Biofilms are considered a significant reason for the failure of disinfection strategies in industrial water systems due to their resistance to antimicrobial agents. This study is designed to investigate the anti-biofilm activity of hydrogen peroxide (H2O2) at combinations of temperatures and contact times. For this purpose, an in vitro microtiter plate (MTP)-based model system was used for biofilm formation using Escherichia coli (E. coli) strain FL-Tbz isolated from the water system of a pharmaceutical plant. To investigate the anti-biofilm activity of H2O2, it was added at different concentrations (2–7% v/v) to biofilms and incubated at different temperatures (20–60 °C) for 10–40 min to find effective conditions to eradicate biofilms. Maximum biofilms were formed when bacterial suspensions were incubated at 37 °C for 96 h. The rate of biofilm formation using an environmental isolate was higher than that of standard strain. H2O2 at concentrations of ≥6.25% (v/v) at temperatures of ≥40 °C incubated for ≥25 min significantly eradicated the biofilms.

  • The time-saving and fairly cheap MTP-based model system is selected for the evaluation of the anti-biofilm activity of hydrogen peroxide.

  • Different behavior of pharmaceutical water system isolates of E. coli in in vitro biofilm formation can be seen.

  • H2O2 is an effective anti-biofilm agent in optimum concentration, temperature, and exposure time against E. coli isolated from a pharmaceutical water system.

Graphical Abstract

Graphical Abstract
Graphical Abstract

Biofilms, a population of microorganisms adhered to a biotic or abiotic surface, significantly contribute to biofouling in industrial water systems (Murthy & Venkatesan 2008). The formation of biofilm is a multistep process in which microbial cells initially attach to a surface reversibly (Gomes et al. 2014) followed by the encapsulation of cells within a self-produced matrix consisting of polysaccharides, proteins, and DNA (Stoodley et al. 2002; Sauer 2003; Coenye & Nelis 2010). Sessile cells, which are physiologically and phenotypically distinct from planktonic (non-adhered) cells, are formed by the association of cells embedded in the extracellular matrix. They are characterized by their enhanced resistance to antimicrobial treatments (Coenye & Nelis 2010). Therefore, the main reason for the failure of antimicrobial treatment in industrial water systems is biofilm generation, which poses significant challenges and has negative consequences (Gün & Ekİncİ 2009).

The biofilms periodically detach from the surface and enter the water, producing severe difficulties such as material damage, production losses, and deterioration of product quality (Murthy & Venkatesan 2008). It is advantageous and necessary to implement effective disinfectants to eliminate, control, and manage biofilms in industrial water systems.

The microtiter plate (MTP)-based system is commonly used to study the complex communities of biofilms due to many advantages. It is user-friendly and relatively cost-effective due to the need for small volumes of reagents (Niu & Gilbert 2004). Additionally, many tests may be conducted simultaneously using this method. MTP-based assays provide a simple way of assessing the influence of coating and material modification on biofilm formation. Therefore, this method is ideal for evaluating various agents' antibacterial and anti-biofilm properties (Ali et al. 2006).

Several disinfectants, including ozone and chlorine compounds (Loret et al. 2005), have been utilized to suppress biofilms in industrial water systems. Although microbial contamination and recolonization of water lines can be decreased or even prevented by these antimicrobial agents, they may have adverse effects on worker health, generate hazardous byproducts, and damage equipment (Di Pippo et al. 2018).

Hydrogen peroxide (H2O2) is a broad-spectrum disinfectant with bactericidal and sporicidal properties, which is effective against most chlorine-resistant bacteria (Linley et al. 2012). Importantly, H2O2 does not produce any byproducts or gases. It is fully soluble in water, and its safety depends on its concentration. It exerts its effects by generating radicals, such as the hydroxyl radical (OH), which react with cellular components such as lipids, proteins, and nucleic acids (Cloete et al. 1998; Perumal et al. 2014). H2O2's efficiency decreases against biofilms. Biofilm structures of exopolysaccharide matrix produce a protective barrier that inhibits antibiotics from reaching bacterial cells (Mah & O'Toole 2001; Gomes et al. 2014). Also, secreted alginate, catalases, and other free radical scavengers break down H2O2 and inhibit its impact on bacterial cells (Perumal et al. 2014).

