ABSTRACT.
Bacterial infections pose a global threat, especially in the pediatric population. Antimicrobials that are used to treat such infections continuously show reduced efficacy, and empirical therapy is a major treatment option in Rwanda. This study aimed to determine the resistance rate of commonly used antibiotics in pediatric patients. The study was conducted from June 1, 2018 to May 30, 2019, and microbiological samples were collected from 712 children with suspected bacterial infections. Antimicrobial sensitivity testing was performed on 177 positive cultures (24%) that were considered for data analysis. The findings show that the major bacterial isolates were Klebsiella pneumoniae (n = 50, 28.2%), Escherichia coli (n = 47, 26.5%), and Staphylococcus aureus (n = 38, 21.4%). In general, the greatest antibiotic resistance rate was observed in ampicillin (n = 125, 86.2%), amoxicillin–clavulanic acid (n = 84, 82.4%), amoxicillin (n = 64, 79%), cefadroxil (n = 83, 69.2%), tetracycline (n = 72, 59.7%), ceftazidime (n = 42, 55.3%), and cefuroxime (n = 14, 53.8%). More specifically, Klebsiella pneumoniae was 100% resistant to amoxicillin-clavulanic acid, cefuroxime, trimethoprim–sulfamethoxazole, ceftazidime, erythromycin, and clindamycin. Staphylococcus aureus was 86.7% resistant to ampicillin, and Escherichia coli was 91.7% resistant to tetracycline, 90.6% resistant to ampicillin, 83.3% resistant to amoxicillin–clavulanic acid, 79.3% resistant to cefadroxil, and 78.6% resistant to ceftazidime. Moreover, Klebsiella pneumoniae from blood and urine was 96.8% and 100% sensitive, respectively, to meropenem. Staphylococcus aureus from blood was 100% sensitive to vancomycin, whereas Escherichia coli from urine was sensitive to clindamycin (100%), nitrofurantoin (80.6%), and ciprofloxacin (72.7%). In conclusion, our findings show a high resistance rate to commonly used antibiotics, which suggests precaution in empirical therapy and continued surveillance of antimicrobial resistance.
INTRODUCTION
Pediatric bacterial infections pose a global threat and cause high morbidity and mortality.1,2 This health threat is the result of a high infection rate, irrational use of antimicrobials, inadequate infection prevention and control practices, and resource-limited or unavailable diagnostic facilities.3,4 Moreover, antibiotics—one of the major achievements of the 20th century—are increasingly having reduced efficacy and thus poor treatment outcome.5–7 In addition, the emergence of resistance is fast and remains unresolved.
Microorganisms have multiple mechanisms of resistance to each and every antibiotic introduced into practice. This is a consequence of the immense genetic plasticity of bacterial pathogens, which trigger specific responses and result in mutational adaptations, acquisition of genetic material, or alteration of gene expression, producing resistance to virtually all antibiotics currently available in clinical practice. The most known resistance acquisitions includes gene mutation, horizontal gene transfer, overexpression of efflux pumps that extrude the drug from the cell, and change in antibiotic target site.8–10 All these phenomena contribute to continuous antibiotic resistance acquisition, inefficacy of the antibiotic, and poor patient prognosis.
Prediction of prognosis and treatment outcome of pediatric patients with sepsis depends on early and accurate diagnosis and treatment with susceptible antimicrobials. Nevertheless, the outcome is not always good as a result of empirical therapy using unsusceptible drugs.11,12 Yet, empirical therapy remains the single option where antimicrobial sensitivity testing is not feasible in poor-resource settings, including those located in Rwanda.13–15
Therapeutic guidelines based on currently available information regarding microbial infection etiology and antimicrobial susceptibility are needed to guide empirical therapy.13,16 However, these data are scarce and are not updated regularly, while antimicrobial resistance continuous to change.17 Thus, it is paramount to know and monitor bacterial infections and their antibiotic resistance profile.18,19 Furthermore, continued surveillance of antimicrobial resistance would help decision makers tackle the emerging threat to saving lives, especially in highly vulnerable pediatric groups.20,21 At the University Teaching Hospital of Butare (UTHB) in Rwanda, we evaluated the bacteria responsible for pediatric infection in relation to their antibiotic resistance profile.
