Article Information

Jean-Marie Liesse Iyamba^{1, 2,} *, Cyprien Mbundu Lukukula^{1, 2}, Joseph Welo Unya^{1, 2}, Benjamin Kodondi Ngbandani^{1, 2}, Edouard Bissingou¹, Musomoni Mabankama², Nelson Nsiata Ngoma³, Thierry Mukendi Kajinga³, Blaise Mabamu Maya⁴, Aline Diza Lubonga⁴, NB Takaisi-Kikuni^{1, 2}

¹University Reference Center for Monitoring of Antimicrobial Resistance (URCM-AMR) Faculty of Pharmaceutical Sciences, University of Kinshasa, Kinshasa, Democratic Republic of Congo

²Laboratory of Experimental and Pharmaceutical Microbiology, Faculty of Pharmaceutical Sciences , University of Kinshasa, Kinshasa, Democratic Republic of Congo

³Saint Joseph Hospital, Department of Clinical Biology and Internal Medicine, Kinshasa/Limete, Democratic Republic of Congo

⁴Biamba Marie Mutombo Hospital, Department of Bacteriology and Pharmacy, Kinshasa Kinshasa/Masina, Democratic Republic of Congo

*Corresponding author: Jean-Marie Liesse Iyamba, University Reference Center of Antimicrobial Resistance Surveillance (URC-AMRS), Faculty of Pharmaceutical Sciences, University of Kinshasa, Democratic Republic of Congo.

Received: 22 August 2022; Accepted: 02 September 2022; Published: 19 October 2022

Citation: Jean-Marie Liesse Iyamba, Cyprien Mbundu Lukukula, Joseph Welo Unya, Benjamin Kodondi Ngbandani, Edouard Bissingou, Musomoni Mabankama, Nelson Nsiata Ngoma, Thierry Mukendi Kajinga, Blaise Mabamu Maya, Aline Diza Lubonga, NB Takaisi-Kikuni. Antibiotic Resistance Pattern and Biofilm Formation of Staphylococcus and Enterobacteriaceae Isolates from Clinical Samples of Patients with Urinary Tract and Surgical Site Infections in Kinshasa, Democratic Republic of Congo. Journal of Pharmacy and Pharmacology Research 6 (2022): 158-168.

Abstract

Keywords

Antibiotic resistance, Staphylococcus aureus, Enterobacteriaceae, Biofilm

Article Details

1. Introduction

Emergence of resistance to multiple antimicrobial agents in pathogenic bacteria has become a significant public health threat as there are fewer, or even sometimes no, effective antimicrobial agents available for infections caused by these bacteria 1). Gram-positive and Gram-negative bacteria are both affected by the emergence and rise of antimicrobial resistance [1]. Treatment of infections is compromised worldwide by the emergence of bacteria that are resistant to multiple antibiotics [2]. Infections caused by multidrug-resistant organisms (MDROs) are associated with increased mortality, morbidity, length of hospitalization, cost of health care, and the cost-effectiveness of antibiotics with different degrees of resistance [3, 4]. MDROs include vancomycin-resistant enterococci (VRE), methicillin-resistant Staphylococcus aureus (MRSA), and resistant gram-negative bacilli (RGNB) [1]. Bacterial resistance to antibiotics is primarily the consequence of a variety of phenomena such as alteration of the target of the drug, impermeability of the bacteria to the antibiotic, destruction of the antibiotic molecule, efflux system able to pump antibiotic out of the cytoplasm of bacteria, and genetically associated changes (mutational events, genetic transfer of resistance genes through plasmids, and mutations of target genes) [5]. Enterobacteriaceae had become resistant to β-lactam antibiotics and carbapenems due to the production of extended-spectrum beta-lactamases (ESBL) and carbapenemase enzymes such as oxacillinase (OXA)-48-like β-lactamases respectively [6, 7]. However, this is not the only reason for antimicrobial treatment failure. Bacteria are able to colonize host tissues or medical devices and to form a biofilm. By definition, biofilms are microbially derived sessile communities characterized by cells that are irreversibly attached to a substratum or interface or each other, are embedded in a matrix of extracellular polymeric substances that they have produced and exhibit an altered phenotype with respect to growth rate and gene transcription [8]. Growth in biofilms enhances the survival of bacterial populations in hospital settings and inside patients, increasing the probability of causing nosocomial infections. Biofilm formation confers pathogenic bacteria increased resistance to convectional antibiotics and host defenses mechanisms [9]). Previous studies showed a correlation between biofilm production and multiple drug resistance in clinical isolates [10, 11].