The anti-biofilm activity of H2O2 on various microorganisms has been studied. Presterl et al. (2007) tested the effects of H2O2 on established biofilms of Staphylococcus epidermidis isolated from patients with cardiac implant infections. Perumal et al. (2014) investigated the efficacy of H2O2-based disinfectants on biofilms formed by Gram-negative pathogens, including Pseudomonas aeruginosa, Klebsiella pneumoniae, and Acinetobacter spp. Christensen et al. (1990) studied the effectiveness of low concentration H2O2 on biofilms formed by two marine bacteria. Also, the anti-biofilm activity of H2O2 against bacteria isolated from dental unit water lines (DUWL) has been studied. Lin et al. (2011) studied the effects of various concentrations of H2O2 on mature waterline biofilms and in controlling planktonic (free-floating) organisms in simulated dental treatment water systems. Orrù et al. (2010) evaluated the anti-biofilm activity of H2O2 on bacteria frequently reported in DUWL (including Pseudomonas spp., Streptococcus spp., and Staphylococcus spp.).

Escherichia coli (E. coli) is a potential water quality hazard in water systems. However, the efficacy of H2O2 was not reported on biofilm formed by E. coli isolated from industrial water systems. It has been proven that the environment can act as a reservoir of antibiotic-resistance genes, and isolated microbes' behavior can be highly different from standard bacteria (Jara et al. 2020; Tarhriz et al. 2020). Therefore, the efficacy study of various agents on biofilms formed by isolated bacteria from the water system of pharmaceutical plants helps generalize the laboratory data to real-world applications. To our knowledge, this is the first study evaluating the anti-biofilm activity of H2O2 on E. coli isolated from the water system of a pharmaceutical plant located in Tabriz, Iran, which can help to mimic the actual efficacy of H2O2 in pharmaceutical water systems.

Inoculum preparation

Water samples were collected from deionizing resins (sampling point) of the water system of the pharmaceutical plant in Tabriz, Iran. The samples were firstly cultured for 48 h at 37 °C in lactose broth medium (enriching microbes) followed by the passage of the growth microbes to MacConkey agar (selective medium for non-fastidious Gram-negative organisms) and incubated for 72 h at 37 °C.

According to the results of preliminary phenotypic and phylogenetic tests, the isolated bacteria belonged to the strain of E. coli, and its 16S rRNA sequences were submitted in GenBank with accession numbers OL798000 (Farjami et al. 2022). Several colonies from the agar plate were transferred to 10 mL of Trypticase Soy Broth (TSB) and incubated at 37 °C for 72 h. Furthermore, according to the United States Pharmacopeia, E. coli ATCC 8739 was used as standard bacteria in this study. Ten milliliters overnight culture of E. coli in TSB were harvested by centrifugation (6,000 g for 5 min), rinsed in sterile phosphate-buffered saline (PBS) of pH 7.2, resuspended to the optical density (OD) of 1.0 at 600 nm, and were used immediately to form biofilms using the MTP-based system. The supplemental file contains more details on the phylogenetic analysis and biochemical characterization of the isolated strain FLp-Tbz.

MTP assay

The formation of biofilm by E. coli grown in TSB was performed using MTP-based assays on 96-well tissue culture plates. Then 200 μL of bacterial suspension with an OD of 1.0 was added to six wells. Six wells with sterile TSB alone were served as controls. Incubation time and temperature are effective parameters in biofilm formation which need to be optimized. Plates were incubated at different temperatures of 25, 30, and 37 °C for 24, 48, 72, and 96 h to achieve maximum biofilm formation. After selecting optimum temperature and incubation time, biofilm formation was conducted using isolated E. coli bacteria from a deionizer.