MATERIALS AND METHODS
Study setting.
Our study was conducted at the UTHB, a tertiary care teaching hospital with 500 beds, located in the Southern Province, Huye District, Rwanda. The hospital serves as a teaching and referral hospital for both inpatients and outpatients referred from different district hospitals within the southern part of the country.
Study population.
The target population was children younger than 15 years who visited the pediatric department during the study period and who were suspected of having bacterial infection with recurrent fever, urinary tract infection (UTI), or any other sign suggesting bacterial infection. The study was conducted from June 1, 2018 to May 30, 2019. Different microbiological samples from 712 children were collected and sent to the laboratory for culture. A total of 177 samples (24.8%) exhibited a positive microbiological culture and were considered for data analysis. All negative microbiological cultures were excluded from data analysis.
Sample collection.
Clinical specimens were collected and sent to the microbiology laboratory for culture. The collected samples included urine, blood, sputum, stool, pus swab, eye swab, throat swab, and a vaginal swab from pediatric patients who were suspected of having a bacterial infection. Blood samples were collected and incubated in a pediatric blood culture bottle (BD Bactec Ped Plus; BD, Franklin Lakes, NJ). Urine, wound, stool, eye swab, throat swab, vaginal swab, and sputum cultures were collected in sterile containers. The material for laboratory testing, including sterile containers, antibiotic disks, culture media, and swabs, was obtained from BD.
Microbiologic sample processing.
All the samples were processed in accordance with current laboratory standard operating procedures of the UTHB Department of Pathology. Briefly, urine samples, after wet mount examination, were cultured on cysteine lactose electrolyte deficient. The colonies were counted to rule out significant growth, and ≥ 105 colony-forming units/mL was considered significant growth. For swab and sputum samples, Gram stain was used to direct culture media selection. Blood culture bottles were incubated directly at 37°C in an automated blood culture machine (Bactec, BD). The subculture of positive bottles was done on sheep blood agar, and the samples were incubated overnight at 37°C at 5% carbon dioxide. Bacterial species were identified manually with Gram stain and biochemical tests. Gram-positive cocci were identified by catalase, coagulase, and homolysis properties. Gram-negative bacilli were identified by colony morphology and biochemical tests, including triple sugar iron, motility indole urease, and citrate tests.
Antibiotic susceptibility testing.
Disk diffusion or the Kirby Bauer method was used for antibiotic susceptibility testing. Different antibiotic disks were used, including ampicillin, 10 μg; ceftazidime, 30 μg; cefotaxime, 30 μg; cefuroxime, 30 μg; ciprofloxacin, 5 μg; trimethoprim–sulfamethoxazole (TMP–SMX or cotrimoxazole), 1.25/23.75 μg; Augmentin (amoxicillin–clavulanic acid), 20/10 u μg; clindamycin, 2 μg; cloxacillin, 1 μg; erythromycin, 10 μg; gentamicin, 10 μg; meropenem, 10 μg; levofloxacin, 5 μg; nalidixic acid, 30 μg; nitrofurantoin, 300 μg; norfloxacin, 10 μg; amoxicillin, 30 μg; chloramphenicol, 30 μg; oxacillin, 1 μg; vancomycin, 30 μg; and tetracycline, 30 μg (BD).
The saline suspension from a pure culture plate was prepared by adding bacterial colonies into sterile saline at 0.9% until the MacFarland turbidity standard of 0.5 was attained. The resulting suspension was inoculated on Muller–Hinton agar by using a sterile cotton swab. After this procedure, the antibiotic disks were added to the plate with at least 20 mm between each disk. Disks were subsequently incubated at 37°C for 18 to 24 hours. Thereafter, interpretation of the diameter of inhibition was done according to the 2017 Clinical and Laboratory Standards Institute (CLSI) guidelines. Extended-spectrum β-lactase (ESBL) and methicillin-resistant Staphylococcus aureus (MRSA) phenotypes were determined in accordance with the 2017 CLSI guidelines.
