Skip to main content
NCBI home page
Saudi Journal of Biological Sciences logoLink to Saudi Journal of Biological Sciences
. 2021 Apr 23;28(8):4472–4477. doi: 10.1016/j.sjbs.2021.04.044

Diagnostic deficiencies of C. difficile infection among patients in a tertiary hospital in Saudi Arabia: A laboratory-based case series

Reem AlJindan a, Doaa M AlEraky b, J Francis Borgio c,d, Sayed AbdulAzeez c, Baha Abdalhamid e, Nehal Mahmoud a, Maha Farhat f,⁎
PMCID: PMC8324924  PMID: 34354432

Abstract

Clostridioides difficile infection (CDI) has become a threatening public health problem in the developed world. In the kingdom of Saudi Arabia, prevalence of CDI is still unknown due to limited surveillance protocols and diagnostic resources. We used a two-step procedure to study and confirm C. difficile cases. We also studied toxin profiles of these isolates.

Stool samples were collected from symptomatic patients and clinically suspected of CDI for almost 12 months. Isolates were confirmed by culture method followed by 16S rRNA sequencing. Multiplex PCR was performed for the identification of toxin A, toxin B and binary toxin genes and compared to Gene Expert results.

Out of the 47 collected samples, 27 were successfully grown on culture media. 18 samples were confirmed as C. difficile by both culture and 16S rRNA sequencing. Interestingly, the rest of the isolates (9 species) belonged to different genera. Our results showed 95% of samples were positive for both toxin A and B (tcdA, tcdB) and all samples exhibited the toxin gene regulator tcdC. All samples were confirmed negative for the binary toxin gene ctdB and 11% of the isolates were positive for ctdA gene. Interestingly, one isolate harbored the binary toxin gene (cdtA+) and tested negative for both toxins A and B.

We believe that combining the standard culture method with molecular techniques can make the detection of C. difficile more accurate.

Keywords: Clostridioides difficile, CCFA medium, 16S rRNA sequencing, Toxin A, Toxin B, Binary toxin

1. Introduction

Clostridioides difficile is a known pathogen discovered in 1935 and recognized as a major cause of antibiotic-associated diarrhea (AAD) and Pseudomembranous colitis in the 1970s (Heinlen and Ballard, 2010, Lessa et al., 2012, Oka et al., 2012, Walter et al., 2014). It is a Gram positive strictly anaerobic, spore-forming bacillus that can live in the gastrointestinal tract of humans or animals with thousands of different microbial species and then shed spores into their feces. These spores (metabolically inactive) can be spread to humans through hands, water and food. C. difficile exists in toxigenic form where pathogenicity can be mainly caused by two virulent genes TcdA and TcdB producing toxin A and toxin B respectively. A third and binary toxin has been shown to enhance C. difficile’s virulence and cause a variety of cytotoxic mechanism leading to cell death (Schwan et al., 2009, Pruitt and Lacy, 2012). Other nontoxic virulent factors such as Putative type IV pilus, capsule and flagella play a role in bacterium pathogenesis (Carey-Ann and Carroll, 2013).

Clostridioides difficile-associated Disease (CDAD) appear mostly in old, hospitalized patients (more than 65 years old) with diminished immune response and recent antibiotic exposure associated with treatment failure (Lessa et al., 2012, Burke and Lamont, 2014; (CDC, 2021). In the last several years, the incidence and severity of CDI has been remarkably raised as one of the most common hospitals acquired infection (Peery et al., 2012, Wiegand et al., 2012, Hensgens et al., 2013, Aldeyab et al., 2014). This infection has become a threatening public health problem in the developed world, with significant high rate of morbidity and mortality reported since the early 2000 s. The centers for disease control and prevention (CDC) declared 223,900 cases of CDI in the united states in 2017 of which 12,800 patients died (Centers for Disease Control and Prevention (U.S.), 2019). European surveillance of CDI started in 2016 with 7711 cases including 5756 (74.6%) healthcare-associated cases (European Centre for Disease Prevention and Control, 2016).

Studies suggested similar rates of CDI in Asia compared to Europe and North America with estimated prevalence in the middle east around 11% (Borren et al., 2017). Although the increasing number of CDI reported worldwide, only few studies has been done in developing countries to evaluate the extent of this infection and the problem may be still underestimated (Cheng et al., 2016a, Cheng et al., 2016b, Curcio et al., 2019).

