Original Article

Implications of Biofilm-Producing Organisms Among Bacteria Isolated From Ear-, Nose-, and Throat-Infected Patients

Please cite this article as follows: Ademakinwa OR, Adeyemo A, Ayodele SO, Nwamuo AS, Oluduro AO. Implications of biofilm-producing organisms among bacteria isolated from ear-, nose-, and throat-infected patients. Avicenna J Clin Microbiol Infect. 2024; 11(3):100-106. doi:10.34172/ajcmi.3557

Introduction

Antimicrobial resistance (AMR) is a significant public health threat, as admitted by the World Health Organization (WHO) (1). The extent of this AMR and the selection of multidrug-resistant (MDR) pathogens have been reported to be a result of non-compliance with proper infection control methods, unfounded use of antibiotics, availability of antibiotics without a prescription, and counterfeit products of dubious quality (2). Upper respiratory tract infections (URTI) are commonly treated with antibiotics based on the known sensitivity patterns of the common causative pathogens encountered in specific regions (3). Antimicrobial agents remain the backbone of infectious disease treatment; however, the unjustifiable uses of antibiotics in many countries have evolved into the emergence of MDR microorganisms (4). Infections become chronic, incalcitrant, or present with complications when treated with antibiotics that are not responsive to infective agents. The abuse of antimicrobial agents is well known to create pressure on the selection of appropriate drugs and resultantly increase the capability of microbes to restrain from being attacked (5). Antibiotic resistance leads to higher medical expenses, prolonged hospital stays, and increased mortality rates (2). Studies have reported an alarming increase in the occurrence of antibiotic-resistant bacteria that include beta-lactam-resistant strains of common pathogens as well as macrolides and fluoroquinolone-resistant strains isolated from URTIs (6,7).

An important element that contributes to AMR is biofilm production by bacteria. The formation of biofilm has been reported as one of the factors causing antibiotic resistance, particularly in URTIs. Bacterial biofilms are naturally resistant to antibioticsdue to the fact that some antibiotics are unable to reach the depths of the biofilm; some cells in biofilms grow slowly or not at all, likely due to nutrient limitation, and certain cells in the biofilms may adopt a unique and safeguarded biofilm phenotype (8).

Studies have identified bacterial biofilm as the main cause of antibiotic resistance in URTIs (9). Biofilm can form on moist biotic and abiotic surfaces, making them common for infection of the ear, nose, and throat (ENT). Bacterial biofilms are known to be “influencers of infections”, especially in patients suffering from rhinosinusitis and otitis media as a result of their propensity to form biofilms in sinuses and adenoid tissues (9–13). Most bacteria (more than 99%) produce biofilms, which can lead to dangerous, incurable illnesses. The bacteria in biofilms interact with each other through molecular mechanisms that enable some cells to resist antibiotics and host immune defenses, hence increasing the likelihood that ENT infections would recur or persist (13).

It has been estimated that at least 25% of cases of chronic rhinosinusitis are caused by biofilm formation. (14) Many microorganisms such as Streptococcus spp., Staphylococcus aureus, Corynebacterium argentoratense, and Micrococcus luteus present in the respiratory tract have been reported to easily produce biofilms. S. aureus strains have been identified intracellularly and in the sub-mucosa of adult patients with chronic rhinosinusitis undergoing endoscopic sinus surgery (15). Microorganisms such as S. aureus,Haemophilus influenza, Pseudomonas aeruginosa,and fungus have been isolated and identified as bacterial biofilms from chronic rhinosinusitis (9).

Despite the knowledge of bacteria as the major etiological agents of URTI and the roles of antibiotic resistance and formation of biofilms in the pathogenesis of URTI, local studies have not focused on biofilm formation potentials in bacteria causing ENT infections in West Africa. Therefore, this study seeks to establish the antibiotic sensitivity pattern and formation of biofilms in bacteria causing ENT infections in our study population and to determine the contribution of biofilm formation to MDR agents and the most effective choice of empiric antibiotics.

Materials and Methods

Sampling Population/Ethical Clearance

Participants with a clinical diagnosis of ENT infections attending the Otorhinolaryngology (ENT) Clinic in Obafemi Awolowo University Teaching Hospitals Complex (OAUTHC), Ile-Ife, Nigeria during the study period formed the study population. Ethical clearance (with protocol number ERC/2018/06/14) was duly obtained from the Ethical and Research Committee of the OAUTHC. In addition, informed consent was obtained from the patient or guardian of the patient as appropriate.

Sample Collection

ENT samples in the form of discharges, aspirates, and infected tissues were collected at the Otorhinolaryngology Clinic and Main Theatre of OAUTHC. Each sample was collected aseptically using sterile swab sticks or sterile bottles as appropriate by a medical doctor who has been pre-trained on the study protocol in the otorhinolaryngology clinic and operating theatre. The samples were collected into freshly prepared sterile transport media (thioglycolate media), properly labelled (with study number, date, gender, age, and time), and immediately transported for bacteriological analysis at the Microbiology Department of Obafemi Awolowo University, Ile-Ife, Nigeria.

