–32) | 19 (51.35) | ST81 (1), ST109 (1), ST37 (17) | blaR1 (36), mecI (35) | ≥32 (15) |
| ≥16 < 32 (4) | ||||
| RIF (0.002–32) | 18 (48.65) | ST37 (18) | rpoB (36), rphB (34) | ≥32 (18) |
| VAN (0.016–256) | 0 | 0 | vanXYG-vanSG-vanTG-vanG gene cluster was only in ZR18; vanRI (37), vanRM (37), vanRA (37), vanRF (36), vanZF (36), vanRE (37), vanRB (37), vanSA (36), vanRG (37), arlR (34) | ≤0.5 |
| MET (0.016–256) | 0 | 0 | None | ≤1 |
a
Values in parentheses show the percentage of isolates resistant to a single drug.
b
Values in parentheses show the number of isolates with the ST, gene, or MIC.
c
Gene name.
Genome-wide association analysis of resistance genes.
According to our analysis, antimicrobial susceptibility does not correlate well with genotypes. To explore other potential sites responsible for drug resistance, we performed genome-wide association study (GWAS) analysis. Significantly related SNPs were detected in relation to resistance to CHL, MFX, and RIF (Fig. 7). Three significant SNPs associated with CHL resistance were located within an intergenic genetic region (IGS) (Fig. 8), which may play a role in methylation and regulation of cell function. Three remarkable nonsynonymous SNPs associated with MFX resistance (Fig. 7) were detected in genes encoding inosine-uridine-preferring nucleoside hydrolase family protein and the ABC transporter and in the gyrA gene, which are reported to be associated with quinolone resistance, and were detected using all three methods (Fig. 7). All MFX-resistant isolates except for GZ11 and 28 displayed the point mutation (Thr→IIe) in gyrA compared with susceptible isolates (Fig. 8). In addition, only one MFX-susceptible isolate ZR66 carried the Thr→IIe mutation in gyrA (Fig. 8). As shown in Fig. 7, four significant SNPs significantly related to RIF resistance were identified as following: a gene (dacC or dacD) encoding D-alanyl-D-alanine carboxypeptidase protein, also known as penicillin-binding protein (WP_022616436.1), gene merR encoding the transcriptional regulator (WP_022616524.1), a gene involved in replication, recombination, and repair (WP_022616400.1), and a gene in an IGS region (Fig. 8). Most of the RIF-resistant isolates carried the point mutation Ala→Val, Thr→IIe, and Val→Gly in these genes, respectively, with the exception of isolates GZ3, 6, 29, and 23 (Fig. 8). The RIF-susceptible isolates 5 and 38 carried the same mutations (Fig. 8).
FIG 7
FIG 8
DISCUSSION
Due to rapid progress in WGS techniques, the global population structure of C. difficile is defined as 5 main clades (clades 1 to 5). ST37 (RT017), which is a well-known representative of clade 4, has caused outbreaks in Europe and in the United States, and is documented as the major type associated with CDI in Asia (8). In addition to ST37, clade 4 contained some diversity, such as ST332 (A+B+), ST39 and ST109 (A−B−), ST81 (A−B+), and ST86 (A−B+). ST332 is a novel type previously identified by our group (37). In this study, we analyzed and compared the whole-genome sequences of 37 clinical C. difficile isolates from clade 4. M68 and CF5 were included as isolates of clade 4, and CD630 was used as reference. We further explored the presence of Tn/CTns, prophages, and CRISPRs and compared their characteristics among the isolates in this clade. The antimicrobial phenotypes of these isolates were determined, and possible antibiotic resistance-related genes were identified. In this study, we aimed to clarify the genetic heterogeneity and microevolution within clade 4 to provide potential explanations for the mechanisms of pathogenesis and antibiotic resistance of C. difficile.
The basic features, including GC% and genome size, of the clade 4 isolates from China investigated in this study were similar to those have been reported previously (3). The C. difficile genome is highly mosaic and dynamic in structure because of a high proportion of MGEs. CTns, also known as integrated conjugative elements (ICEs), are capable of excision, transfer, and integration into the chromosome or plasmids, resulting in the formation of a circular intermediate (12). Within C. difficile, there are two main families of CTns, known as the Tn916- and the Tn1549-like elements. Tn916 is an 18-kb conjugative transposon that encodes resistance to tetracycline and minocycline via the ribosomal protection protein, Tet (M), and contains 24 potential ORFs. Tn1549, a 34-kb vancomycin resistance element controlled via the vanB operon, was originally described in E. faecalis. In CD630, six putative CTns belonging either to the Tn916-like family (CTn1, CTn6, CTn7 plus Tn5397, previously known as CTn3) or to the Tn1549-like family (CTn2, CTn4, and CTn5) were identified. Tn916-like, CTn1-like, and CTn5-like were reported from M68, while CTn5-like was found in CF5. In this study, CTn5-like, Tn5397, and Tn916 elements were identified, with CTn5-like found to exist widely among the investigated clade 4 isolates. However, other transposons, which did not belong to either the Tn916-like or Tn1549-like families, were also discovered, including Tn4453a/b-like, Tn5398-like, and Tn6194-like. Furthermore, two novel putative transposons having regions of homology with CTn2 and CTn7 were identified in isolates ZR9 and ZR18, respectively. It is remarkable that in Tn4453a/b, catD was replaced by aac(6′) aph(2′′) in isolates 2, 7, and 28, resulting in susceptibility to CHL but resistance to gentamicin and amikacin. The three isolates, which were all ST8, were obtained from elderly hospitalized patients treated with quinolone and carbapenem for 5 days. However, isolate ZR18 carried a typical Tn4453a/b element with catD, leading to CHL resistance. The aac(6′) aph(2′′) gene, displaying 100% identity and coverage with C. jejuni, was first reported in C. difficile. This gene encodes a bifunctional AME, accounting for HLGR resistance rates in >90% E. faecalis and E. faecium isolates (38). This illustrated that HGT may occur between C. difficile and other intestinal bacteria, leading to unique microevolution of individual isolates. However, the mechanisms by which the local microenvironment causes HGT and the ability of Tn4453a/b containing aac(6′) aph(2′′) to transfer between C. difficile isolates and/or C. difficile and other bacteria remains to be elucidated. Furthermore, two novel transposons with regions with homology to CTn2 and CTn7 were identified. One novel transposon, in isolate ZR9, contained 10 unique genes showing identity with diverse transposons of diverse origins in C. difficile, although the remaining 26 genes were homologous with CTn2. The other transposon, in isolate ZR18, lacked 10 out of 24 coding genes present in CTn7. The 10 novel genes also displayed identity with C. difficile or other gut pathogens of heterogeneous origins. This situation further suggested that “personalized” HGT occurred in individual isolates, resulting in diversity and genome evolution in C. difficile. Great diversity was also exhibited among the other CTns identified in this study even within the same STs, and several types were classified according to their gene structure and composition.