Information concerning the true extent of the problem of AMR in the African Region is limited because surveillance of drug resistance is carried out in only a few countries. In order to provide data on antibiotic resistance, our laboratory collects bacterial strains from hospitals in order to monitor the resistance of certain important pathogens. The purpose of the work reported here was to evaluate the antibiotic resistance of S. aureus and Enterobacteriaceae strains from patients with urinary tract and surgical site infection respectively at Biamba Marie Mutombo Hospital and Saint Joseph Hospital located in Eastern Kinshasa city, to determine the prevalence OXA-48-producing Enterobacteriaceae, and to study the formation of the biofilm by clinical strains isolated.

2. Material and Methods

2.1 Bacteria isolates

From Biamba Marie Mutombo Hospital, a total of 13 clinical isolates of S. aureus isolates (from urines, vaginal smears, prostatic fluid, infected devices and from surgical site infections[SSI]), and 19 clinical isolates of Enterobacteriaceae (10 Escherichia coli and 9 Enterobacter sp.) from urinary tract samples (UTI) were investigated. From Saint Joseph Hospital, 5 S. aureus and 41 Enterobacteriaceae (19 E. coli, 8 Enterobacter sp., 9 Citrobacter sp. and 5 Serratia sp.) isolates from SSI were tested. The clinical samples were collected for diagnostic purposes by the bacteriology laboratories of these hospitals, and were from hospitalized and non-hospitalized patients.

All Staphylococcus sp. were initially identified by standard microbiological methods including Gram stain, catalase and coagulase tests. In the microbiology laboratory of the Faculty of Pharmaceutical Sciences, University of Kinshasa, the identification of Staphylococcus aureus strains was performed with latex agglutination test (Pastorex Staph-Plus, BioRad, Marnes-la-Coquette, France) and DNase test. All staphylococcal strains, negative for latex agglutination and DNase tests, were considered as coagulase negative staphylococci.

Isolated strains of Gram negative bacilli were identified using microbiological conventional methods including Gram staining, oxydase tests, indole and urease production, citrate utilization, hydrogen sulphide, gas production and fermentation of sugars, phenylalanine deaminase, lysine decarboxylase (L.D.C.), ornithine decarboxylase (O.D.C.), arginine dihydrolase (A.D.H.) tests, and methyl red reaction. In our laboratory Gram negative bacilli were confirmed as Enterobacteriaceae species using the same tests. All cultures were maintained on trypticase soy agar (Liofilchen, Roseto degli Abruzzi, Italy).

2.2 Antibiotic susceptibility tests

Antibiograms of each isolated Staphylococcus spp strains using the diffusion method on Mueller Hinton Agar were realized with the following antibiotic disks (Liofilchen, Roseto degli Abruzzi, Italy): amikacin (30 µg), amoxicillin + clavulanic acid (30 µg), ampicillin (30µg), ampicillin- sulbactam (30/20 µg), azithromycin (15 µg), aztreonam (30 µg), ceftazidime (30 µg), cefixime (5 µg), ciprofloxacin (5 µg), clarithromycin (15 µg), erythromycin (15 µg), fosfomycin (200 µg), kanamycin (30 µg), levofloxacin (5 µg), netilmicin (30 µg), piperacillin - tazobactam (100/10 µg), teicoplanin (30 µg), temocillin (30 µg), tobramycin (10 µg), trimethoprim (5 µg), and vancomycin (30 µg). Test for methicillin resistance was performed with diffusion method using oxacillin (1 μg) on Mueller Hinton agar with 4 % NaCl.

Enterobacteriaceae were tested against the following antibiotic disks (Liofilchen, Roseto degli Abruzzi, Italy): ampicillin (30 µg), amikacin (10 µg), amoxicillin (10 µg), ampicillin (30 µg), ampicillin-sulbactam (20 µg), aztreonam (30 µg), cefixime (5 µg), cefotaxime (5 µg), cefuroxime (30 µg), ceftazidime (30 µg), fosfomycin (200 µg), imipenem (10 µg), norfloxacin (5 µg), levofloxacin (5 µg), tobramycin (10 µg), temocillin (30 µg), and piperacillin-tazobactam (100/10 µg). After incubation of plates at 37°C for 24 hours, diameters of zone of inhibition were measured. Evaluation of the results was done according to the criteria of Clinical Laboratory Standards Institute (CLSI) [12]. E. coli ATCC 25922 and S. aureus ATCC 25923 were used for quality control.

