The effect of diluted 1% baby shampoo on biofilm reduction in chronic rhinosinusitis with nasal polyposis

Study population

This study received the Research Ethical Committee, Universiti Kebangsaan Malaysia (REC UKM) approval with the approval number of JEP-2016-510. All patients underwent ESS for CRSwNP within the study period of 2 years who fulfilled both the inclusion and exclusion criteria were recruited into the study. The inclusion criteria were age 18 years old and above, who underwent ESS for CRSwNP. Patients who have sinonasal tumour and/or diagnosed with granulomatous nasal diseases were excluded from the study. Written informed consent in accordance to institutional guidelines was taken from patients who agreed to be involved in this study.

Methodology

Two sections of the samples of nasal polyp were obtained from each patient and samples were sent to the histopathology (for one section) and microbiology lab (for another section) as per laboratory protocols on the same day (Fig. 1).

Histopathology lab metholology

H&E on Biofilm. One section was stained with H&E as protocol modified from Hochstim et al. (2010). The section was deparaffinized in xylene twice for 5 min (2 × 5 min) and rehydrated with successive 1-minute washes in 100%, 96%, 80%, and 70% ethanol. Subsequently, the specimen was stained with haematoxylin for two minutes, rinsed with distilled water, rinsed with 0.1% hydrochloric acid in 50% ethanol, rinsed with tap water for 15 min, stained with eosin for one minute, and rinsed again with distilled water. The slide was dehydrated with 95% and 100% ethanol, successively followed by xylene twice for five minutes (2 × 5 min) and mounted with coverslips.

Biofilm identification for H&E. Slides were examined and evaluated by two researchers. Presence of irregularly shaped groupings of small basophilic bacterial clusters, one third of the size of the surrounding epithelial or inflammatory cells, a biofilm seen over the epithelial lining, not in or under the epithelial lining, biofilm seen tightly adherent to the surface epithelium or pulled away slightly; or a dense extracellular polysaccharide substance (EPS) material with embedded basophilic bacteria, occasionally entrapped erythrocytes, leukocytes and partially sheared from epithelial surface substance coating the epithelial surface, indicating biofilm-positive (Tóth et al., 2011) (Fig. 2).

Gram staining of the samples. Gram staining of the samples following the protocols of Gram stain kit Bio-Optica 04-100802 was employed to complement the histopathological examination (HPE) findings of biofilm (Mangels, Cox & Lindberg, 1984, Bortholomew, 1962).

Biofilm identification criteria. Histological criteria to denote the presence of biofilms were the morphologic features of biofilms on H&E staining as mentioned above, Gram positive or Gram negative and microcolonies seen in Gram staining, and the presence of surrounding polysaccharide layer.

Microbiology lab methodology

Tissue culture plate assay. Another section of the tissue specimens obtained from the operating theatre were immediately transported to the microbiology lab. The tissue culture plate assay employed in this study was based on the method described previously by Ha et al. (2008). The tissue specimens were homogenized. Specimens were inoculated on blood agar, MacConkey agar, mannitol salt agar and cetrimide selective agar, and incubated at 37 °C for 24 h. Pure colonies from the isolates above were then inoculated on to nutrient agars until further testing. Yellow colonies grew on mannitol salt agar were subjected to deoxyribonuclease (DNase) and coagulase test. DNase positive and coagulase positive colonies were considered as S. aureus. Green-pigmented colonies which grew on cetrimide agar were subjected to the API 20 NE test to confirm the identification of P. aeruginosa. Three to four colonies of bacterial isolates were suspended in Mueller-Hinton broth until the turbidity matches a standard of 0.5 McFarland and were incubated aerobically at 37 °C for 24 h. One mL of the bacterial suspension was diluted 1:100 with fresh Mueller-Hinton broth (10⁶ colony forming unit (CFU) mL). One mL of this 1:100 bacterial suspension was added to 1ml of sterile Mueller-Hinton broth to achieve a final concentration of 5 × 10⁵ CFU/mL. Reference strain of biofilm-forming S. aureus ATCC 25923 and P. aeruginosa ATCC 27853 obtained from the American Tissue Culture Collection (ATCC) were used as a positive control in this study. Control strains were cultured with the same method. Biofilm presence of the tissue specimen was confirmed using H&E and Gram staining. Only tissues with biofilm and culture positive for S. aureus or/and P. aeruginosa were included in the analysis. The organisms were tested for biofilm-forming properties.