Sequencing of a number of C. difficile genomes has facilitated the identification of numerous putative prophages by bioinformatic analyses, and the role of prophages in its evolution and virulence has become the focus of recent research. So far, only a limited number of temperate phages infecting C. difficile have been fully characterized in terms of WGS and morphology by transmission electron microscopy (TEM). ΦCD119, ΦC2, ΦCD27, ΦMMP02, and ΦMMP04 are members of the Myoviridae family, while ΦCD38-2 and ΦCD6356 are members of the Siphoviridae family. In this study, WGS revealed distribution of various numbers of prophages in all isolates. The top seven main prophages in clade 4 were ΦCDHM19, ΦCD506, ΦMMP01, ΦMMP03, ΦMMP02, ΦMMP04, and ΦC2, all of which belong to the Myoviridae family. Some prophages (ΦMMP01, ΦMMP03, ΦMMP02, and ΦMMP04) were induced and isolated from fecal supernatant of CDI patients (34). In addition, ΦC2 mediates transduction of Tn6215, encoding erythromycin resistance, between C. difficile isolates (11). Our next step is to induce phages of these isolates and confirm their function experimentally.
The CRISPR-Cas systems, which are present in most archaea and many bacteria, provide adaptive resistance against invasive genetic elements, such as phages and plasmids (40, 41). CRISPR array consists of partial palindromic repeats (DR) interspersed by unique DNA sequences (spacers) that are derived from foreign genetic elements in a linear, time-resolved manner (41, 42). Whole-genome analysis of C. difficile strain 630 (ST54/RT012) showed that the CRISPR spacer sequence exhibited similarity to C. difficile phages and plasmids, suggesting CRISPR interference against these mobile elements, which was later experimentally confirmed in the R20291 strain (ST1/RT027/B1) (3, 43). Recently, in a comprehensive analysis of the CRISPR-Cas system in 217 C. difficile genomes, a single type IB CRISPR-Cas system and a total of 1,865 CRISPR arrays were identified with cas gene clusters present at conserved chromosomal locations (44). The CRISPR arrays of C. difficile were markedly enriched (12.5 arrays/genome) in this study compared with other species and previous reports of C. difficile (8.5 arrays/genome) (45). Numerous CRISPR arrays (90/464) were homologous to prophages identified among the clade 4 C. difficile isolates in this study. CRISPR-Cas has been used for outbreak tracking in Yersinia pestis (46) and Salmonella enterica subspecies (47), and correlating with important phenotypes, such as antibiotic -resistance cassettes in enterococci (48), and prophages in Streptococcus pyogenes genomes (49). All these correlations reflect the role of CRISPR-Cas systems in controlling HGT, as well as the uptake and dissemination of particular genes and operons involved in bacterial adaptation and pathogenesis (50), and hence the evolution of specific species. In our study, we identified another potential method of phylogenetic analysis and genotyping within clade 4 or closely related isolates according to the CRISPR arrays.
Rates of antibiotic resistance vary considerably in different studies, probably depending on the geographic regions and local or national antibiotic policy. In our study, the MDR isolates account for 94.6% of all resistant isolates, which is a much higher proportion than reported previously (55%) in Europe and China (19, 51, 52). Many of the hypervirulent RT027 and RT078 strains are known to exhibit MDR. In a 4-year study in China, MDR isolates with ST37 accounted for the highest proportion (78.2%) of all the tested isolates (53–55). A similar situation was observed in our study, which further illustrates that antibiotic usage promotes independent acquisition of drug resistance, regardless of molecular type.
Resistance to fluoroquinolones has increased dramatically in recent years; for example, the rates of resistance to LVX and CIP reached almost 100% in many studies around the world (51), including our study. Genomic mutations in gyrA and/or gyrB, known as the quinolone resistance-determining region, QRDR, are involved with resistance to fluoroquinolones (56). The amino acid substitution Thr-82Ile in GyrA was found to be responsible for moxifloxacin resistance (57). Some other substitutions in gyrA and/or gyrB have been reported in other studies, such as Ser-366Val, Ser-416Ala, Asp-426Asn, and Glu-466Val (58). According to our GWAS analysis, the Thr→IIe mutation in the gyrA gene is highly related to MXF resistance in clade 4 C. difficile isolates.
RIF resistance was very recently detected, and studies conducted from 2008 to 2010 in different European countries showed variability in the rates of resistance to this antibiotic in C. difficile strains, ranging from 6.3% to 36.8% (58). In our study, the rate of RIF resistance reached 48.65%, which is even higher than that reported in a systematic meta-analysis of drug resistance of C. difficile infections in mainland China (52). Different SNPs (His-502Asn and Arg-505Lys) within the rpoB gene encoding the β-subunit of the RNA polymerase have been identified in RIF-resistant strains (59). Another RIF resistance-related gene, rphB, was also detected in 34 isolates in this study but at a low related rate. The existence of some specific substitution in rphB that is responsible for RIF remains to be investigated.