2.3 Detection of OXA-48 producers

OXA-48-producing Enterobacteriaceae were detected on Chromatic^TM OXA-48 chromogenic medium (Liofilchem, Roseto degli Abbruzzi, Italy). After incubation at 37°C/24-48 hours, the color and the morphology of the colonies were observed and the results interpreted as follow: red colony (E. coli-producing OXA-48), blue-violet colony (Klebsiella sp. producing OXA-48), blue-green (Enterobacter sp. producing OXA-48), blue colony with red halo (Citrobacter sp. producing OXA-48). E. coli ATCC 25922 was used for quality control.

2.4 Biofilm formation assay

In present study, we screened all isolates for their ability to form biofilm by Crystal Violet Staining method as previously described [13]), with modifications. A suspension equivalent to the McFarland 0.5 turbidity standard was prepared in Trypticase Soya broth (Becton Dickinson, Franklin Lake) for each strain. Accuracy of bacterial counts in the suspension was confirmed by serial dilution in log steps. Polystyrene sterile strips were inoculated with 200 μL of each calibrated bacterial suspension and incubated for 24 hours at 35°C in a humid atmosphere. A control well was inoculated with sterile medium. Each strain was evaluated in triplicate. Medium was removed from the wells which were washed 3 times with 200 μL sterile distilled water. The strips were air- dried for 45 min and the adherent cells were stained with 200 μL of 0.1% Crystal violet solution. After 45 min, the dye was eliminated and the wells were washed 5 times with 300 μL of sterile distilled water to remove excess stain. The dye incorporated by the cells forming a biofilm was dissolved with 200 μL of 33% (v/v) glacial acetic acid and the absorbance of the well was obtained by means of enzyme-linked immunosorbent assay (ELISA) reader, at the wavelength of 540 nm. The results were expressed as variation of Optical density (OD)540 nm (OD540 nm sample - OD540 nm control). These OD values were considered as an index of bacteria adhering to surface and forming biofilms. For interpretation of biofilm production, the average of the three wells was calculated, and the criterion proposed by Stepanovic et al. [14] was adopted: non-adherent (OD < 0.12), moderate producer (0.12 < OD < 0.24) and strong producer (OD > 0.24).

3. Results

3.1 Antibiotic susceptibility

The S. aureus isolates in Biamba Marie Mutombo Hospital and from UTI were 100 % resistant to ampicillin-sulbactam, piperacillin-tazobactam, levofloxacin, and amoxicillin-clavulanic acid. With the exception for fosfomycin, netilmycin and amikacin, the resistance rates of clarithromycin, azithromycin, cefixime, ceftazidime, tobramycin, trimethoprim, and aztreonam to S. aureus was within the range 69 - 92 %. All Staphylococcus studied were MRSA and resistant to glycopeptide antibiotics, vancomycin and teicoplanin (Table 1).

Table 1: Antibiotic susceptibility pattern of S. aureus isolates from UTI and SSI

The 5 S. aureus strains isolated in Saint Joseph Hospital (Kinshasa) from SSI were highly resistant to ampicillin (100 %), ceftazidime (80 %), fosfomycin (100 %), amoxicillin + clavulanic acid (100 %), aztreonam (100 %), temocillin (80 %), erythromycin (100 %). All strains were MRSA. All MRSA strains were fully resistant to vancomycin and teicoplanin (Table 1).

In E. coli isolates, imipenem, cefixime, cefotaxime, ceftazidime, aztreonam, norfloxacin, temocillin, amoxicillin, ampicillin-sulbactam, and piperacillin-tazobactam resistance was observed in 100 % of cases. All Enterobacter sp. strains were fully resistant to imipenem, cefixime, temocillin, cefotaxime, aztreonam, amoxicillin, ampicillin-sulbactam, and piperacillin-tazobactam. E. coli and Enterobacter sp. strains demonstrated good sensitivity to fosfomycin. For other antibiotics, resistance was over 70 %, with the exception of amikacin (Table 2).