Bacteria Isolation and Identification

The sample collected in the transport media was incubated over 24 hours at 37 ºC. The incubated culture was then inoculated separately on sterile blood agar, nutrient agar, MacConkey agar, and Mannitol salt agar (Lab M Ltd., UK) by the streak plate method for discrete colonies and incubated at 37 °C for 24 hours. The organisms were purified by successive subculturing on a nutrient agar plate. The isolates were identified by morphological and physiological characteristics according to Bergey’s Manual of Determinative (16). Furthermore, the isolates were identified by using the MICROBACT^TM identification kits 24E (Oxoid Ltd., UK) for Gram-negative, and the STPY gene was used to identify S. aureus using the polymerase chain reaction method.

Antibiotic Susceptibility Test and Biofilm Formation

The antibiotic susceptibility profile of the isolates was determined on Mueller-Hinton agar (MHA; Lab M Ltd., UK) according to Kirby-Bauer’s disc diffusion technique (17). The antibiotic discs, including single (Oxoid Ltd., Basingstoke, Hampshire, England) and combined (Abtek Biological Ltd., UK) discs of varying and specific concentrations, were employed and aseptically placed on the inoculated MHA plate with sterile forceps. The antibiotic discs were properly placed on MHA plates, seeded with standardized (10⁶ CFU/mL of 0.5 McFarland Standard) inoculum, and the plates were incubated at 37 °C for 18–24 hours, after which the diameter of zones of inhibition was compared with the Clinical and Laboratory Standard Institute (18) chart of interpretative zones as sensitive, resistance, and intermediate resistance. The isolates were described as resistant to multiple antibiotics when they were resistant to ≥ 3 separate classes of the tested antibiotics. The qualitative method for the biofilm formation of the isolates was performed using the tube method. A loop full of test organisms was inoculated in 10 mL of nutrient broth (Lab M Ltd., UK) with 1% glucose in test tubes. After incubation, the tubes were decanted and washed with the use of the phosphate buffer saline (pH = 7.3). The tubes were dried and then stained with crystal violet (0.1%). Deionized water was used to wash off excess stains, and the tubes were dried in an inverted position. The tube method was scored in line with the results from the control strains. As depicted in Figure 1, biofilm formation was considered positive when a visible film lined the wall and the bottom of the tube. No biofilm was produced when the tube was clear, implying that the wall and bottom of the tube were not lined by any visible film (19).

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Figure 1.

An Image of Biofilm-Producing Bacteria Recovered From ENT Infections. Note. ENT: Ear, nose, and throat

Figure 1.

An Image of Biofilm-Producing Bacteria Recovered From ENT Infections. Note. ENT: Ear, nose, and throat

Statistical Analysis

The data collected for analysis included patient sociodemographic details, previous exposure to antibiotics in current ENT disease, nature of the specimen, nature of isolates, antibiotic resistance, and biofilm production. Statistical Product and Service Solutions Statistics (version 22) was utilized to perform statistical analysis with the level of statistical significance set at P ≤ 0.05.

Results

One hundred and fifty samples, including ear (n = 87), nasal (n = 22) throat (n = 8), and ENT surgical aspirates and tissues (n = 33), were obtained from 150 patients [77 (51.3%) females and 73 (48.7%) males] diagnosed with various ENT infections. The age range of 0–5 years had the highest population in this study, followed by age ≥ 46, while 41–45 years had the lowest population. Table 1 provides a summary of the collected samples.

As outlined in Tables 2 and 3, a total of 15 different bacterial species were identified from 192 bacteria isolated from 150 samples [110 (73.3%) mono-bacterial were cultured from collected samples, and poly-bacterial culture was found in 40 (26.70%) samples]. These species were Proteus spp., P. aeruginosa, Klebsiella pneumoniae, Escherichia coli, Citrobacter freundii, Enterobacter cloacae, Serratia spp., Salmonella spp., Acinetobacter baumannii, Staphylococcus spp., Bacillus spp., Corynebacterium spp., Streptococcus spp., Micrococcus spp., and Enterococcus spp. Antibiotic susceptibility showed meropenem, vancomycin, and ofloxacin as the most effective antibiotics, while the isolates are more resistant to commonly used empirical antibiotics such as augmentin, cotrimoxazole, ceftazidime, cefuroxime, gentamycin, erythromycin, chloramphenicol, and ampicillin.

Based on the results (Table 4), 192 bacteria were recovered from collected samples, 113 isolates were isolated from ear infections, and 51 (45.1%) were multiple antibiotic resistant (MAR). In addition, 46 isolates were recovered from nose infection, and 20 (43.5%) were MAR. Twenty-three isolates were recovered from the throat, while 12 (52.2%) were MAR.