Quality control.
Quality control was performed according to the standard operating procedures of the UTHB Department of Pathology, with reference to CLSI guidelines. Culture media sterility control and Kirby Bauer disk diffusion test control were performed using different American Type Culture Collection (ATCC) strains, including Escherichia coli ATCC 25922 (Oxoid, UK), Staphylococcus aureus ATCC 25923 (Oxoid, UK), and Pseudomonas spp. ATCC 27853 (Oxoid, UK). Suspensions of the organisms were prepared as described earlier, and the inhibition diameter obtained was compared with the standard range expected from the CLSI for ATCC strains.
Data analysis.
Data were captured, recorded, and stored in Microsoft Excel 2016 (Microsoft Corporation, Redmond, WA) and analyzed further using SPSS (version 22, SPSS Inc., Chicago, IL). The frequency, percentages, and means were calculated in SPSS.
Ethical consideration.
The ethical clearance for the study was obtained from the Institution Review Board of the UTHB (CHUB/DG/SA/11/2301/2019). All participants/parents of the children gave written consent and agreed to participate voluntarily in the study.
RESULTS
Characteristics of the study participants.
In our study, 177 children exhibited positive microbiologic culture results and were considered for data analysis, with 105 positive results (59.3%) from girls and 72 positive results (40.7%) from boys. Infection was more distributed in children 1 to 5 years of age (n = 34.1%), followed by children older than 5 years (n = 58, 33%) and neonates (n = 39, 22.2%), with a median age of 4 years. Among all positive samples, blood cultures were acquired from 70 participants (39.5%), urine cultures from 60 (33.8%), and pus cultures from 40 (22.5%). Other cultured samples (eye swab, stool, sputum, throat swab, and vaginal swab) were less frequent (n = 7, 3.9%; Table 1).
Table 1.
Demographic characteristics of the participants with positive microbiologic cultures
| Variable | Frequency, n (%) |
|---|---|
| Gender | |
| Female | 105 (59.3) |
| Male | 72 (40.7) |
| Age group* | |
| Neonate (0–28 days) | 39 (22.2) |
| < 6 months | 10 (5.7) |
| 6–12 months | 9 (5.1) |
| 1–5 years | 60 (34.1) |
| > 5 years | 58 (33) |
| Culture type | |
| Blood | 70 (39.5) |
| Urine | 60 (33.8) |
| Pus | 40 (22.5) |
| Other† | 7 (3.9) |
| Total | 177 (100) |
Median age, 4.0 years.
Includes eye swab, stool, sputum, throat swab, and vaginal swab.
Frequency of bacterial isolates in different samples.
Clinical management of sepsis varies depending on the affected body site. we investigated the frequencies of bacterial isolates in different body sites. Our data show that Klebsiella pneumoniae (n = 32, 45.7%) and S. aureus (n = 19, 27.1%) were major blood isolates. Escherichia coli (n = 39, 65.9%) was a major cause of UTIs, whereas S. aureus (n = 18, 45%) and K. pneumoniae (n = 11, 27.5%) were predominant isolates from pus. Taken together, K. pneumoniae, E. coli, and S. aureus were the major causes of pediatric infections in our study subjects (Table 2).
Table 2.