Three 96-well tissue-culture treated microplates were filled with 200 µL of Mueller-Hinton broth and 10 µL of the final inoculums (5 × 10⁵ CFU/mL) and were incubated at 37 °C for 8 days without agitation for biofilm growth. 50 µL of media were removed from each well and replaced with 50–150 µL of fresh media on alternate days to achieve maximal biofilm growth. After incubation for 8 days, the wells were washed twice with distilled water to remove planktonic forms of bacteria and were left to dry. One of the 96-well tissue-culture treated microplates was filled with 250 µL of methanol and left for 15 min for fixation of adherent bacteria. Subsequently methanol was removed, and the microplate was air dried for another 15 min. Subsequently 200 µL of 0.1% crystal violet (CV) solution was applied to stain the adherent bacteria for 5 min. Then, the CV solution was decanted, and the wells were washed three times with distilled water. The stained wells were filled with 250 µL of 95% ethanol and were incubated for 1 h on a rocking platform at room temperature to solubilize the adherent material.

Quantification of biofilm formation. Quantification of biofilm growth was determined by reading the optical density (OD) of each well at 595 nm (OD595nm) using a microplate reader. Reference strains were used as control. Negative control wells were filled with only sterile broth. Test was carried out in a triplicate manner, and the mean OD was recorded. A cut-off optical density (ODc) needs to be established to note the presence or absence of biofilm. The cut-off optical density (ODc) was defined as 3 standard deviations above the mean OD of the negative control (culture medium): ODc=average OD of negative control+(3 ×SD of negative control). The mean OD of the strains will be compared to the OD of the negative control. Analysis was done as described by Stepanović et al. (2007), whereby strains were classified as follows: OD ≤ ODc no biofilm producer, ODc<OD ≤ 2 × ODc weak biofilm producer, 2 × ODc<OD ≤ 4 × ODc moderate biofilm producer and 4 × ODc<OD strong biofilm producer.

Preparation of diluted 1% baby shampoo dilution

An amount of 0.1 ml of Johnson & Johnson baby shampoo was diluted in 10 mls of normal saline to achieve a 1% concentration which was used as nasal irrigation (Joss et al., 2016). This concentration was determined and adapted based on previous literature (Chiu et al., 2008). Solution was prepared 30 min prior to test.

Biofilm reduction testing

Once the biofilm had been established and confirmed on the first microplate on day 8, wells on the second and third microplates containing the same inoculums were filled with 100 µL of 1% baby shampoo diluted in normal saline, and normal saline separately. The microplates were incubated for another 24 h at 37 °C. Positive control wells contained only the inoculums and the media without the test component, and negative control wells contained only test component and media, without the inoculums. The microplates were analysed using a crystal violet assay and optical density and measured with a microplate reader as described earlier. A flow chart of the methodology is shown in Fig. 1. All tests were carried out in a triplicate fashion. The relative inhibition of biofilm was calculated as described by Ha et al. (2008), as follows: ${\displaystyle \mathbf{Percentage of biofilm inhibition}=100-\left[O{D}_{595}\text{of treated well}/O{D}_{595}\text{of untreated well}\right]\times 100.}$

Statistical analysis

Data was entered and analysed using SPSS 25.0. Wilcoxon signed-rank test was employed to analyse the reduction of biofilm using diluted 1% baby shampoo and normal saline only, and to compare biofilm reduction between normal saline and diluted 1% baby shampoo. Fisher’s exact test was used to determine the statistical difference in biofilm formation and biofilm forming capacity of S. aureus and P. aeruginosa. p < 0.05 was considered as statistically significant.