In C. difficile, resistance to CLI and ERY is common in Europe (55% and 47%, respectively) (19), and in China (70% to 80%) (60), while in our study the rate was more than 90%. In this study, several genes were identified with ermB, high copy numbers of macB, and cfrC, although the correlation between the phenotype and the harbored gene is low. In pan-Europe studies, the rate of TET resistance has been reported to range from 2.4% to 41.67% (51), and the rate in the present study was slightly lower than the highest level at 35.14%. Moreover, the emergence of reduced TET susceptibility increased in our study. In C. difficile, TET resistance is commonly conferred by a tet(M) carried by a Tn5397 transposon. In our study, 29 isolates contained Tn5397; however, tetM was absent from isolates ZR73 and 5. Among these 27 isolates, the rate of TET resistance was 40.74% (11/27). For eight isolates without Tn5397, two resistant isolates (15 and 16) and one intermediate strain (11032) were still identified. In addition, no other TET resistance-determining genes, including tet 44 and tetW (other important genes responsible for TET resistance in C. difficile especially with animal and environmental origins) were detected in these three isolates. This indicates the possible existence of another mechanism of TET resistance.
Although CHL resistance is not so common in Europe (3.5%) and China (2%) (19, 52), the rate obviously increased in our study (29.73%). It is known that catD in Tn4453a/b mediates CHL resistance; however, in our study, only one (ZR18) of 11 resistant isolates carried catD in Tn4453a/b and cata11 in the genome, which may represent another mechanism of gaining CHL resistance in C. difficile. The great heterogeneity in the genetic arrangement of those resistance determinants confirms that genetic exchange and recombination frequently occur in individual clinical strains.
As reported in a previous study in China, no resistant isolates to MTZ and VAN or isolates with reduced MTZ susceptibility were found here, although reduced MTZ susceptibility was occasionally reported in Europe (61). The mechanism of MTZ resistance is still not completely understood. Nitroimidazole (nim) genes conferring resistance to MTZ in Helicobacter pylori have not been identified in C. difficile. The defined mechanism of resistance is a multifactorial process involving alterations in different metabolic pathways, such as nitroreductase activity, iron uptake, and DNA repair (59). A vanB operon originally described in E. faecalis harboring Tn1549 is responsible for resistance to VAN (62). Although several Tn1549-like elements have been reported in C. difficile, no functional vanB operon was identified. Recently, a vanG-like gene cluster, homologous to the cluster found in E. faecalis, has been reported in a number of C. difficile isolates. However, although this cluster is expressed, it does not promote resistance to VAN (63). The same situation was found in isolate ZR18 in our study with a series of vanG-like gene clusters identified, without resistance to VAN.
In conclusion, we analyzed HGT-related elements involved in shaping the genome and biological features, including antibiotic susceptibility phenotype of C. difficile from clade 4 with varied STs isolated in China. Our findings provide important new insights into the mechanism of genome remodeling within clade 4 and offer a new method for typing and tracing the origins of closely related isolates.
MATERIALS AND METHODS
Isolates and preparation of genomic DNA.
A total of 37 clinical C. difficile isolates from clade 4 with divergent origins in China were included in this study (Table 1). These isolates comprised five ST types: one ST332 isolate (a new type identified in our previous study), four ST 81 (RT 017), one each of ST 39 (RT 085) and ST109, and 30 ST 37 (RT 017) isolates. Details of these isolates are summarized in Table 1. Strains CD630 (GenBank accession no. AM180355), CF5 (FN665652), and M68 (FN668375) were used as references throughout the investigation.
All 37 isolates were cultured on brain heart infusion (BHI) agar plates (Oxoid, Basingstoke, UK) supplemented with 5% sheep blood (BaoTe, Beijing, China) in an anaerobic environment (80% nitrogen, 10% hydrogen, and 10% carbon dioxide) (Mart, NL) at 37°C for 48 h. Typical colonies were picked and recultured on BHI for 24 h before preparation of genomic DNA using the Wizard Genomic DNA purification kit (Promega, Madison, WI, USA) according to the manufacturer’s instructions.
MLST, PCR ribotyping, and toxin gene profiling.
Whole-genome sequencing and assembly.
Whole-genome sequencing (WGS) was performed on the Illumina PE150 (Illumina Inc., USA) with average insert lengths of 350 bp. Raw data were processed in four steps: removal of reads with 5 bp of ambiguous base, reads with 20 bp of low quality (≤Q20), adapter contamination, and duplicated reads. Finally, filtered reads were assembled using SOAP denovo v2.04 (65, 66).
Genome annotation and phylogenetic analysis.
Genes were predicted using GeneMarkS version 3.05 (67) and aligned with databases to obtain the annotation corresponding to their homologs, with the highest quality alignment result chosen as the gene annotation. Gene functions were predicted by whole-genome BLAST (http://blast.ncbi.nlm.nih.gov/Blast.cgi) searches against the following databases: Gene Ontology (GO) (68), Kyoto Encyclopedia of Genes and Genomes (KEGG) (69), Clusters of Orthologous Groups (COG) (70), Non-Redundant Protein (NR) databases (71), Transporter Classification Database (TCDB) (72), and Swiss-Prot (73). tRNA genes were predicted by tRNA scan-SE (74). rRNA genes were analyzed by rRNAmmer (75). Small nuclear RNAs (snRNA) were predicted by BLAST searches against the Rfam (76) database.
Single core genes were obtained from the clustering results of all sample genes used by CD-HIT version 4.6. A phylogenetic tree was built using the neighbor-joining method based on 2,456 single core genes using MEGA6.0 with 1,000 replicates (77).
Analysis of MGEs.