Frequency of bacterial isolates according to sample type
| Bacterial species | Sample type, n (%) | ||||
|---|---|---|---|---|---|
| Blood | Urine | Pus | Other* | Total | |
| Acinetobacter spp. | 6 (8.5) | 0 (0.0) | 1 (2.5) | 0 (0.0) | 7 (3.9) |
| Escherichia coli | 4 (5.7) | 39 (65.0) | 4 (10.0) | 0 (0.0) | 47 (26.5) |
| Enterobacter spp. | 2 (2.8) | 3 (5.0) | 1 (2.5) | 0 (0.0) | 6 (3.3) |
| Klebsiella oxytoca | 2 (2.8) | 7 (4.2) | 1 (2.5) | 2 (28.5) | 12 (6.7) |
| Klebsiella pneumoniae | 32 (45.7) | 6 (10.0) | 11 (27.5) | 1 (14.2) | 50 (28.2) |
| Pseudomonas aeruginosa | 3 (4.2) | 1 (1.6) | 0 (0.0) | 0 (0.0) | 4 (2.2) |
| Proteus spp. | 0 (0.0) | 2 (3.3) | 3 (7.5) | 1 (14.2) | 6 (3.3) |
| Staphylococcus aureus | 19 (27.1) | 1 (1.6) | 18 (45.0) | 0 (0.0) | 38 (21.4) |
| Salmonella spp. | 1 (1.4) | 0 (0.0) | 0 (0.0) | 1 (14.) | 2 (1.1) |
| Shigella spp. | 0 (0.0) | 0 (0.0) | 0 (0.0) | 1 (14.2) | 1 (0.5) |
| Streptococcus spp. | 1 (1.4) | 1 (1.6) | 1 (2.5) | 1 (14.2) | 4 (2.2) |
| Total | 70 (100) | 60 (100) | 40 (100) | 7 (100) | 177 (100) |
Includes eye swab, stool, sputum, throat swab, and vaginal swab.
Frequency of bacterial isolates in different age categories
Epidemiology and prognosis of bacterial infection differ in different age categories, which may require special clinical intervention. In our study, we investigated bacterial pathogens responsible for pediatric infection in different age groups. Our findings show that K. pneumoniae (n = 22, 55%) was the most frequent isolate from neonates. In children younger than 6 months, and 6 to 12 months, Klebsiella oxytoca (n = 4, 40%) and S. aureus (n = 3, 33.3%) were the major isolates. In addition, E. coli was more frequent in children 1 to 5 years (n = 23, 38.3%) and older than 5 years (n = 18, 31%). Our findings highlight that K. pneumoniae was a prominent cause of neonate infection, whereas E. coli and S. aureus were the leading causes of infection in other age groups (Table 3).
Table 3.
Bacterial isolate by age group, measured as a percentage
| Bacterial species | Age group | |||||
|---|---|---|---|---|---|---|
| Neonates | > 6 Months | 6 Months–1 year | 1–5 Years | > 5 Years | Total | |
| Acinetobacter spp. | 5 (12.5) | 0 (0.0) | 1 (11.1) | 0 (0.0) | 1 (1.7) | 7 (4.0) |
| Escherichia coli | 3 (7.5) | 1 (10.0) | 2 (22.2) | 23 (38.3) | 18 (31.0) | 47 (26.6) |
| Enterobacter spp. | 1 (2.5) | 1 (10.0) | 0 (0.0) | 2 (3.3) | 2 (3.4) | 6 (3.4) |
| Klebsiella oxytoca | 1 (2.5) | 4 (40.0) | 0 (0.0) | 4 (6.7) | 3 (5.2) | 12 (6.8) |
| Klebsiella pneumoniae | 22 (55.0) | 2 (20.0) | 2 (22.2) | 13 (21.7) | 11 (19.0) | 50 (28.2) |
| Pseudomonas aeruginosa | 0 (0.0) | 0 (0.0) | 1 (11.1) | 2 (3.3) | 1 (1.7) | 4 (2.3) |
| Proteus spp. | 1 (2.5) | 0 (0.0) | 0 (0.0) | 3 (5.0) | 1 (1.7) | 6 (3.4) |
| Staphylococcus aureus | 7 (17.5) | 1 (10.0) | 3 (33.3) | 12 (20.0) | 15 (25.9) | 38 (21.5) |
| Salmonella spp. | 0 (0.0) | 0 (0.0) | 0 (0.0) | 0 (0.0) | 2 (3.4) | 2 (1.1) |
| Shigella spp. | 0 (0.0) | 0 (0.0) | 0 (0.0) | 0 (0.0) | 1 (1.7) | 1 (0.6) |
| Streptococcus spp. | 0 (0.0) | 1 (10.0) | 0 (0.0) | 1 (1.7) | 2 (3.4) | 4 (2.3) |
| Total | 40 (100) | 10 (100) | 9 (100) | 60 (100) | 58 (100) | 177 (100) |
Antimicrobial resistance profile of bacterial isolates.