Demographic data

A total of 60 samples were collected from 30 patients who underwent ESS for CRSwNP. The age ranged from 26 to 78 years, with a mean ± standard deviation (SD) of 58 ± 15 years. There were 21 male and nine female which comprised of 70% and 30% respectively, and 13 (43.3%) of them were Malays, 13 (43.3%) were Chinese and four (13.3%) were Indian. All patients were treated with intranasal corticosteroids and normal saline nasal douche. Half of the population underwent repeated ESS.

Data on biofilms

Biofilms were detected in 13 out of 60 samples (21.7%) using H&E complimented with Gram stain method (Fig. 2). Out of these 13 samples, 23% (n = 3) comprised of P. aeruginosa, 15% (n = 2) of S. aureus, 54% (n = 7) had no bacterial growth and 8% (n = 1) belonged to other bacteria (Table 1). All samples which grew S. aureus (n = 2) or P. aeruginosa (n = 3) were bio-film-positive. There was no sample which grew S. aureus and P. aeruginosa simultaneously. There was statistical significance in biofilm formation of S. aureus or P. aeruginosa with the p-values of 0.0002 (p < 0.05) (Table 2).

Biofilm forming capacity

All samples which grew S. aureus or P. aeruginosa with biofilm seen on H&E staining were subjected for tissue culture plate assay to determine the bacteria biofilm forming capacity. They were all graded after a cut-off point of optical density of the negative control was determined. The biofilm forming capacity categorizes the isolates into either a non-, weak-, moderate-, or strong biofilm producer, based on the optical density reading as described in the methodology. All samples with P. aeruginosa (100%) isolates were strong biofilm forming capacity, whereas one sample with S. aureus had strong biofilm forming capacity and one weak biofilm forming capacity (50%). There was no statistical difference in biofilm forming strength between S. aureus and P. aeruginosa with the p-value of 0.4.

Effects of diluted 1% baby shampoo on biofilm mass

The five samples with biofilms (three of P. aeruginosa and two of S. aureus) were then tested for biofilm reduction using normal saline alone, and with a diluted 1% baby shampoo. The mean reduction in biofilm was 26.87% with normal saline alone and 57.31% after treatment with diluted 1% baby shampoo (Table 3). There was significant reduction of biofilm with the treatment of diluted 1% baby shampoo with the p-value of 0.043, however there was no significant reduction of biofilm with normal saline with the p-value of 0.080 (Table 4). Overall analysis diluted 1% baby shampoo was significantly more effective in biofilm reduction than normal saline with the p-value of 0.043 (p < 0.05) (Table 4, Fig. 3).

Study limitations and suggestions

The primary limitation of this study is the limited availability of financial resources, which may have influenced methodological design and strategy. Biofilm detection was conducted without CSLM or FISH due to their high cost, although we acknowledge that these methods may enhance biofilm detection. Nevertheless, in our study biofilm was successfully identified using H&E staining and light microscopy, supported by Gram staining. The methods are supported by previous studies, which have demonstrated that biofilm detection using H&E staining is reliable in CRS patients (Hochstim et al., 2010; Hong et al., 2014; Tóth et al., 2013). Furthermore, the methods are easily reproducible and cost effective, making it suitable to be used particularly in centers without CSLM and FISH services. We selected only the two most commonly found bacteria biofilms in CRSwNP based on previous studies, due to the high cost of ATCC strains. As a result, the reductions of biofilms using diluted 1% baby shampoo of the other biofilms was not investigated.

We suggest using CSLM method with BacLight methods or FISH analysis with species-specific oligonucleotides probes in the next study on biofilm, which is currently most widely used for biofilm detection and to speciate the organisms involved. Sampling methods should be improved by taking samples from all involved paranasal sinuses in view of the reported heterogenicity of organism community in the nasal cavity. We recommend a prospective multi center studies in the future to evaluate the effectiveness of diluted baby shampoo 1% in biofilm reduction comparing populations of urban and rural areas to achieve improved tailored management in our local population.