In the Eastern Province of Saudi Arabia, there is a growing concern on the C. difficile infections in hospitals (Hudhaiah and Elhadi, 2019, Al-Tawfiq et al., 2020). C. difficile toxin was detected in 9.5% of patients diagnosed with gastroenteritis at a major referral center in Saudi Arabia in 1994 (Akhter et al., 1994). A more recent study done in 2007–2008 estimated the annual incidence rate of CDI to be around 2.4 and 1.7 per 10,000 patient days respectively (Al-Tawfiq and Abed, 2010). However, prevalence of CDI in the Kingdom of Saudi Arabia is still unknown where difficulties in estimating and comparing the incidence of CDI to the international rates comes from the inconsistent diagnostic, limited surveillance protocols and diagnostic resources.

The techniques that are most often used to detect C. difficile include culture methods, enzyme immunoassay (EIA) and nucleic acid amplification tests (NAATs) (Xiao et al., 2020). PCR is known to have higher sensitivity for toxin genes testing than the toxin EIA routinely used (de Jong et al., 2012). Establishing a routine detection method that is sensitive and fast is essential for the clinical management of patients with CDI (Tenover et al., 2012). The call for the current study was recommended by an earlier study which reported in a recent study last year investigates the prevalence and genotype of nosocomial infection of C. difficile in Eastern Province as there is lack and urgent need to develop a diagnostic protocol for rapid identification of virulent strains of C. difficile (Al-Tawfiq et al., 2020).

Since a fast and accurate laboratory diagnostic is crucial to evaluate the extent of CDI problem in developing countries, we used a two-step procedure to study and confirm C. difficile isolates. The culture method was used followed by 16S rRNA sequencing. Molecular detection was also performed for the identification of toxin A, toxin B and binary toxin genes.

2. Materials and methods

2.1. Sample collection

Patients recruited for the study have been consented as per the Institutional Review Board (IRB) regulations and the proposed project has been submitted for the ethical approval by the ethical committee of Imam Abdulrahman Bin Faisal University (IRB-2017–338-Dent) and all procedures performed with the 1964 Helsinki declaration and its later amendments or comparable ethical standards. Stool samples were collected from patients exhibiting diarrhea and clinically suspected of C. difficile infection from Feb 2017 till May 2018.

All patients enrolled in the study had a history of recent antibiotic exposure, the majority of the samples isolated from male patients (88%) and the average age was 54.8 years. Patients showed different diagnosis such as, chronic diarrhea, ulcerative colitis, fistula of intestine, decompensated liver cirrhosis, urosepsis, acute pancreatitis, recurrent urinary tract infection, end stage renal failure on peritoneal dialysis and cauda equina syndrome.

2.2. C. Difficile isolates toxigenic culture study and bacterial culture

Frozen stool samples have been received from King Fahad Hospital of the University (Imam Abdulrahman Bin Faisal University, Dammam), symptomatic patients who were positive for both toxin A and toxin B by GeneXpert were proceeded for bacterial culture. Identification of C. difficile was performed by boiling fecal samples in PBS (phosphate-buffered saline) with pH 7.4, then cultured on the selective medium, cycloserine cefoxitin fructose agar medium (CCFA) in 86% N2, 7% H2 and 7% CO2 atmosphere at 37℃ for 48 h.

Preliminary identification of C. difficile growth showed shiny, grayish, and flat colonies on the selective medium under anaerobic conditions with a horse manure odor and microscopically gram-positive rods.

Isolated pure colonies were prepared for DNA extraction. The DNA was extracted right away from the pure colonies.

2.3. Genetic study

2.3.1. Confirmation of C. Difficile isolates by 16S rRNA sequencing

DNA was extracted using Gentra Puregene Yeast/Bact. Kit (Qiagen, Hilden, Germany) according to the manufacturer protocol. The extracted DNA was washed and purified three times before the final elution in 100 μl of elution buffer. DNA concentrations were measured by spectrophotometer (nanodrop 2000) and the extracted DNA was stored at −80 °C for further molecular analysis. In order to confirm C. difficile isolates, DNA from all the individual colonies were PCR amplified for 16S rRNA gene, sequenced and analysed as we described earlier (Rehman et al., 2019).

Nucleotide sequence accession numbers. 16S rRNA sequences of C. difficile and other detected bacteria were submitted to the GenBank database under the following accession numbers; MK909920-MK909925, MN080438, MN053902, MN049824, MN049823, MN049821, MN049793, MN049583, MN049570, MN049549, MN049543, MK791717, MK791684.

2.3.2. Virulence genes identification using multiplex PCR

A multiplex PCR was established using single tube PCR reaction for the identification of the virulent genes. A total of 12 primers were used for the detection of 16SrDNA, tcdA, tcdb, ctdB, ctdA and tcdC genes with amplicon size 1062, 629, 410, 262, 221 and 475 bp respectively (Table 1). Except tcdC gene, all the other genes were identified using single tube multiplex PCR (Persson et al., 2008).