Although antibiotics are used to treat bacterial infections effectively, antimicrobial resistance acquisition and superbugs constitute a global challenge, and antimicrobial sensitivity testing is not common or is unavailable in low-income countries. We evaluated the antimicrobial resistance of all isolates toward different antibiotics. Our findings show that the greatest resistance rates were seen in ampicillin (n = 125, 86.2%), amoxicillin–clavulanic acid (n = 84, 82.4%), amoxicillin (n = 64, 79%), cefadroxil (n = 83, 69.2%), tetracycline (n = 43, 59.7%), ceftazidime (n = 42, 55.3%), and cefuroxime (n = 14, 53.8%). In addition, ESBL phenotypes and MRSA rates were 5 (31.3%) and 12 (38.7%) respectively. Moreover, the greatest antimicrobial sensitivity rate was observed in meropenem (n = 118, 95.9%), levofloxacin (n = 31, 83.8%), nitrofurantoin (n = 48, 78.7%), vancomycin (n = 7, 77.8%), and ciprofloxacin (n = 56, 75.7%). In summary, our findings show a greater resistance rate in penicillin and cephalosporin, whereas carbapenems, quinolone, peptides, and nitrofuran were the most sensitive antimicrobials (Table 4).
Table 4.
Overall antimicrobial resistance in all clinical isolates
| Antibiotic | n | R, n (%) | S, n (%) | ESBL*, n (%) | MRSA, n (%) |
|---|---|---|---|---|---|
| Ampicillin | 145 | 125 (86.2) | 20 (13.8) | – | – |
| Nitrofurantoin | 66 | 13 (21.3) | 48 (78.7) | – | – |
| Negram | 58 | 21 (36.2) | 37 (63.8) | – | – |
| Norfloxacin | 50 | 15 (30) | 35 (70) | – | – |
| Cefadroxil | 120 | 83 (69.2) | 37 (30.8) | – | – |
| Ciprofloxacin | 74 | 18 (24.3) | 56 (75.7) | – | – |
| Meropenem | 123 | 5 (4.1) | 118 (95.9) | – | – |
| Ceftazidime | 76 | 42 (55.3) | 34 (44.7) | 12 (38.7) | – |
| Amoxicillin–clavulanate | 102 | 84 (82.4) | 18 (17.6) | – | |
| Cefotaxime | 31 | 12 (38.7) | 19 (61.3) | – | |
| Cefuroxime | 26 | 14 (53.8) | 12 (46.2) | – | – |
| Trimethoprim–sulfamethoxazole | 13 | 8 (61.5) | 5 (38.5) | – | – |
| Tetracycline | 72 | 43 (59.7) | 29 (40.3) | – | – |
| Cloxacillin | 28 | 9 (32.1) | 19 (67.9) | – | – |
| Erythromycin | 34 | 10 (29.4) | 24 (70.6) | – | – |
| Gentamycin | 118 | 51 (43.2) | 67 (56.8) | – | – |
| Cefoxitin | 16 | 5 (31.3) | 11 (68.8) | – | 5 (31.3) |
| Vancomycin | 9 | 2 (22.2) | 7 (77.8) | – | – |
| Clindamycin | 23 | 3 (13) | 20 (87) | – | – |
| Levofloxacin | 37 | 6 (16.2) | 31 (83.8) | – | – |
| Chloramphenicol | 43 | 16 (37.2) | 27 (62.8) | – | – |
| Amoxicillin | 81 | 64 (79) | 17 (21) | – | – |
ESBL = extended-spectrum β-lactamase; MRSA = methicillin-resistant Staphylococcus aureus; R = resistant, S = sensitive.