Table2: Antibiotic susceptibility pattern of Enterobacteriaceae isolates from UTI (Biamba Marie Mutombo Hospital)

The E. coli, Citrobacter sp., Enterobacter sp., Serratia sp. strains from SSI isolated in Biamba Marie Mutombo Hospital were highly resistant to the majority of antibiotics tested. E. coli isolates were particularly 100 % resistant to ampicillin, temocillin, kanamycin, amoxicillin – clavulanic acid, cefotaxime, and imipenem (Table 3).

Table3: Antibiotic susceptibility pattern of Enterobacteriaceae isolates from SSI Saint Joseph Hospital, Kinshasa

Multidrug resistance (MDR) was observed in Staphylococcus and Enterobacteriaceae isolated from UTI and SSI.

3.2 Detection of OXA-48-producing Enterobacteriaceae

Cultures in Chromatic^TM OXA-48 chromogenic medium revealed 48(87.2%) OXA-48 producers in general. All Enterobacteriaceae strains from SSI were OXA-48 producers (Table 4).

Table 4: OXA-48-producing Enterobacteriaceae strains

3.3 Biofilm formation

The results of biofilm formation of different clinical isolates studied are presented in Table 5).

Table 5: Biofilm phenotype of Enterobacteriaceae and S. aureus isolates from UTI and SSI

3.3.1 Enterobacteriaceae and S. aureus isolates from UTI

From the total number of 13 S. aureus isolates from Biamba Marie Mutombo Hospital and tested for biofilm formation, strong biofilm producers (SBP) were 4 (30.8%), 7 (53,8%) were moderate producers (MBP), and 2 (15,4%) were non- biofilm producers (NBP). Out of 10 E. coli tested for biofilm formation, 2 (20.0%) were SBP, 4 (40.0%) MBP, and 4 (40.0%) NBP. In E. cloaceae strains, 3 (33.3%) were SBP, 4 (44.5%) MBP, and 2 (22.2%) NBP (Table 5).

3.3.2 Enterobacteriaceae and S. aureus isolates from SSI

Among 5 S. aureus strains isolated from SSI in Saint Joseph Hospital and tested for biofilm formation, 4 (80.0%) were SBP, and 1 (20.0%) was NBP. Ten (52.6%), 9 (47.4%) of E. coli strains were SBP and MBP respectively. For a total of 9 Enterobacter sp. studied for biofilm formation, 6 (62.5%) were SBP and 3 (33.5%) were MBP. Five (66.7%) of Citrobacter strains have formed a strong biofilm and 3 (33.3%) have produced moderate biofilm. Out of 5 Serratia sp. strains, 3 (60.0%) were SBP and 2 (40.0%) were MBP (Table 5).

3.4 Resistance pattern of S. aureus and Enterobacteriaceae isolates among biofilm producers and non-biofilm producers

To determine whether biofilm formation was correlated with resistance to any particular antibiotic(s), we compared the biofilm forming capacities among isolates from UTI and SSI with different resistance profiles for the all antibiotics (Table 6 and 7).

Table 6: Biofilm formation and antibiotic resistance pattern Enterobacteriaceae and S. aureus isolates from UTI (Biamba Marie Mutombo Hospital

SBP: strong biofilm producers; MBP: moderate producers; NBP: non- biofilm producers; ND: not determined

Table 6 Continued: Biofilm formation and antibiotic resistance pattern of Enterobacteriaceae and S. aureus isolates from UTI (Biamba Marie Mutombo Hospital)

SBP: strong biofilm producers; MBP: moderate producers; NBP: non- biofilm producers; ND: not determined

Table 7: Biofilm formation and antibiotic resistance pattern of Enterobacteriaceae and S. aureus isolates from SSI (Saint Joseph Hospital)

Table 7 Continued: Biofilm formation and antibiotic resistance pattern of Enterobacteriaceae and S. aureus isolates from SSI (Saint Joseph Hospital)

SBP: strong biofilm producer; MBP: moderate biofilm producer; NBP: non-biofilm producer

Table 8: Occurrence of multidrug resistant pattern and their associations with biofilm phenotype in Enterobacteriaceae and S. aureus isolates from UTI (Biamba Marie Mutombo Hospital)