The effect of albumin was investigated on promoting the adhesion of the bacterial cells to the surface of polystyrene wells (Ali et al. 2006; Wuren et al. 2014). For this purpose, before adding bacterial suspension to wells, 100 μL of albumin (Octalbin 25%, Swiss) was transferred to a U-bottom 96-well MTP and incubated for 24 h at room temperature without shaking. Then, the excess albumin was gently drained by a sampler.

Quantitation of biofilm by crystal violet staining

Before crystal violet (CV) staining, the contents of the MTP wells were emptied, and the wells were rinsed with 200 μL of sterile PBS to remove non-adherent bacteria. The cells adhered to wells were stained for 10 min with 200 μL of 0.05% (v/v) CV (Applichem, Canada), and 200 μL of 33% (v/v) acetic acid (Merck, Germany) was added to solubilize the dye bound to bacterial cells. The OD was measured at 600 nm using an MTP reader (BioTek, synergy HT, USA). Quantification of biofilms in each well section was done using six replica wells per experiment. Three independent experiments were carried out.

Study of the anti-biofilm activity of H2O2

Various parameters, including contact time, concentration, and temperature, affect the antimicrobial activity of disinfectants. The biofilms were incubated with 200 μL of H2O2 at concentrations of 2, 3.25, 4.5, 6.25, and 7% (v/v) for 10, 25, and 40 min to test the anti-biofilm activity of H2O2 (Merck, Germany). After selecting the optimum concentration of H2O2, the experiments were done at temperatures of 20, 40, and 60 °C. Six replicate wells were used per experiment, and three independent experiments were carried out.

Statistical analysis

P-values were compared for all three experimental groups, and p < 0.05 was considered statistical significance. The experiment results are expressed as the mean ± SD. Statistical analysis was performed by two-tailed unpaired Student's t-test or ANOVA as appropriate, followed by post hoc Tukey's test. All data were analyzed with GraphPad Prism Software vs. 6.0 (GraphPad Software, Inc.).

Optimization of MTP-based model for biofilm formation

According to the obtained maximum OD value (OD = 1.60 ± 0.06), incubation at 37 °C for 96 h caused the maximum formation of biofilm. It was selected as an optimum condition for biofilm formation using an MTP-based model system (Figure 1). A clear difference in the rate of biofilm formation between standard strain and isolated E. coli behavior in biofilm formation can be seen in Figure 2. Also, some controversial results were observed after adding albumin to wells containing the isolated bacteria compared with standard bacteria.
Figure 1

OD of E. coli biofilms formed by the MTP-based model system at different incubation times (24, 48, 72, and 96 h) and different temperatures (a: 25 °C, b: 30 °C, and c: 37 °C) using standard bacteria as well as isolated E. coli (d: 37 °C) in uncoated and coated wells with albumin. *Significantly different (comparison between with albumin and without albumin columns) (unpaired Student's t-test, p < 0.05).

Figure 1

OD of E. coli biofilms formed by the MTP-based model system at different incubation times (24, 48, 72, and 96 h) and different temperatures (a: 25 °C, b: 30 °C, and c: 37 °C) using standard bacteria as well as isolated E. coli (d: 37 °C) in uncoated and coated wells with albumin. *Significantly different (comparison between with albumin and without albumin columns) (unpaired Student's t-test, p < 0.05).

Close modal
Figure 2

Decrease in the OD of isolated E. coli biofilms compared with the control after H2O2 treatment at different concentrations of 2–7% for a contact time of 10–40 min at 20–60 °C. C: control (untreated biofilm). *Significantly different (one-way ANOVA followed by Tukey's test, p < 0.05).

Figure 2

Decrease in the OD of isolated E. coli biofilms compared with the control after H2O2 treatment at different concentrations of 2–7% for a contact time of 10–40 min at 20–60 °C. C: control (untreated biofilm). *Significantly different (one-way ANOVA followed by Tukey's test, p < 0.05).