ESBL phenotypes were detected by the double disk-diffusion method, and synergy with ceftazidime, amoxicillin–clavulanate, and cefotaxime were evaluated.
MRSA was detected by evaluating resistance on the cefoxitin disk.
Resistance rate in major bacterial isolates in commonly used drugs.
Isolates from different body sites could exhibit different susceptibility patterns and require different treatment regimen. We studied the antimicrobial resistance of major bacterial isolates from blood, urine, and pus. In blood, K. pneumoniae was 100% resistant to amoxicillin–clavulanic acid, cefuroxime, TMP–SMZ, ceftazidime, erythromycin, and clindamycin. Staphylococcus aureus from blood was 86.7% resistant to ampicillin and 57.1% resistant to cefoxitin, suggesting MRSA. In urine isolates, K. pneumoniae was 100% resistant to cefadroxil, amoxicillin–clavulanic acid, cefuroxime, and ceftazidime. Escherichia coli was 91.7% resistant to tetracycline, 90.6% resistant to ampicillin, 83.3% resistant to amoxicillin–clavulanic acid, 79.3% resistant to cefadroxil, and 78.6% resistant to ceftazidime. In pus isolates, K. pneumoniae was 100% resistant to amoxicillin–clavulanic acid and ceftazidime. However, a low antimicrobial resistance rate was observed in blood and pus isolates for vancomycin and S. aureus, and for meropenem and Klebsiella pneumonia. In urine, meropenem, nitrofurantoin, ciprofloxacin, and clindamycin were less resistant. Taken together, our findings highlight a high resistance rate in the most commonly used drugs. However, meropenem was the most sensitive drug in Gram-negative pathogens, and vancomycin was the most sensitive drug in Gram-positive pathogens. Urine isolates were mostly sensitive to meropenem, nitrofurantoin, ciprofloxacin, and clindamycin (Table 5).
Table 5.
Antimicrobial resistance, measured as a percentage, of major bacterial isolates on commonly used antibiotics
| Culture type | Bacterium | Antibiotic | |||||||||||||||||||||||||||||
|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|
| Ampicillin | Cefadroxil | Amox clav | Cefuroxime | TMP-SMZ | Tetracycline | Ceftazidime | Amoxicillin | Nitrofurantoin | Ciprofloxacin | Meropenem | Erythro | Clindamycin | Cefoxitin | VA | |||||||||||||||||
| R | S | R | S | R | S | R | S | R | S | R | S | R | S | R | S | R | S | R | S | R | S | R | S | R | S | R | S | R | S | ||
| Blood | Klebsiella pneumoniae | 96.4 | 3.6 | 69.2 | 30.8 | 100 | 0.0 | 100 | – | 100 | 0.0 | 87.5 | 12.5 | 15.4 | 100 | – | – | – | – | 35 | 65 | 3.2 | 96.8 | 100 | 0.0 | 100 | 0.0 | – | – | – | – |
| Staphylococcus aureus | 86.7 | 13.3 | 66.7 | 33.3 | 36.4 | 63.6 | 0.0 | 100 | 0.0 | 100 | 66.7 | 33.3 | 12.5 | 63.6 | 63.6 | 36.4 | – | – | 40 | 60 | – | – | 40 | 60 | 16.7 | 83.3 | 57.1 | 42.9 | 0.0 | 100 | |
| Urine | K. pneumoniae | 80 | 20 | 100 | 0.0 | 100 | 0.0 | 100 | 0.0 | – | – | – | 100 | 25 | 100 | – | – | 25 | 75 | 0.0 | 100 | 100 | 100 | 0.0 | 100 | – | – | – | – | ||
| Escherichia coli | 90.6 | 9.4 | 79.3 | 20.7 | 83.3 | 16.7 | 0.0 | 100 | 66.7 | 33.3 | 91.7 | 8.3 | 73.3 | 78.6 | 0.0 | 21.4 | 19.4 | 80.6 | 27.3 | 72.7 | 0.0 | 100 | – | – | 0.0 | 100 | – | – | – | – | |