3.4.1 Enterobacteriaceae and S. aureus from UTI

For S. aureus isolates, resistance to oxacillin, ampicillin-sulbactam, amoxicillin-clavulanic acid, piperacillin-tazobactam, ceftazidime, cefixime, aztreonam, vancomycin, teicoplanin, levofloxacin, tobramycin, trimethoprim, clarithromycin, and azithromycin were higher in MBP and SBP than in NBP. Resistance to ampicillin-sulbactam; cefotaxime, cefuroxime, amoxicillin, piperacillin-tazobactam, ceftazidime, cefixime, imipenem, aztreonam, levofloxacin, norfloxacin, and tobramycin were higher in MBP and NBP than in SBP in E. coli isolates. Among Enterobacter cloaceae, resistance to ampicillin-sulbactam; cefotaxime, cefuroxime, amoxicillin, piperacillin-tazobactam, ceftazidime, cefixime, imipenem, aztreonam, levofloxacin, norfloxacin, amikacin, and tobramycin were higher in MBP and SBP than in NBP (Table 6).

3.4.2 Enterobacteriaceae and S. aureus from SSI

Among S. aureus isolates, resistance to oxacillin, ampicillin, amoxicillin-clavulanic acid, ceftazidime, aztreonam, vancomycin, teicoplanin, amikacin, levofloxacin, ciprofloxacin, trimethoprim, fosfomycin, erythromycin, and temocillin were notably high in SBP than in NBP. Resistance to ampicillin, amoxicillin-clavulanic acid, cefotaxime, amikacin, kanamycin, norfloxacin, and imipenem were higher in SBP than in MBP in E. coli isolates. Similar results were obtained for Enterobacter sp., Citrobacter sp., and Serratia sp. isolates (Table 7).

3.5 Occurrence of multidrug resistant pattern and their associations with biofilm phenotype

Regarding MDR, no relationships were found between the ability to form biofilm and antimicrobial resistance (Table 8 and Table 9).

4. Discussion

Enterobacteriaceae and Staphylococcus are known as a significant cause of infections in both community and nosocomial settings. The emergence of microorganisms resistant to multiple antibiotics used in the treatment of infections has become an important health problem worldwide, particularly in African countries [15]. The present study analyzed the resistance profile of pathogens involved in community and hospital acquiring infections and their capability to form and to produce a biofilm. The results showed an alarmingly increase of antibiotic resistance among Enterobacteriaceae and Staphylococcus aureus strains from UTI and SSI isolated in Biamba Marie Mutombo and Saint Joseph Hospitals.

All S. aureus isolates from UTI and SSI were MRSA. The results of studies conducted on S. aureus antibiotic resistance in Central Africa region are in concordance with the results of the present study. 82 % of S. aureus strains isolated from different clinical samples (wounds, urines, pus) were MRSA [16]. 100 % of these MRSA strains were also resistant to ceftazidime, cefotaxime, amoxicillin- clavulanic acid and cefixime as demonstrated in our study. Reports from Uganda showed MRSA prevalence of 57.2%, where 100% of MRSA strains resistant to amoxicillin-clavulanic acid, ceftriaxone, and imipenem (15). Another study from East Africa revealed an overall MRSA prevalence of 53.4% [17]). In contrast to our data, MRSA isolates from these last studies remained highly susceptible to teicoplanin and vancomycin [18, 19].

Our data demonstrates very high prevalence rates of antibiotic resistance of Enterobacteriaceae strains from UTI and SSI to ampicillin, imipenem, cephalosporins, ciprofloxacin, levofloxacin, norfloxacin, amoxicillin-clavulanic acid, amoxicillin, ampicillin-sulbactam, aztreonam, and tobramycin. These results are consistence with previous reports. In Nigeria, E. coli isolates demonstrated remarkable high rates of resistance to the β-lactam antibiotics, except the carbapenems and piperacillin-tazobactam. High resistance rates were also observed for E. cloacae against ampicillin (90%), aztreonam (80%), cefepime (70%), cefotaxime (80%), ceftazidime (60%), and cefuroxime (100%) (17). A study conducted in Rwandan referral hospital have demonstrated that out of 241 Gram-negative isolates tested for ceftriaxone, 183 (75.9%) were resistant [20].