All the transposons (Tns) and conjugative transposons (CTns) were identified using BLAST searches of the C. difficile genome sequences or transposon sequences available at NCBI ( https://www.ncbi.nlm.nih.gov/) with identity and coverage requirements set at >80%. Prophages were identified using the PHASTER web server (http://phast.wishartlab.com). The intact, questionable, and incomplete prophage sequences were defined by score values of >90, 70 to 90, and <70, respectively (78). To identify plasmids, reads were assembled into contigs using the SOAP denovo. Contigs were then screened for plasmids using Microbial Genome BLAST searches against the NCBI complete plasmid database (ftp://ftp.ncbi.nlm.nih.gov/genomes/refseq/plasmid/). The potential plasmids were defined as ≥70% coverage and ≥80% identity. The CRISPRFinder (79) was used for CRISPR identification.
Sequence analysis of PaLoc and CdtLoc.
The PaLoc and CdtLoc were confirmed by comparison with the reference CD630 genome. Orthologous genes were detected by BLAST (version 2.2.12) searches with >80% coverage and >90% nucleotide identity. The genetic structure as well as insertions and deletions (indels) were studied.
Antimicrobial susceptibility tests.
C. difficile isolates were tested for susceptibility to moxifloxacin (MXF), vancomycin (VAN), clindamycin (CLI), tetracycline (TET), erythromycin (ERY), rifampin (RIF), levofloxacin (LVX), chloramphenicol (CHL), metronidazole (MTZ), ciprofloxacin (CIP), and meropenem (MEM) using Etest strips (bioMérieux, France, and Liofilchem, Italy). Tests to gentamicin (CEN) and amikacin (AMK) were also performed with isolates 2, 7, 28, and ZR18 using Etest strips (bioMérieux, France, and Liofilchem, Italy). The MICs for MTZ, MXF, CLI, CIP, LVX, and TET were determined according to recommendations of the Clinical and Laboratory Standards Institute (CLSI) M11-A7 and M100-S24 (80, 81), and the European Committee on Antimicrobial Susceptibility Testing (EUCAST) (http://www.eucast.org). The breakpoints for VAN, RIF, ERY, CHL, and meropenem were determined according to a previous study (54, 58). Multidrug resistance (MDR) was defined as resistance to at least three antimicrobial classes. C. difficile ATCC 700057 was included as a control in each experiment (82).
Antimicrobial resistance genes were predicted through comparison with the Antibiotic Resistance Genes Database, ARDB https://ardb.cbcb.umd.edu/ (35), and the Comprehensive Antibiotic Resistance Database, CARD https://card.mcmaster.ca/ (36). Heatmap analysis was performed using the pheatmap package, and statistics packages in R software (version 2.15.3).
Single nucleotide polymorphisms detected and association analysis.
Single nucleotide polymorphisms (SNPs) were detected by comparing the sequence of each isolate with that of strain BJ08. SNP calling was detected using the NUCmer program in the MUMer package (V3.23). All primary SNPs were further evaluated in contigs to validate alleles. Sequence regions of 201 bp, which included 100 bp extracted from either side of the referenced SNP site, were subjected to BLAST analysis with sample contigs. The SNPs were removed if the alignment length was less than 101 bp. In addition, the SNPs located in repeat regions predicted by RepeatMasker (version open-4.05) and TRF (Tandem Repeats Finder; version 4.07b) of the reference were also excluded. Genome-wide association studies (GWAS) were conducted using the open-source R package treeWAS, which is freely available at https://github.com/caitiecollins/treeWAS (83). Three methods (terminal, simultaneous, and subsequent tests) were used to confirm the results in this study.
Data availability.
The sequence information from this whole-genome shotgun project of 37 C. difficile isolates has been submitted to DDBJ/EMBL/GenBank under the following BioProject accession number PRJNA479396.
ACKNOWLEDGMENTS
This research was supported by the National Sci-Tech Key Project (grant 2018ZX10733402).
We thank Lei Zhao from Iowa State University for critically reviewing the manuscript.
Supplemental Material
REFERENCES
1.
He M, Miyajima F, Roberts P, Ellison L, Pickard DJ, Martin MJ, Connor TR, Harris SR, Fairley D, Bamford KB, D'Arc S, Brazier J, Brown D, Coia JE, Douce G, Gerding D, Kim HJ, Koh TH, Kato H, Senoh M, Louie T, Michell S, Butt E, Peacock SJ, Brown NM, Riley T, Songer G, Wilcox M, Pirmohamed M, Kuijper E, Hawkey P, Wren BW, Dougan G, Parkhill J, Lawley TD. 2013. Emergence and global spread of epidemic healthcare-associated Clostridium difficile. Nat Genet 45:109–113.
2.
Warriner K, Xu C, Habash M, Sultan S, Weese SJ. 2017. Dissemination of Clostridium difficile in food and the environment: significant sources of C. difficile community-acquired infection? J Appl Microbiol 122:542–553.
3.
Sebaihia M, Wren BW, Mullany P, Fairweather NF, Minton N, Stabler R, Thomson NR, Roberts AP, Cerdeno-Tarraga AM, Wang H, Holden MT, Wright A, Churcher C, Quail MA, Baker S, Bason N, Brooks K, Chillingworth T, Cronin A, Davis P, Dowd L, Fraser A, Feltwell T, Hance Z, Holroyd S, Jagels K, Moule S, Mungall K, Price C, Rabbinowitsch E, Sharp S, Simmonds M, Stevens K, Unwin L, Whithead S, Dupuy B, Dougan G, Barrell B, Parkhill J. 2006. The multidrug-resistant human pathogen Clostridium difficile has a highly mobile, mosaic genome. Nat Genet 38:779–786.
4.
Monot M, Boursaux-Eude C, Thibonnier M, Vallenet D, Moszer I, Medigue C, Martin-Verstraete I, Dupuy B. 2011. Reannotation of the genome sequence of Clostridium difficile strain 630. J Med Microbiol 60:1193–1199.
5.