Close modal

Anti-biofilm activity of H2O2

The effect of H2O2 is shown in Figure 2 at different concentrations, contact times, and temperatures on the in vitro biofilm model system. According to the data, biofilm was eradicated when treated with H2O2 at concentrations of ≥6.25% with temperatures of ≥40 °C incubated for ≥25 min.

Resistance of biofilms to antimicrobial compounds and the severity of problems caused by biofilms in industrial settings all over the world highlight the critical need to find effective and nontoxic biocides for biofilm control in industrial water systems (Murthy & Venkatesan 2008). In addition to production losses and negative impacts on the quality of the products, biofilms prevalent in industrial water systems are a health threat (Muhammad et al. 2020) and have a significant role in bacterial infections (Parsek & Singh 2003; Tarhriz et al. 2020). The release of cell clusters from the sloughing of biofilms results in the periodic release of bacteria in water systems, which causes infection (Hunt et al. 2004; Castonguay et al. 2006).

In this research, an in vitro MTP-based biofilm model system was applied to imitate the production of biofilms in water systems. Since we intended to evaluate the effect of H2O2 on the disinfection of the pharmaceutical water system (no added pollution), the interfering influence of the potential matrix was ignored.

According to the data (Figure 1), both standard and isolated E. coli exhibited a time- and temperature-dependent rise in biofilm development. In the case of isolated E. coli, however, the rate of biofilm production is much greater than that of standard E. coli, as the highest OD value of biofilms produced after 96 h at 37 °C was 1.60 ± 0.06 versus 0.96 ± 0.01 at 600 nm, respectively. Consistent with our result, Castonguay et al. (2006) revealed that laboratory strains of E. coli PHL565 have no substantial ability to generate biofilms. E. coli isolates from a water system had better adherence to glass than the E. coli PHL565 lab strain. The capacity of adhesion-proficient isolates to attach to glass is substantially associated with their ability to produce curli. It is claimed that curli is the key adhesion characteristic in environmental E. coli isolates.

Adding albumin to wells with isolated bacteria versus standard bacteria yielded controversial results. In the presence or absence of albumin, biofilm formation did not follow a pattern. In research by Garca-Bonillo et al. (2020), albumin enhanced biofilm formation on a hydrophobic urinary medical device. The biofilm model system was developed using an uropathogenic strain grown in LB broth and LB with agar. Due to the great affinity of bacteria for albumin structure, bacteria are securely entrenched in albumin or even integrated; consequently, they claimed that albumin is the primary parameter in bacterial attachment and the growth and maturation of biofilm (García-Bonillo et al. 2020). In another study, Hammond et al. (2010) found that albumin reduces P. aeruginosa biofilm development on plastic. They indicated that albumin works as a physical barrier, preventing biofilm development (Hammond et al. 2010). According to two research works (Hammond et al. 2010; García-Bonillo et al. 2020), albumin might have opposing effects (promotor or inhibitor) on biofilm development depending on surface type and microorganism. So, the albumin influence on in vitro biofilm development should be tuned according to strain type and experiment conditions.

E. coli monitoring as a water quality threat is the cornerstone of water microbiological management (Ram et al. 2008; Carrillo-Gómez et al. 2019). E. coli control by various chemical agents has been evaluated (Cloete et al. 1992, 1998; Loukili et al. 2004; Kauppinen et al. 2012; Azargun et al. 2020). Due to hazardous byproduct production and the inability to penetrate biofilms, chlorine's medicinal uses are restricted (Cloete et al. 1998). Also, ozone as a water disinfectant and an anti-biofilm agent has two fundamental limitations, such as instability in water and interaction with organic compounds, which creates low-molecular-weight oxygenated byproducts (Cloete et al. 1998).