In this study, we detected OXA-48-producing strains among different enterobacterial species isolated in samples from patients with UTI and SSI. The prevalence of 87.2% of OXA-48-producing Enterobacteriaceae observed in our study was higher than those obtained from studies conducted in some African countries, such as in a Nigerian hospital and in Tanzania with respectively 3.4 % and 4.9 % of OXA-48 producers among multidrug-resistant Enterobacteriaceae isolates [11,15]. Investigations done in many African countries such as Tunisia, Libya, Tanzania, Senegal, and Morocco, had shown that K. pneumoniae was the most frequently OXA-48 producer [10]. But in this study, we observed an emerging rate of OXA-48 producers among Enterobacter sp and Citrobacter sp strains (100%). In contrast, 22 of the 29 strains of E. coli were OXA-48 producers.

In this study the detection of biofilm formation was performed using Microtiter plate method. The results showed that 11 (84.6%) S. aureus, 6 (60%) E. coli, and 7 (77.7%) Enterobacter sp. isolates from UTI were biofilms producers. All Enterobacteriaceae and 4 (80.0%) S. aureus isolates from SSI were biofilm producers. Microbial cell adherence to surfaces and the development of multi-cellular communities is a key step in infection. Furthermore, bacteria biofilms can play a critical role in SSI and in in recurrent UTI [21, 22]. In this study the results showed that the capability of bacteria isolates to form a biofilm was very high in clinical strains from SSI than those from UTI. We demonstrated also a high variability in biofilm biomass production among isolates from UTI and SSI. Biofilm formation depends on many factors such as environment, sugar content and concentration (glucose versus lactose), geographical origin, types of specimen, surface adhesion characteristics, proteolytic enzymes, and biofilm associated genes [23 - 27]. These factors could be involved in the high prevalence of biofilm formation in bacteria strains from SSI as observed in the present study. Biofilm infections are clinically important because bacteria in biofilms exhibit recalcitrance to antimicrobial compounds. Microbes growing within a biofilm have been reported to be up to 1000 times more tolerant to antimicrobials than their planktonic counterparts [28]. The biofilm producing - Enterobacteriaceae and Staphylococcus aureus as well as non-biofilm producers from UTI were very resistant to antibiotics. Our results are in contrast with those obtained by Neaopane et al. in which 86.7% of biofilm-producing S. aureus were MDR; whereas all MRSA non- biofilm producers were non-MDR [29]. Our results are also in contrast with dose obtained by Neupane et al., [30]. In this last study authors showed that the antibiotic resistance of biofilm producing - E. coli was found significantly higher than that of biofilm non- producing E. coli. In our study 3 E. coli negative for biofilm formation were resistant to 12 different antibiotics (Table 7). Among biofilm producing-Enterobacteriaceae and S. aureus from SSI, higher antibiotic resistance was observed in strong and moderate biofilm producers. In this case, our results are in agreement with previous reports [26, 30]. Globally, the results of the current study are in agreement with report in which no relationship was observed between global resistance or MDR and biofilm formation [31].

Many factors could be responsible for the increasing of resistance in Kinshasa. Among them are some frequent societal behaviors (such as self-medication), inadequate healthcare infrastructure (insufficiently trained prescribers and inadequate diagnostic tools), and an uncontrolled drug sector (antibiotics sold over-the-counter, improperly stored, counterfeit, and/or expired [32] as well as biofilm ability of strains and the acquisition of resistance genes [33].

5. Conclusion

The alarming increase of S. aureus and Enterobacteriaceae isolates from Biamba Marie Mutombo and Saint Joseph Hospital to antibiotics limits the treatment of patients with UTI and SSI. The study showed that non- biofilm and biofilm producers were MDROs. The results of the present study showed that antibiotic resistance is a major public health problem that requires a range of urgent interventions. So, public health authorities should implement and develop comprehensive national policies and plans to prevent and combat the spread of MDROs in community and hospital setting.

Conflict of Interest

None

Acknowledgments

We thank Microbiology Laboratory staff members of Biamba Marie Mutombo and Saint Joseph Hospitals, Kinshasa, for their cooperation and technical assistance during the study.

Abbreviations

MDROs-Multidrug-Resistant Organisms; MRSA-methicillin-resistant Staphylococcus aureus; MDR- Multidrug resistance; OXA-oxacillinase; UTI-Urinary tract infection; SSI-Surgical site Infections, SBP-Strong biofilm producers; MBP-Moderate producers; NBP-Non- biofilm producers.

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