Brouwer MS, Allan E, Mullany P, Roberts AP. 2012. Draft genome sequence of the nontoxigenic Clostridium difficile strain CD37. J Bacteriol 194:2125–2126.
6.
Stabler RA, He M, Dawson L, Martin M, Valiente E, Corton C, Lawley TD, Sebaihia M, Quail MA, Rose G, Gerding DN, Gibert M, Popoff MR, Parkhill J, Dougan G, Wren BW. 2009. Comparative genome and phenotypic analysis of Clostridium difficile 027 strains provides insight into the evolution of a hypervirulent bacterium. Genome Biol 10:R102.
7.
He M, Sebaihia M, Lawley TD, Stabler RA, Dawson LF, Martin MJ, Holt KE, Seth-Smith HM, Quail MA, Rance R, Brooks K, Churcher C, Harris D, Bentley SD, Burrows C, Clark L, Corton C, Murray V, Rose G, Thurston S, van Tonder A, Walker D, Wren BW, Dougan G, Parkhill J. 2010. Evolutionary dynamics of Clostridium difficile over short and long time scales. Proc Natl Acad Sci U S A 107:7527–7532.
8.
Knight DR, Elliott B, Chang BJ, Perkins TT, Riley TV. 2015. Diversity and evolution in the genome of Clostridium difficile. Clin Microbiol Rev 28:721–741.
9.
Didelot X, Eyre DW, Cule M, Ip CL, Ansari MA, Griffiths D, Vaughan A, O'Connor L, Golubchik T, Batty EM, Piazza P, Wilson DJ, Bowden R, Donnelly PJ, Dingle KE, Wilcox M, Walker AS, Crook DW, Peto TE, Harding RM. 2012. Microevolutionary analysis of Clostridium difficile genomes to investigate transmission. Genome Biol 13:R118.
10.
Mullany P, Allan E, Roberts AP. 2015. Mobile genetic elements in Clostridium difficile and their role in genome function. Res Microbiol 166:361–367.
11.
Goh S, Hussain H, Chang BJ, Emmett W, Riley TV, Mullany P. 2013. Phage phiC2 mediates transduction of Tn6215, encoding erythromycin resistance, between Clostridium difficile strains. mBio 4:e00840-13.
12.
Brouwer MS, Warburton PJ, Roberts AP, Mullany P, Allan E. 2011. Genetic organisation, mobility and predicted functions of genes on integrated, mobile genetic elements in sequenced strains of Clostridium difficile. PLoS One 6:e23014.
13.
Wasels F, Monot M, Spigaglia P, Barbanti F, Ma L, Bouchier C, Dupuy B, Mastrantonio P. 2014. Inter- and intraspecies transfer of a Clostridium difficile conjugative transposon conferring resistance to MLSB. Microb Drug Resist 20:555–560.
14.
Griffiths D, Fawley W, Kachrimanidou M, Bowden R, Crook DW, Fung R, Golubchik T, Harding RM, Jeffery KJ, Jolley KA, Kirton R, Peto TE, Rees G, Stoesser N, Vaughan A, Walker AS, Young BC, Wilcox M, Dingle KE. 2010. Multilocus sequence typing of Clostridium difficile. J Clin Microbiol 48:770–778.
15.
Eyre DW, Cule ML, Wilson DJ, Griffiths D, Vaughan A, O'Connor L, Ip CLC, Golubchik T, Batty EM, Finney JM, Wyllie DH, Didelot X, Piazza P, Bowden R, Dingle KE, Harding RM, Crook DW, Wilcox MH, Peto TEA, Walker AS. 2013. Diverse sources of C. difficile infection identified on whole-genome sequencing. N Engl J Med 369:1195–1205.
16.
Stabler RA, Dawson LF, Valiente E, Cairns MD, Martin MJ, Donahue EH, Riley TV, Songer JG, Kuijper EJ, Dingle KE, Wren BW. 2012. Macro and micro diversity of Clostridium difficile isolates from diverse sources and geographical locations. PLoS One 7:e31559.
17.
Janezic S, Rupnik M. 2015. Genomic diversity of Clostridium difficile strains. Res Microbiol 166:353–360.
18.
Bauer MP, Notermans DW, van Benthem BH, Brazier JS, Wilcox MH, Rupnik M, Monnet DL, van Dissel JT, Kuijper EJ. 2011. Clostridium difficile infection in Europe: a hospital-based survey. Lancet 377:63–73.
19.
Freeman J, Vernon J, Morris K, Nicholson S, Todhunter S, Longshaw C, Wilcox MH, Pan-European Longitudinal Surveillance of Antibiotic Resistance among Prevalent Clostridium difficile Ribotypes’ Study Group. 2015. Pan-European longitudinal surveillance of antibiotic resistance among prevalent Clostridium difficile ribotypes. Clin Microbiol Infect 21:248.e9–248.e16.
20.
Lim SK, Stuart RL, Mackin KE, Carter GP, Kotsanas D, Francis MJ, Easton M, Dimovski K, Elliott B, Riley TV, Hogg G, Paul E, Korman TM, Seemann T, Stinear TP, Lyras D, Jenkin GA. 2014. Emergence of a ribotype 244 strain of Clostridium difficile associated with severe disease and related to the epidemic ribotype 027 strain. Clin Infect Dis 58:1723–1730.
21.
O'Connor JR, Johnson S, Gerding DN. 2009. Clostridium difficile infection caused by the epidemic BI/NAP1/027 strain. Gastroenterology 136:1913–1924.
22.
Tickler IA, Goering RV, Whitmore JD, Lynn AN, Persing DH, Tenover FC. 2014. Strain types and antimicrobial resistance patterns of Clostridium difficile isolates from the United States, 2011 to 2013. Antimicrob Agents Chemother 58:4214–4218.
23.