| Pus | K. pneumoniae | 90.9 | 9.1 | 77.8 | 22.2 | 100 | 0.0 | – | – | – | – | 50 | 50 | 25 | 100 | 100 | 0 | – | – | 33.3 | 66.7 | 12.5 | 87.5 | – | – | – | – | – | – | – | – |
| S. aureus | 70 | 30 | 14.3 | 85.7 | 66.7 | 33.3 | – | – | – | – | 25 | 75 | 25 | 55.6 | 53.6 | 44.4 | – | – | 12.5 | 87.5 | – | – | 0.0 | 100 | 0.0 | 100 | 12.5 | 87.5 | 0.0 | 100 | |
The disk diffusion method was performed on different antibiotics. Amox clav = amoxicillin–clavulanic acid; Erythro = erythromycin; R = resistant, S = sensitive; TMP-SMZ = trimethoprim–sulfamethoxazole; VA = vancomycin.
DISCUSSION
Bacterial infection is a public health concern with an increased risk in pediatric patients.1,22 In addition, the emergence of antimicrobial resistance and lack of microbiologic diagnostic facilities in low-income counties constitute major challenges and handicap treatment options. We evaluated bacterial infections in pediatric patients in Rwanda. Our findings show that the infection rate was more distributed in children younger than 5 years (34.1%). Among all positive microbiologic cultures, blood cultures were 39.5% followed by urine culture at 33.8%. Similar findings were described in recent reports on bacterial infections and antimicrobial resistance.7,23,24 Children younger than 5 years were more susceptible to infection than other age groups, and this is a result of their immunity, which is not fully developed at the time of infection.
Understanding major bacterial isolates according to sample type provides key guidance on affected organs and helps to choose appropriate treatment. We found that 45.7% of bloodstream infections was caused by K. pneumoniae, 65% of UTIs were mainly caused by E. coli, and 45% of S. aureus were isolated from pus. Our findings are in accordance with previous reports in which E. coli, K. pneumoniae, and S. aureus were the main pathogen isolates.7,23,25 It is also in line with WHO surveillance data from 22 countries, in which among 500,000 isolates, E. coli, K. pneumoniae, and S. aureus were the most commonly reported bacteria.26,27 These findings highlight the clinical importance of these bacterial species as major causes of bloodstream infections, UTIs, and wound-associated infections.
Furthermore, we have shown that K. pneumoniae was a prominent cause of neonate infections (55%), whereas E. coli (38.3%) and S. aureus (25.9%) were the leading causes of infections in other age groups, which is in line with previous reports.28,29 Klebsiella pneumoniae in neonates is most likely linked to hospital-acquired infections, as reported previously.30 Escherichia coli, an uropathogen from the gut, as reported previously,31 was a more common cause of UTIs in our participants. This is a result of its nature with virulence factors that help to colonize the urinary tract. Other explanations include self-contamination with gut flora or urinary catheters. Isolation of S. aureus in pus indicates its association with wound or surgical site infections. Staphylococcus aureus is a pyogenic bacterium that is part of human normal flora that easily infect wounds or cause surgical site infections, especially when aseptic procedures are not followed.