Regarding the constraints previously mentioned, H2O2 is typically utilized as a low-toxic disinfectant to treat individual water sources without byproducts. It inhibits corrosion and the production of odors and colors by microorganisms and pollutant degradation. H2O2 disintegrates into water and oxygen, raising water oxygen levels (Linley et al. 2012). The emission of free oxygen radicals is H2O2’s disinfection mechanism.
formula

Higher temperatures and organic pollutants enhance this process. Free oxygen radicals with oxidizing and disinfecting powers decompose pollution, leaving only water. H2O2 at low quantities oxidizes cell components such as proteins, DNA, enzymes, and others, affecting a broad spectrum of species. H2O2’s efficiency decreases against biofilms. Biofilm structures of exopolysaccharide matrix produce a protective barrier that inhibits antibiotics from reaching bacterial cells (Mah & O'Toole 2001; Gomes et al. 2014). Also, secreted alginate, catalases, and other free radical scavengers break down H2O2 and inhibit its impact on bacterial cells (Perumal et al. 2014).

H2O2 is affected by peroxide concentration, reaction time, and temperature. Our findings (Figure 2) showed that increasing H2O2 concentration, temperature, and contact time decreased the OD of isolated E. coli biofilms and increased its efficacy against biofilm. Since H2O2 penetrates slowly into the biofilm matrix, as indicated by a reaction–diffusion interaction hypothesis (Lu 1996), contact time affects its anti-biofilm activity.

As indicated in pharmaceutical microbiology (Hugo & Russell 1998), H2O2 is suitable for general disinfection. Presterl et al. (2007) demonstrated that H2O2 at 3 and 5% (v/v) could eradicate S. epidermidis bacteria and clear biofilms formed by the MTP-based model system. Armon et al. (2000) showed that H2O2 (30 ppm) efficiently inhibited biofilm formation in drinking and wastewater pipes. In potable surface water distribution systems with residual H2O2 at 16.5 mg L−1, biofilm regrowth occurred the day following disinfection (Momba et al. 1998). Reducing residual disinfectant leads to biofilm growth. H2O2 may inhibit biofilm renewal owing to its long-lasting impact on microorganisms (Momba et al. 1998).

Our research indicated that H2O2 at concentrations of ≥6.25% with temperatures of ≥40 °C and contact time of ≥25 min eradicates E. coli biofilms. Lin et al. (2011) found no biofilm elimination even with 7% H2O2 for 24 h. Controversy may be related to the isolate type and experiment conditions (temperature and contact time). In another study, H2O2 at 6 and 10% (v/v) reduced Listeria monocytogenes in biofilm on food processing equipment (Yun et al. 2012). According to the above literature review, H2O2’s influence on biofilm development and anti-biofilm functions are significantly variable and need to be examined depending on isolate type, microbial flora of water, and real-world parameters like temperature and age of the distribution system.

The maximum biofilm formation was achieved by incubating E. coli isolated from the pharmaceutical water system for 96 h at 37 °C using the MTP method. The rate of biofilm formation was higher for the environmental isolate than for the standard strain. In an MPT-based system, H2O2 concentrations of 6.25% (v/v) and higher, a minimum incubation time of 25 min, and a temperature of 40 °C successfully eliminated E. coli biofilms. Biofilm control in water systems requires the prevention of biofilm regrowth. Therefore, in-house validation of disinfection procedures and periodic modification of the sanitization strategy play a crucial role in the biofilm management of water systems.

The authors acknowledge the Food and Drug Safety Research Center, Tabriz University of Medical Sciences.

S.J. and A.F. performed the experiments; F.L. and A.F. analyzed data; A.F., S.J., and M.S. prepared and wrote the manuscript; A.F. and M.S. edited the manuscript; F.L. designed the experiments; F.L. led and supervised the project.

There is no involvement of humans or animals in this study.

This work was supported by the Tabriz University of Medical Sciences.

All relevant data are included in the paper or its Supplementary Information.

The authors declare there is no conflict.

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