Kuijper EJ, Barbut F, Brazier JS, Kleinkauf N, Eckmanns T, Lambert ML, Drudy D, Fitzpatrick F, Wiuff C, Brown DJ, Coia JE, Pituch H, Reichert P, Even J, Mossong J, Widmer AF, Olsen KE, Allerberger F, Notermans DW, Delmee M, Coignard B, Wilcox M, Patel B, Frei R, Nagy E, Bouza E, Marin M, Akerlund T, Virolainen-Julkunen A, Lyytikainen O, Kotila S, Ingebretsen A, Smyth B, Rooney P, Poxton IR, Monnet DL. 2008. Update of Clostridium difficile infection due to PCR ribotype 027 in Europe, 2008. Euro Surveill 13(31):pii=18942. https://doi.org/10.2807/ese.13.31.18942-3en.
24.
Chen R, Feng Y, Wang X, Yang J, Zhang X, Lu X, Zong Z. 2017. Whole genome sequences of three Clade 3 Clostridium difficile strains carrying binary toxin genes in China. Sci Rep 7:43555.
25.
Kuijper EJ, de Weerdt J, Kato H, Kato N, van Dam AP, van der Vorm ER, Weel J, van Rheenen C, Dankert J. 2001. Nosocomial outbreak of Clostridium difficile-associated diarrhoea due to a clindamycin-resistant enterotoxin A-negative strain. Eur J Clin Microbiol Infect Dis 20:528–534.
26.
Drudy D, Harnedy N, Fanning S, Hannan M, Kyne L. 2007. Emergence and control of fluoroquinolone-resistant, toxin A-negative, toxin B-positive Clostridium difficile. Infect Control Hosp Epidemiol 28:932–940.
27.
Alfa MJ, Kabani A, Lyerly D, Moncrief S, Neville LM, Al-Barrak A, Harding GK, Dyck B, Olekson K, Embil JM. 2000. Characterization of a toxin A-negative, toxin B-positive strain of Clostridium difficile responsible for a nosocomial outbreak of Clostridium difficile-associated diarrhea. J Clin Microbiol 38:2706–2714.
28.
Collins DA, Hawkey PM, Riley TV. 2013. Epidemiology of Clostridium difficile infection in Asia. Antimicrob Resist Infect Control 2:21.
29.
Knight DR, Thean S, Putsathit P, Fenwick S, Riley TV. 2013. Cross-sectional study reveals high prevalence of Clostridium difficile non-PCR ribotype 078 strains in Australian veal calves at slaughter. Appl Environ Microbiol 79:2630–2635.
30.
Suo J, Yan Z, Wu Y, Li WG, Zhang WZ, Liu XS, Liu Y, Lu J. 2017. Clostridium difficile RT 078/ST11: a threat to community population and pigs identified in elder hospitalized patients in Beijing, China. Infect Control Hosp Epidemiol 38:1383–1385.
31.
Hallgren A, Saeedi B, Nilsson M, Monstein HJ, Isaksson B, Hanberger H, Nilsson LE. 2003. Genetic relatedness among Enterococcus faecalis with transposon-mediated high-level gentamicin resistance in Swedish intensive care units. J Antimicrob Chemother 52:162–167.
32.
Donabedian SM, Thal LA, Hershberger E, Perri MB, Chow JW, Bartlett P, Jones R, Joyce K, Rossiter S, Gay K, Johnson J, Mackinson C, Debess E, Madden J, Angulo F, Zervos MJ. 2003. Molecular characterization of gentamicin-resistant Enterococci in the United States: evidence of spread from animals to humans through food. J Clin Microbiol 41:1109–1113.
33.
Roberts AP, Allan E, Mullany P. 2014. The impact of horizontal gene transfer on the biology of Clostridium difficile. Adv Microb Physiol 65:63–82.
34.
Meessen-Pinard M, Sekulovic O, Fortier LC. 2012. Evidence of in vivo prophage induction during Clostridium difficile infection. Appl Environ Microbiol 78:7662–7670.
35.
Liu B, Pop M. 2009. ARDB–Antibiotic Resistance Genes Database. Nucleic Acids Res 37:D443–D447.
36.
McArthur AG, Waglechner N, Nizam F, Yan A, Azad MA, Baylay AJ, Bhullar K, Canova MJ, De Pascale G, Ejim L, Kalan L, King AM, Koteva K, Morar M, Mulvey MR, O'Brien JS, Pawlowski AC, Piddock LJ, Spanogiannopoulos P, Sutherland AD, Tang I, Taylor PL, Thaker M, Wang W, Yan M, Yu T, Wright GD. 2013. The comprehensive antibiotic resistance database. Antimicrob Agents Chemother 57:3348–3357.
37.
Liu XS, Li WG, Zhang WZ, Wu Y, Lu JX. 2018. Molecular characterization of Clostridium difficile isolates in China from 2010 to 2015. Front Microbiol 9:845.
38.
Kao SJ, You I, Clewell DB, Donabedian SM, Zervos MJ, Petrin J, Shaw KJ, Chow JW. 2000. Detection of the high-level aminoglycoside resistance gene aph(2”)-Ib in Enterococcus faecium. Antimicrob Agents Chemother 44:2876–2879.
39.
Reference deleted.
40.
Stern A, Mick E, Tirosh I, Sagy O, Sorek R. 2012. CRISPR targeting reveals a reservoir of common phages associated with the human gut microbiome. Genome Res 22:1985–1994.
41.
Makarova KS, Haft DH, Barrangou R, Brouns SJ, Charpentier E, Horvath P, Moineau S, Mojica FJ, Wolf YI, Yakunin AF, van der Oost J, Koonin EV. 2011. Evolution and classification of the CRISPR-Cas systems. Nat Rev Microbiol 9:467–477.
42.
Makarova KS, Wolf YI, Alkhnbashi OS, Costa F, Shah SA, Saunders SJ, Barrangou R, Brouns SJ, Charpentier E, Haft DH, Horvath P, Moineau S, Mojica FJ, Terns RM, Terns MP, White MF, Yakunin AF, Garrett RA, van der Oost J, Backofen R, Koonin EV. 2015. An updated evolutionary classification of CRISPR-Cas systems. Nat Rev Microbiol 13:722–736.