Table 1.

List of primers for the Clostridioides difficile Molecular characterization.

Gene target Primer name
Sequence (5′–3′) 		Primer concentration (μM) Amplicon size (bp)
5-plex PCR
tcdA tcdA-F3345
GCATGATAAGGCAACTTCAGTGGTAa 		0.6 629
tcdA-R3969
AGTTCCTCCTGCTCCATCAAATG		 0.6
tcdB tcdB-F5670
CCAAARTGGAGTGTTACAAACAGGTG		 0.4 410
tcdB-R6079A
GCATTTCTCCATTCTCAGCAAAGTA		 0.2
cdtA cdtA-F739A
GGGAAGCACTATATTAAAGCAGAAGC		 0.05 221
cdtA-F739B
GGGAAACATTATATTAAAGCAGAAGC		 0.05
ctdB ctdB-F617
TTGACCCAAAGTTGATGTCTGATTG		 0.1 262
cdtB-R878
CGGATCTCTTGCTTCAGTCTTTATAG		 0.1
16S rDNA PS13
GGAGGCAGCAGTGGGGAATA		 0.05 1062
PS14
TGACGGGCGGTGTGTACAAG		 0.05
tcdC analysis
tcdC tcdC-F(−17)
AAAAGGGAGATTGTATTATGTTTTC		 0.2 475c
tcdC-R(+462)
CAATAACTTGAATAACCTTACCTTCA		 0.2
Open in a new tab

The Taq polymerase mediated amplifications were carried out in total volumes of 25 Âµl with the following recipe: 1 × PCR buffer, 2.6 mM MgCl2, 260 µM each of dNTP and 1.25 U of Taq polymerase (MoleQule-ON, New Zealand), and each primer (Table 1). Thermocycler profile: 10 min at 94℃; 35 cycles of 94℃ for 50 s, 54℃ for 40 s and at 72℃ for 50 s, and a final extension at 72℃ for 3 min. Similar protocol was applied for a single amplicon for PCR of tcdC (475 bp) gene with 2 primers.

3. Results

3.1. Confirmation of C. difficile by 16S rRNA sequencing

A total of 47 samples were collected over a period of twelve months. Those samples were positive by GeneXpert for both toxin A and toxin B. However, only 27 samples were successfully grown on the selective media. A total of 18 sequences were related to Clostridioides difficile strains using 16S rRNA and 9 samples belonged to other bacterial strains as follow; Terrisporobacter sp. (MK793698), Enterococcus faecalis (MK791713, MK791687, MK791686), Terrisporobacter petrolearius (MN049790), Clostridium perfringens (MN049820), Clostridium ventriculi (MN080440), Erysipelatoclostridium ramosum (MN049572) and Bacillus sp.

Eleven antimicrobial agents have been associated with C. difficile infection, ciprofloxacin followed by ceftriaxone, tazocin and gentamicin were the most prevalent predisposing antimicrobial agents.

3.2. Virulence genes identification using multiplex PCR

The results of multiplex PCR analysis showed different grouping of samples (Table 2, Fig. 1). All samples were revealed toxigenic where the majority tested positive for the two toxin genes tcdA, tcdB accounting for 89% of the strains with tcdC+. The two remaining strains (MD2505, MD1742) harbored the binary toxin gene cdtA accounting for 11% of the strains. These two latter strains had different toxin profiles of (tcdA + B + C+, cdtA+) and (cdtA+, tcdC+) respectively (see Fig. 2).

Table 2.

Identification of toxin genes in Clostridioides difficile using multiplex PCR.

Sample No. 16S rDNA tcdA tcdB ctdB cdtA tcdC
MD837 (+) (+) (+) (−) (−) (+)
MD500 (+) (+) (+) (−) (−) (+)
MD895 (+) (+) (+) (−) (−) (+)
MD949 (+) (+) (+) (−) (−) (+)
MD1400 (+) (+) (+) (−) (−) (+)
CDT25 (+) (+) (+) (−) (−) (+)
MD814 (+) (+) (+) (−) (−) (+)
MD991 (+) (+) (+) (−) (−) (+)
MD1346 (+) (+) (+) (−) (−) (+)
MD2136 (+) (+) (+) (−) (−) (+)
MD1904 (+) (+) (+) (−) (−) (+)
CDT18 (+) (+) (+) (−) (−) (+)
95,731 (+) (+) (+) (−) (−) (+)
6603 (+) (+) (+) (−) (−) (+)
MRN121781 (+) (+) (+) (−) (−) (+)
MRN66043 (+) (+) (+) (−) (−) (+)
MD2505 (+) (+) (+) (−) (+) (+)
MD1742 (−) (−) (−) (−) (+) (+)
Open in a new tab

(+): indicates positive for the corresponding toxin gene, (−): indicates negative for the corresponding toxin gene.