Antimicrobial sensitivity testing is not common or is unavailable in poor-resource countries, and empirical antimicrobial therapy remains a single option, regardless of its association with poor treatment outcome.32 Our results show a high resistance rate of almost 100% in penicillin and cephalosporin drugs, whereas carbapenems, quinolone, peptides, and nitrofuran exhibited a greater sensitivity rate. Our findings also show that K. pneumoniae was 100% resistant to amoxicillin–clavulanic acid, cefuroxime, TMP-SMZ, ceftazidime, erythromycin, and clindamycin. Staphylococcus aureus from blood was 86.7% resistant to ampicillin. Escherichia coli was 91.7% resistant to tetracycline, 90.6% resistant to ampicillin, 83.3% resistant to amoxicillin–clavulanic, 79.3% resistant to cefadroxil, and 78.6% resistant to ceftazidime. Our findings are in line with a previous study33 in which the level of multidrug-resistant strains was alarmingly high and treatment outcome was affected negatively, prompting the need for regular, strict guidelines for empirical antibiotic use.
Our findings are also in line with a study34 conducted on the clinical outcome of oral amoxicillin and TMP-SMZ in children between 2 and 59 months at Mulago Hospital in Uganda, in which amoxicillin showed a resistance rate of 89.9% and TMP-SMZ was 77% resistant. Similar results were obtained in a previous study35 conducted in Rwanda, in which ampicillin had 100% resistance, and resistance to third-generation cephalosporins was 71.7%. Concurrently, research conducted in Bangladesh36 on antibiotic resistance as a complex system driven by socioeconomic growth and antibiotic misuses showed a high resistance rate to cotrimoxazole (68.5%), ampicillin (84.5%), and amoxicillin–clavulanic acid (81%). Although the exact same resistance rate was not obtained, antimicrobial resistance is a dynamic phenomenon, and variability in different studies could be associated with recruited participants, geographic location, and whether the study participants were ambulatory or hospitalized.
Nevertheless, E. coli and K. pneumoniae exhibited synergy with ceftazidime, amoxicillin–clavulanic acid, and cefotaxime at 38.7%, suggesting ESBL producers, whereas S. aureus resistance to cefoxitin was 31.3%, suggesting MRSA phenotypes. Our findings are in line with a previous study37 conducted in Mauritania, in which the activity of penicillin G against S. aureus isolates was almost zero, and the MRSA rate was 34%. These findings show increased resistance to penicillin and cephalosporin drugs.
In our study, we show that the greatest antimicrobial sensitivity rate was 95.9% for Gram-negative isolates and meropenem, and a 77.8% sensitivity of Gram-positive isolates and vancomycin. In addition, nitrofurantoin, ciprofloxacin, and clindamycin had the greatest sensitivity rates for uropathogenic isolates. This is in line with the work by Zhanel et al.,38 who found a sensitivity of 89.8% for meropenem, 82.9% for levofloxacin, 87.5% for clindamycin, and 78.8% for chloramphenicol. In addition, the treatment effectivity of imipenem and meropenem showed 91% susceptibility.38 Nevertheless, reduced vancomycin susceptibility has also been reported.39,40 Antimicrobial resistance seems to be a complex mechanism, and the acquisition of resistance genes is unceasing and requires continued surveillance.
Our study had some limitations. First, we did not differentiate outpatients from inpatients to determine hospital and community-acquired infections easily. Second, we show the rate of ESBL and MRSA phenotypes, but we could not perform genotyping and determine resistance mechanisms. Future multicenter studies with a bigger sample size are needed to update the epidemiological status of antimicrobial resistance in Rwanda.
CONCLUSION
Our results highlight that E. coli, K. pneumoniae, and S. aureus were the most common pathogens responsible for pediatric bacterial infection in our study subjects. Penicillin and third-generation cephalosporin drugs have shown a high antimicrobial resistance rate whereas carbapenems, vancomycin, quinolone, and nitrofuran exhibited a greater sensitivity rate. These findings suggest more precautions with regard to empirical treatment, and continued surveillance of antimicrobial resistance.
ACKNOWLEDGMENTS
Our sincere thanks to the administration of the University Teaching Hospital of Butare for their facilitation during this study. The American Society of Tropical Medicine and Hygiene (ASTMH) assisted with publication expenses.
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