43.
Boudry P, Semenova E, Monot M, Datsenko KA, Lopatina A, Sekulovic O, Ospina-Bedoya M, Fortier LC, Severinov K, Dupuy B, Soutourina O. 2015. Function of the CRISPR-Cas system of the human pathogen Clostridium difficile. mBio 6:e01112-15.
44.
Andersen JM, Shoup M, Robinson C, Britton R, Olsen KE, Barrangou R. 2016. CRISPR diversity and microevolution in Clostridium difficile. Genome Biol Evol 8:2841–2855.
45.
Grissa I, Vergnaud G, Pourcel C. 2007. The CRISPRdb database and tools to display CRISPRs and to generate dictionaries of spacers and repeats. BMC Bioinformatics 8:172.
46.
Barros MP, Franca CT, Lins RH, Santos MD, Silva EJ, Oliveira MB, Silveira-Filho VM, Rezende AM, Balbino VQ, Leal-Balbino TC. 2014. Dynamics of CRISPR loci in microevolutionary process of Yersinia pestis strains. PLoS One 9:e108353.
47.
Pettengill JB, Timme RE, Barrangou R, Toro M, Allard MW, Strain E, Musser SM, Brown EW. 2014. The evolutionary history and diagnostic utility of the CRISPR-Cas system within Salmonella enterica ssp. enterica. PeerJ 2:e340.
48.
Palmer KL, Gilmore MS. 2010. Multidrug-resistant enterococci lack CRISPR-cas. mBio 1:e00227-10.
49.
Nozawa T, Furukawa N, Aikawa C, Watanabe T, Haobam B, Kurokawa K, Maruyama F, Nakagawa I. 2011. CRISPR inhibition of prophage acquisition in Streptococcus pyogenes. PLoS One 6:e19543.
50.
Louwen R, Staals RH, Endtz HP, van Baarlen P, van der Oost J. 2014. The role of CRISPR-Cas systems in virulence of pathogenic bacteria. Microbiol Mol Biol Rev 78:74–88.
51.
Freeman J, Vernon J, Pilling S, Morris K, Nicholson S, Shearman S, Longshaw C, Wilcox MH. 2018. The ClosER study: results from a three-year pan-European longitudinal surveillance of antibiotic resistance among prevalent Clostridium difficile ribotypes, 2011-2014. Clin Microbiol Infect 24:724–731.
52.
Tang C, Cui L, Xu Y, Xie L, Sun P, Liu C, Xia W, Liu G. 2016. The incidence and drug resistance of Clostridium difficile infection in Mainland China: a systematic review and meta-analysis. Sci Rep 6:37865.
53.
Cheng JW, Xiao M, Kudinha T, Kong F, Xu ZP, Sun LY, Zhang L, Fan X, Xie XL, Xu YC. 2016. Molecular epidemiology and antimicrobial susceptibility of Clostridium difficile isolates from a university teaching hospital in China. Front Microbiol 7:1621.
54.
Jin D, Luo Y, Huang C, Cai J, Ye J, Zheng Y, Wang L, Zhao P, Liu A, Fang W, Wang X, Xia S, Jiang J, Tang YW. 2017. Molecular epidemiology of Clostridium difficile infection in hospitalized patients in eastern China. J Clin Microbiol 55:801–810.
55.
Chen YB, Gu SL, Shen P, Lv T, Fang YH, Tang LL, Li LJ. 2018. Molecular epidemiology and antimicrobial susceptibility of Clostridium difficile isolated from hospitals during a 4-year period in China. J Med Microbiol 67:52–59.
56.
Peng Z, Jin D, Kim HB, Stratton CW, Wu B, Tang YW, Sun X. 2017. Update on antimicrobial resistance in Clostridium difficile: resistance mechanisms and antimicrobial susceptibility testing. J Clin Microbiol 55:1998–2008.
57.
Dridi L, Tankovic J, Burghoffer B, Barbut F, Petit JC. 2002. gyrA and gyrB mutations are implicated in cross-resistance to ciprofloxacin and moxifloxacin in Clostridium difficile. Antimicrob Agents Chemother 46:3418–3421.
58.
Spigaglia P, Barbanti F, Mastrantonio P. 2011. Multidrug resistance in European Clostridium difficile clinical isolates. J Antimicrob Chemother 66:2227–2234.
59.
Caspers P, Locher HH, Pfaff P, Diggelmann S, Rueedi G, Bur D, Ritz D. 2017. Different resistance mechanisms for cadazolid and linezolid in Clostridium difficile found by whole-genome sequencing analysis. Antimicrob Agents Chemother 61:e00384-17.
60.
Gao Q, Wu S, Huang H, Ni Y, Chen Y, Hu Y, Yu Y. 2016. Toxin profiles, PCR ribotypes and resistance patterns of Clostridium difficile: a multicentre study in China, 2012-2013. Int J Antimicrob Agents 48:736–739.
61.
Peláez T, Cercenado E, Alcalá L, Marín M, Martín-López A, Martínez-Alarcón J, Catalán P, Sánchez-Somolinos M, Bouza E. 2008. Metronidazole resistance in Clostridium difficile is heterogeneous. J Clin Microbiol 46:3028–3032.
62.
Tsvetkova K, Marvaud JC, Lambert T. 2010. Analysis of the mobilization functions of the vancomycin resistance transposon Tn1549, a member of a new family of conjugative elements. J Bacteriol 192:702–713.
63.
Knight DR, Androga GO, Ballard SA, Howden BP, Riley TV. 2016. A phenotypically silent vanB2 operon carried on a Tn1549-like element in Clostridium difficile. mSphere 1:e00177-16.
64.