Fig. 1.

Fig. 1

Open in a new tab

Representative gel pictures of PCR amplicons of toxic genes in Clostridioides difficile. A: Multiplex PCR for 16Sr DNA (1062 bp) tcdA (629 bp) tcdB (410 bp) ctdB (262 bp) and cdtA (221 bp) genes using the following primers: (PS13, PS14) - (F3345, R3989) – (F5670, R6079A, R6079B) - (F617, R878) - (F739A, F739B, R958), respectively. L: 100 bp ladder; Lane 1 to 18: C. difficile strains; Lane 19: Internal C. difficile control. B: Representatives of single amplicon for PCR of tcdC (475 bp) gene by F (17), R (+462) primers. L: 100 bp ladder; Lane 1 to 13: C. difficile strains; Lane 14 and 15: Enterococcus faecalis; Lane 16: Internal C. difficile control. Lane 17: Negative PCR control.

Fig. 2.

Fig. 2

Open in a new tab

Representative gel pictures of PCR amplicons of toxic genes in Clostridioides difficile. A: Multiplex PCR for 16Sr DNA (1062 bp) tcdA (629 bp) tcdB (410 bp) ctdB (262 bp) and cdtA (221 bp) genes using the following primers; (PS13, PS14) - (F3345, R3989) – (F5670, R6079A, R6079B) - (F617, R878) - (F739A, F739B, R958), respectively. B: Single amplicon for PCR of tcdC (475 bp) gene by F (17), R (+462) primers.

4. Discussion

In this research work, we investigated a group of 47 samples that were collected from February 2017 for a period of one year in the Eastern Province of Saudi Arabia. We used a two-step test to isolate C. difficile from stool samples; culture followed by 16S rRNA sequencing. We also studied toxin profiles of these isolates. Only few studies were done in this field in our region despite the high mortality rates and financial burden encountered worldwide due to hospital-acquired infections including CDI (Al-Tawfiq et al., 2020, Alasmari et al., 2014, Aldeyab et al., 2014, Shajan et al., 2014, Alzouby et al., 2020). Our study had limitations where only 47 samples were studied. However, this small sample size does not invalidate our interesting results as discussed below.

Samples were taken from patients exhibiting diarrhea and clinically suspected of C. difficile infection. Detection of Clostridioides toxin was carried out first as part of routine clinical investigations using GeneXpert assay to test both toxin A and toxin B. Then, culture-based detection was conducted to isolate Clostridioides from stool samples on the selective media and under the appropriate growth requirements. This standard method showed more sensitivity than PCR and EIA tests used in the few studies done in our region (Al-Tawfiq et al., 2020). These two latter techniques have shown low sensitivity to detect low concentrations of Clostridioides in the medium studied (Lessa et al., 2012, Cadnum et al., 2014, Fang et al., 2017). 16S rRNA sequencing was used to confirm our culture results and only 18 samples were confirmed as C. difficile out of the 27 isolates grown successfully on the culture medium. The rest of 9 isolates belonged to other bacterial strains as showed in the results.

Terrisporobacter glycolicus and Erysipelatoclostridium ramosum are part of Clostridioides cluster XI and XVIII clostridial cluster, respectively. Terrisporobacter glycolicus was initially classified as Clostridium glycolicum and it is placed within the family Peptostreptococcaceae along with Clostridioides difficile (Cheng et al., 2016a, Ohashi and Fujisawa, 2019). Erysipelatoclostridium ramosum belongs to the human gut microbiota. It is a Gram-positive anaerobic enteric bacterium most widely known as Clostridioides ramosum (Zakham et al., 2019). Hence, the growth of these two bacteria on the culture medium and their disapproval as C. difficile isolates by 16S rRNA sequencing describes the requirement of large scale studies on patients with CDI to differentiate the pathogens and to develop the most appropriate diagnostic tools and treatment strategies.

Enterococcus faecalis is an enterococcus, a nosocomial pathogen that has similar risk factors as C. difficile including antibiotic treatment and hospitalization (Özsoy and İlki, 2017). This Vancomycin-Resistant Enterococcus faecalis (VRE) is often detected in the C. difficile toxin positive samples (Rodriguez et al., 2016). Therefore, based on the above results comparing two different methods for detecting C. difficile, we are reporting herein the recovery of nine bacterial species belonging to different genera on the C. difficile culture medium. Despite the negative impact that wrong diagnosis might have putting the patients at risk, delaying the treatment and transmitting the infection, few studies have shed the light on this area in the developing countries leading to the underestimation of CDI.

The 18 samples confirmed as C. difficile by both culture and 16S rRNA sequencing were also subject to toxinotyping by GeneXpert test and Multiplex PCR to identify the virulence genes tcdABC and ctdAB. The predisposing antimicrobial agents that showed the highest association with C. difficile infection were ciprofloxacin and tazocin.

These findings coincide with the meta-analytic study by Brown el al who noted that tetracyclines and penicillin were associated with the lowest risk, while fluoroquinolones, clindamycin, and expanded spectrum cephalosporins were associated with the highest risk of CDI. It also concurs with that literature reported that the excessive use of cephalosporins is related to the occurrence of CDI than other antibiotics (Brown et al., 2013, Czepiel et al., 2019). In this current study, all patients with positive Gene Expert test were having symptoms, as this expensive test is not performed for screening. According to the recommendations for the first line of treatment, both metronidazole and vancomycin were used and patients with asymptomatic colonization should not be treated (McDonald et al., 2018). The laboratory investigations to identify C. difficile are still critical (Carey-Ann and Carroll, 2013).

Our results showed that GeneXpert test is a reliable diagnostic test to identify the toxins, yet it can be associated with other bacteria. The major virulence factors of C. difficile are toxins A and B but, the presence of binary toxin genes is significantly related to the threat of mortality (Berry et al., 2017). Although the role of binary toxin genes has been shown to cause severe pathogenicity, the role of this toxin remains unclear. Our results showed 95% of samples were positive for both toxin A and B (tcdA, tcdb) and all samples exhibited the toxin gene regulator tcdC. Recent studies reported that tcdB could provoke the characteristics of CDI without tcdA and consequently, they are focusing on the toxin-mediated treatments to broaden the options for eliminating the risk of CDI (Chandrasekaran and Lacy, 2017, Czepiel et al., 2019). All samples were confirmed negative for the binary toxin gene ctdB and 11% of the isolates were positive for ctdA gene.

Many studies have used NAAT to study CDI and many of them reported false positive results for colonized patients without disease (Fang et al., 2017). Interestingly, one of the samples (MD1742, Table 2) that has been confirmed as C. difficile by 16S rRNA sequencing was negative for 16Sr DNA gene testing included in the multiplex PCR. The primers sequences were examined and compared to the sequence of the C. difficile obtained by 16S rRNA sequencing. Only forward primer sequence matched with the bacterium sequence, while no match was found with the reverse primer region. We suggest that this can be the result of a mutation in the region corresponding to the reverse primer sequence and consequently led to the false negative results in the multiplex PCR. This same isolate (MD1742) harbored the binary toxin gene (cdtA+) and tested negative for both toxins A and B. However, most of the binary toxin-producing C. difficile strains reported to date also produced TcdA and/or TcdB.

To conclude, our study has many salient features which are crucial to explore this life threating infection in the Eastern province. The present study sheds the light on the importance of the methods of detection in the C. difficile study. Improper detection leads to wrong diagnosis, treatments delay and ineffective infection control measures. We believe that combining the standard culture method with 16SrRNA sequencing can make the detection of C. difficile more accurate. In addition, our results confirm the presence of locus of pathogenicity with 5 toxin genes in C. difficile isolates from the patients. The locus of pathogenicity in the isolates clearly signifies the multidrug resistance of the isolates and the clinical implication of the patients. These observations strongly suggest that nation-wide large-scale studies are necessary on the locus of pathogenicity and clinical severity for the C. difficile infection to identify the most appropriate drug targets and to design more sensitive diagnostics tools and appropriate preventive measures.

Declaration of Competing Interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Acknowledgements

This work was carried out under Imam AbdulRahman Bin Faisal University Research Grant 2017-256-Dent and approval of Institutional Review Board (IRB-2017‐338-Dent). The authors acknowledge the University for funding, the technical staff (Mr. Ranilo, M. Tumbaga) at the microbiology laboratory and at the Institute for Research and Medical Consultations for their technical support, and the student Nora Hadi for her helpful assistance in the literature review.

Footnotes

Peer review under responsibility of King Saud University.

Contributor Information

Reem AlJindan, Email: raljindan@iau.edu.sa.

Doaa M AlEraky, Email: dmaleraky@iau.edu.sa.

J. Francis Borgio, Email: fbalexander@iau.edu.sa.

Sayed AbdulAzeez, Email: asayed@iau.edu.sa.

Baha Abdalhamid, Email: babdalhamid@unmc.edu.

Nehal Mahmoud, Email: nmhosin@iau.edu.sa.

Maha Farhat, Email: mFarhat@iau.edu.sa.