Liao F, Li W, Gu W, Zhang W, Liu X, Fu X, Xu W, Wu Y, Lu J. 2018. A retrospective study of community-acquired Clostridium difficile infection in southwest China. Sci Rep 8:3992.
65.
Li R, Zhu H, Ruan J, Qian W, Fang X, Shi Z, Li Y, Li S, Shan G, Kristiansen K, Li S, Yang H, Wang J, Wang J. 2010. De novo assembly of human genomes with massively parallel short read sequencing. Genome Res 20:265–272.
66.
Li R, Li Y, Kristiansen K, Wang J. 2008. SOAP: short oligonucleotide alignment program. Bioinformatics 24:713–714.
67.
Besemer J, Lomsadze A, Borodovsky M. 2001. GeneMarkS: a self-training method for prediction of gene starts in microbial genomes. Implications for finding sequence motifs in regulatory regions. Nucleic Acids Res 29:2607–2618.
68.
Ashburner M, Ball CA, Blake JA, Botstein D, Butler H, Cherry JM, Davis AP, Dolinski K, Dwight SS, Eppig JT, Harris MA, Hill DP, Issel-Tarver L, Kasarskis A, Lewis S, Matese JC, Richardson JE, Ringwald M, Rubin GM, Sherlock G. 2000. Gene Ontology: tool for the unification of biology. The Gene Ontology Consortium. Nat Genet 25:25–29.
69.
Kanehisa M, Goto S, Hattori M, Aoki-Kinoshita KF, Itoh M, Kawashima S, Katayama T, Araki M, Hirakawa M. 2006. From genomics to chemical genomics: new developments in KEGG. Nucleic Acids Res 34:D354–D357.
70.
Tatusov RL, Fedorova ND, Jackson JD, Jacobs AR, Kiryutin B, Koonin EV, Krylov DM, Mazumder R, Mekhedov SL, Nikolskaya AN, Rao BS, Smirnov S, Sverdlov AV, Vasudevan S, Wolf YI, Yin JJ, Natale DA. 2003. The COG database: an updated version includes eukaryotes. BMC Bioinformatics 4:41.
71.
Li W, Jaroszewski L, Godzik A. 2002. Tolerating some redundancy significantly speeds up clustering of large protein databases. Bioinformatics 18:77–82.
72.
Saier MH, Jr, Reddy VS, Tamang DG, Vastermark A. 2014. The Transporter Classification Database. Nucleic Acids Res 42:D251–D258.
73.
Bairoch A, Apweiler R. 2000. The SWISS-PROT protein sequence database and its supplement TrEMBL in 2000. Nucleic Acids Res 28:45–48.
74.
Lowe TM, Eddy SR. 1997. tRNAscan-SE: a program for improved detection of transfer RNA genes in genomic sequence. Nucleic Acids Res 25:955–964.
75.
Lagesen K, Hallin P, Rodland EA, Staerfeldt HH, Rognes T, Ussery DW. 2007. RNAmmer: consistent and rapid annotation of ribosomal RNA genes. Nucleic Acids Res 35:3100–3108.
76.
Gardner PP, Daub J, Tate JG, Nawrocki EP, Kolbe DL, Lindgreen S, Wilkinson AC, Finn RD, Griffiths-Jones S, Eddy SR, Bateman A. 2009. Rfam: updates to the RNA families database. Nucleic Acids Res 37:D136–D140.
77.
Tamura K, Stecher G, Peterson D, Filipski A, Kumar S. 2013. MEGA6: Molecular Evolutionary Genetics Analysis version 6.0. Mol Biol Evol 30:2725–2729.
78.
Arndt D, Grant JR, Marcu A, Sajed T, Pon A, Liang Y, Wishart DS. 2016. PHASTER: a better, faster version of the PHAST phage search tool. Nucleic Acids Res 44:W16–W21.
79.
Grissa I, Vergnaud G, Pourcel C. 2007. CRISPRFinder: a web tool to identify clustered regularly interspaced short palindromic repeats. Nucleic Acids Res 35:W52–W57.
80.
Clinical and Laboratory Standards Institute. 2014. Performance standards for antimicrobial susceptibility testing. Twenty-fourth informational supplement. CLSI M 100-S24. Clinical and Laboratory Standards Institute, Wayne, PA.
81.
Clinical and Laboratory Standards Institute. 2007. Methods of antimicrobial susceptibility testing of anaerobic bacteria. Approved standard M11-A7, 7th ed. Clinical and Laboratory Standards Institute, Wayne, PA.
82.
Magiorakos AP, Srinivasan A, Carey RB, Carmeli Y, Falagas ME, Giske CG, Harbarth S, Hindler JF, Kahlmeter G, Olsson-Liljequist B, Paterson DL, Rice LB, Stelling J, Struelens MJ, Vatopoulos A, Weber JT, Monnet DL. 2012. Multidrug-resistant, extensively drug-resistant and pandrug-resistant bacteria: an international expert proposal for interim standard definitions for acquired resistance. Clin Microbiol Infect 18:268–281.
83.
Collins C, Didelot X. 2018. A phylogenetic method to perform genome-wide association studies in microbes that accounts for population structure and recombination. PLoS Comput Biol 14:e1005958.
Information & Contributors
Information
Published In
mSystems
Volume 4 • Number 2 • 30 April 2019
eLocator: 10.1128/msystems.00252-18
Editor: Sergio Baranzini, University of California, San Francisco
PubMed: 30944881
Copyright
© 2019 Wu et al. This is an open-access article distributed under the terms of the Creative Commons Attribution 4.0 International license.
History
Received: 11 October 2018
Accepted: 18 January 2019
Published online: 26 March 2019
Keywords
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Editor
Sergio Baranzini
Editor
University of California, San Francisco
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Citation
2019.
Independent Microevolution Mediated by Mobile Genetic Elements of Individual Clostridium difficile Isolates from Clade 4 Revealed by Whole-Genome Sequencing. mSystems
4:10.1128/msystems.00252-18.
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