References

  1. Akhter J., Burdette J.M., Qadri S.M., Myint S.H. Aetiology of gastroenteritis at a major referral centre in Saudi Arabia. J. Int. Med. Res. 1994;22:47–54. doi: 10.1177/030006059402200106. [DOI] [PubMed] [Google Scholar]
  2. Alasmari F., Seiler S.M., Hink T., Burnham C.-A.-D., Dubberke E.R. Prevalence and risk factors for asymptomatic Clostridium difficile carriage. Clin. Infect. Dis. 2014;59:216–222. doi: 10.1093/cid/ciu258. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Aldeyab M.A., Cliffe S., Scott M., Flanagan P., Kearney M., McElnay J., Aldiab M. Risk factors associated with Clostridium difficile infection severity in hospitalized patients. Am. J. Infect. Control. 2014;42:689–690. doi: 10.1016/j.ajic.2014.02.023. [DOI] [PubMed] [Google Scholar]
  4. Al-Tawfiq J.A., Abed M.S. Clostridium difficile-associated disease among patients in Dhahran, Saudi Arabia. Travel Med. Infect. Dis. 2010;8:373–376. doi: 10.1016/j.tmaid.2010.10.003. [DOI] [PubMed] [Google Scholar]
  5. Al-Tawfiq J.A., Rabaan A.A., Bazzi A.M., Raza S., Noureen M. Clostridioides (Clostridium) difficile-associated disease: Epidemiology among patients in a general hospital in Saudi Arabia. Am. J. Infect. Control. 2020;48:1152–1157. doi: 10.1016/j.ajic.2020.01.011. [DOI] [PubMed] [Google Scholar]
  6. Alzouby S., Baig K., Alrabiah F., Shibl A., Al-Nakhli D., Senok A.C. Clostridioides difficile infection: Incidence and risk factors in a tertiary care facility in Riyadh, Saudi Arabia. J Infect Public Health. 2020;13:1012–1017. doi: 10.1016/j.jiph.2019.10.014. [DOI] [PubMed] [Google Scholar]
  7. Berry C.E., Davies K.A., Owens D.W., Wilcox M.H. Is there a relationship between the presence of the binary toxin genes in Clostridium difficile strains and the severity of C. difficile infection (CDI)? Eur. J. Clin. Microbiol. Infect. Dis. 2017;36:2405–2415. doi: 10.1007/s10096-017-3075-8. [DOI] [PubMed] [Google Scholar]
  8. Borren N.Z., Ghadermarzi S., Hutfless S., Ananthakrishnan A.N. The emergence of Clostridium difficile infection in Asia: A systematic review and meta-analysis of incidence and impact. PLoS ONE. 2017;12 doi: 10.1371/journal.pone.0176797. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Brown, K., Khanafer, N., Daneman, N., Fisman, D., 2013. Meta-analysis of antibiotics and the risk of community-associated Clostridium difficile infection. 57, 5, 2326–2332. https://doi: 10.1128/AAC.02176-12. [DOI] [PMC free article] [PubMed]
  10. Burke K.E., Lamont J.T. Clostridium difficile infection: a worldwide disease. Gut Liver. 2014;8:1–6. doi: 10.5009/gnl.2014.8.1.1. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Cadnum J.L., Hurless K.N., Deshpande A., Nerandzic M.M., Kundrapu S., Donskey C.J. Sensitive and selective culture medium for detection of environmental Clostridium difficile isolates without requirement for anaerobic culture conditions. J. Clin. Microbiol. 2014;52:3259–3263. doi: 10.1128/JCM.00793-14. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Carey-Ann B.D., Carroll K.C. Diagnosis of Clostridium difficile Infection: an Ongoing Conundrum for Clinicians and for Clinical Laboratories. Clin. Microbiol. Rev. 2013;26:604–630. doi: 10.1128/CMR.00016-13. [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. CDC, 2021. Antibiotics can be lifesaving… [WWW Document]. Centers for Disease Control and Prevention. URL https://www.cdc.gov/cdiff/index.html (accessed 1.31.21).
  14. Centers for Disease Control and Prevention (U.S.), 2019. Antibiotic resistance threats in the United States, 2019. Centers for Disease Control and Prevention (U.S.). https://doi.org/10.15620/cdc:82532
  15. Chandrasekaran R., Lacy D.B. The role of toxins in Clostridium difficile infection. FEMS Microbiol. Rev. 2017;41:723–750. doi: 10.1093/femsre/fux048. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Cheng J.-W., Xiao M., Kudinha T., Kong F., Xu Z.-P., Sun L.-Y., Zhang L., Fan X., Xie X.-L., Xu Y.-C. Molecular Epidemiology and Antimicrobial Susceptibility of Clostridium difficile Isolates from a University Teaching Hospital in China. Front. Microbiol. 2016;7:1621. doi: 10.3389/fmicb.2016.01621. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Cheng, M.P., Domingo, M.-C., Lévesque, S., Yansouni, C.P., 2016. A case report of a deep surgical site infection with Terrisporobacter glycolicus/T. Mayombei and review of the literature. BMC Infect. Dis. 16. https://doi.org/10.1186/s12879-016-1865-8. [DOI] [PMC free article] [PubMed]
  18. Curcio D., Cané A., Fernández F.A., Correa J. Clostridium difficile-associated Diarrhea in Developing Countries: A Systematic Review and Meta-Analysis. Infect. Dis. Ther. 2019;8:87–103. doi: 10.1007/s40121-019-0231-8. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Czepiel J., Dróżdż M., Pituch H., Kuijper E.J., Perucki W., Mielimonka A., Goldman S., Wultańska D., Garlicki A., Biesiada G. Clostridium difficile infection: review. Eur. J. Clin. Microbiol. Infect. Dis. 2019;38:1211–1221. doi: 10.1007/s10096-019-03539-6. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. de Jong, E., de Jong, A.S., Bartels, C.J.M., van der Rijt-van den Biggelaar, C., Melchers, W.J.G., Sturm, P.D.J., 2012. Clinical and laboratory evaluation of a real-time PCR for Clostridium difficile toxin A and B genes. Eur. J. Clin. Microbiol. Infect. Dis. 31, 2219–2225. https://doi.org/10.1007/s10096-012-1558-1. [DOI] [PMC free article] [PubMed]
  21. European Centre for Disease Prevention and Control. Healthcare-associated infections: Clostridium difficile infections - Annual Epidemiological Report for 2016. Available at: https://www.ecdc.europa.eu/sites/default/files/documents/AER_for_2016-C-difficile_0.pdf. Accessed March, 2020.
  22. Fang F.C., Polage C.R., Wilcox M.H. Point-Counterpoint: What Is the Optimal Approach for Detection of Clostridium difficile Infection? J. Clin. Microbiol. 2017;55:670–680. doi: 10.1128/JCM.02463-16. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Heinlen L., Ballard J.D. Clostridium difficile infection. Am. J. Med. Sci. 2010;340:247–252. doi: 10.1097/MAJ.0b013e3181e939d8. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Hensgens M.P.M., Goorhuis A., Dekkers O.M., van Benthem B.H.B., Kuijper E.J. All-cause and disease-specific mortality in hospitalized patients with Clostridium difficile infection: a multicenter cohort study. Clin. Infect. Dis. 2013;56:1108–1116. doi: 10.1093/cid/cis1209. [DOI] [PubMed] [Google Scholar]
  25. Hudhaiah D., Elhadi N. Prevalence and Genotypes of Nosocomial Clostridium difficile Infections in the Eastern Province of the Kingdom of Saudi Arabia: A Multi-Centre Prospective Study. JCDR. 2019 doi: 10.7860/JCDR/2019/38321.12694. [DOI] [Google Scholar]
  26. Lessa F.C., Gould C.V., McDonald L.C. Current status of Clostridium difficile infection epidemiology. Clin. Infect. Dis. 2012;55(Suppl 2):S65–S70. doi: 10.1093/cid/cis319. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. McDonald L.C., Gerding D.N., Johnson S., Bakken J.S., Carroll K.C., Coffin S.E., Dubberke E.R., Garey K.W., Gould C.V., Kelly C., Loo V., Shaklee Sammons J., Sandora T.J., Wilcox M.H. Clinical Practice Guidelines for Clostridium difficile Infection in Adults and Children: 2017 Update by the Infectious Diseases Society of America (IDSA) and Society for Healthcare Epidemiology of America (SHEA) Clin. Infect. Dis. 2018;66:e1–e48. doi: 10.1093/cid/cix1085. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Ohashi Y., Fujisawa T. Analysis of Clostridium cluster XI bacteria in human feces. Biosci. Microbiota Food Health. 2019;38:65–68. doi: 10.12938/bmfh.18-023. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Oka K., Osaki T., Hanawa T., Kurata S., Okazaki M., Manzoku T., Takahashi M., Tanaka M., Taguchi H., Watanabe T., Inamatsu T., Kamiya S. Molecular and microbiological characterization of Clostridium difficile isolates from single, relapse, and reinfection cases. J. Clin. Microbiol. 2012;50:915–921. doi: 10.1128/JCM.05588-11. [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Özsoy S., İlki A. Detection of vancomycin-resistant enterococci (VRE) in stool specimens submitted for Clostridium difficile toxin testing. Braz. J. Microbiol. 2017;48:489–492. doi: 10.1016/j.bjm.2016.12.012. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Peery A.F., Dellon E.S., Lund J., Crockett S.D., McGowan C.E., Bulsiewicz W.J., Gangarosa L.M., Thiny M.T., Stizenberg K., Morgan D.R., Ringel Y., Kim H.P., DiBonaventura M.D., Carroll C.F., Allen J.K., Cook S.F., Sandler R.S., Kappelman M.D., Shaheen N.J. Burden of gastrointestinal disease in the United States: 2012 update. Gastroenterology. 2012;143:1179–1187.e3. doi: 10.1053/j.gastro.2012.08.002. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Persson S., Torpdahl M., Olsen K.E.P. New multiplex PCR method for the detection of Clostridium difficile toxin A (tcdA) and toxin B (tcdB) and the binary toxin (cdtA/cdtB) genes applied to a Danish strain collection. Clin. Microbiol. Infect. 2008;14:1057–1064. doi: 10.1111/j.1469-0691.2008.02092.x. [DOI] [PubMed] [Google Scholar]
  33. Pruitt R.N., Lacy D.B. Toward a structural understanding of Clostridium difficile toxins A and B. Front. Cell. Infect. Microbiol. 2012;2:28. doi: 10.3389/fcimb.2012.00028. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Rehman S., Jermy B.R., Akhtar S., Borgio J.F., Abdul Azeez S., Ravinayagam V., Al Jindan R., Alsalem Z.H., Buhameid A., Gani A. Isolation and characterization of a novel thermophile; Bacillus haynesii, applied for the green synthesis of ZnO nanoparticles. Artif. Cells Nanomed. Biotechnol. 2019;47:2072–2082. doi: 10.1080/21691401.2019.1620254. [DOI] [PubMed] [Google Scholar]
  35. Rodriguez C., Warszawski N., Korsak N., Taminiau B., Van Broeck J., Delmée M., Daube G. Laboratory identification of anaerobic bacteria isolated on Clostridium difficile selective medium. Acta Microbiol. Immunol. Hung. 2016;63:171–184. doi: 10.1556/030.63.2016.2.3. [DOI] [PubMed] [Google Scholar]
  36. Schwan C., Stecher B., Tzivelekidis T., van Ham M., Rohde M., Hardt W.-D., Wehland J., Aktories K. Clostridium difficile Toxin CDT Induces Formation of Microtubule-Based Protrusions and Increases Adherence of Bacteria. PLoS Pathog. 2009;5 doi: 10.1371/journal.ppat.1000626. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Shajan S.E., Hashim M.F., Michael A. Prevalence of Clostridium difficile toxin in diarrhoeal stool samples of patients from a general hospital in eastern province, Saudi Arabia. Inte. Jour. of Medi. Res. & Health Sci. 2014;3:302. doi: 10.5958/j.2319-5886.3.2.064. [DOI] [Google Scholar]
  38. Tenover F.C., Tickler I.A., Persing D.H. Antimicrobial-Resistant Strains of Clostridium difficile from North America. Antimicrob. Agents Chemother. 2012;56:2929–2932. doi: 10.1128/AAC.00220-12. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Walter B.M., Rupnik M., Hodnik V., Anderluh G., Dupuy B., Paulič N., Žgur-Bertok D., Butala M. The LexA regulated genes of the Clostridium difficile. BMC Microbiol. 2014;14:88. doi: 10.1186/1471-2180-14-88. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Wiegand P.N., Nathwani D., Wilcox M.H., Stephens J., Shelbaya A., Haider S. Clinical and economic burden of Clostridium difficile infection in Europe: a systematic review of healthcare-facility-acquired infection. J. Hosp. Infect. 2012;81:1–14. doi: 10.1016/j.jhin.2012.02.004. [DOI] [PubMed] [Google Scholar]
  41. Xiao Y., Liu Y., Qin X. Comparative Study of Clostridium difficile Clinical Detection Methods in Patients with Diarrhoea [WWW Document] Can. J. Infect. Dis. Med. Microbiol. 2020 doi: 10.1155/2020/8753284. [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Zakham F., Pillonel T., Brunel A.-S., Zambelli P.-Y., Greub G., Croxatto A., Bertelli C. Molecular diagnosis and enrichment culture identified a septic pseudoarthrosis due to an infection with Erysipelatoclostridium ramosum. Int. J. Infect. Dis. 2019;81:167–169. doi: 10.1016/j.ijid.2019.02.001. [DOI] [PubMed] [Google Scholar]

Articles from Saudi Journal of Biological Sciences are provided here courtesy of Elsevier

RESOURCES