In Vitro Biofilm Formation Kinetics of Pseudomonas aeruginosa and Escherichia coli on Medical-Grade Polyether Ether Ketone (PEEK) and Polyamide 12 (PA12) Polymers

Abstract

Biofilms, structured communities of microorganisms encased in an extracellular matrix, are a major cause of persistent infections, particularly when formed on medical devices. This study investigated the kinetics of biofilm formation by Escherichia coli and Pseudomonas aeruginosa, two clinically significant pathogens, on two medical-grade polymers: polyether ether ketone (PEEK) and polyamide 12 (PA12). Using a modified crystal violet staining method and spectrophotometric quantification, we evaluated biofilm development over time on polymer granules and catheter segments composed of these materials. Results revealed that PEEK surfaces supported significantly more biofilm formation than PA12, with peak accumulation observed at 24 h for both pathogens. Conversely, PA12 demonstrated reduced bacterial adhesion and lower biofilm biomass, suggesting surface characteristics less conducive to microbial colonization. Additionally, the study validated a reproducible protocol for assessing biofilm formation, providing a foundation for evaluating anti-biofilm strategies. While the assays were performed under static in vitro conditions, the findings highlight the importance of material selection and early prevention strategies in the design of infection-resistant medical devices. This work contributes to the understanding of how surface properties affect microbial adhesion and underscores the critical need for innovative surface modifications or coatings to mitigate biofilm-related healthcare risks.

1. Introduction

Biofilms are structured communities of microorganisms, primarily bacteria, that adhere to surfaces and are encased within a self-produced extracellular polymeric substance (EPS) matrix [1]. This matrix comprises polysaccharides, proteins, lipids, and nucleic acids, providing structural integrity and protection to the microbial community [2]. Biofilm formation is a dynamic process involving initial attachment, microcolony formation, maturation, and eventual dispersion [3]. Within biofilms, microorganisms exhibit altered phenotypes, including changes in growth rates and gene expression, which contribute to their resilience in various environments [4].

The physiology of biofilms confers several advantages to the resident microorganisms. The EPS matrix serves as a physical barrier, protecting cells from environmental stresses, including desiccation, ultraviolet radiation, and antimicrobial agents. Additionally, the proximity of cells within the biofilm facilitates horizontal gene transfer, including the spread of antibiotic resistance genes [5]. Nutrient gradients within the biofilm create microenvironments that support diverse metabolic activities, allowing for the survival of different microbial species under varying conditions (Figure 1).

Biofilms pose a significant threat to human health, particularly when they form on medical devices such as catheters, prosthetic heart valves, and pacemakers [4]. The biofilms on these devices can lead to persistent infections that are difficult to treat due to their inherent antibiotic resistance and ability to evade the host immune response [6]. For instance, biofilm-associated infections are associated with conditions such as cystic fibrosis and chronic wounds. Biofilm’s resilience necessitates the development of novel strategies to prevent its formation and effectively manage biofilm-related infections in clinical settings [5,7].

Polyether ether ketone (PEEK) and polyamide 12 (PA12) are high-performance polymers with exceptional properties that are extensively utilized in medical devices [8,9,10] (Figure 2). PEEK is a semi-crystalline thermoplastic known for its high mechanical strength, chemical resistance, and biocompatibility. Its molecular structure consists of repeating units of ether and ketone linkages between aromatic rings, contributing to its rigidity and thermal stability [9]. PA12 is part of the nylon family and is characterized by its flexibility, low moisture absorption, and good chemical resistance. Its structure comprises repeating units of amide linkages, which balance toughness and flexibility [10].

PEEK’s biocompatibility and radiolucency in medical applications make it suitable for implantable devices, such as spinal fusion cages and dental implants. Its mechanical properties resemble human bone, reducing stress shielding and promoting better integration [11]. PA12 is commonly used in medical tubing, catheters, and other components that require flexibility and durability. Its low moisture absorption ensures dimensional stability, which is crucial for precision medical applications [12].

Combining the unique properties of these polymers enables their application in various medical devices, thereby enhancing performance and patient outcomes. Advancements in processing techniques, such as additive manufacturing, have further expanded their use, enabling the production of complex, patient-specific medical components [8,9].

In a previous dental study of biofilm formation analysis using PEEK, a comparison was made with materials such as titanium and zirconium [13]. The methodology used was able to reveal the formation of biofilms; however, the study’s objective was to detect antimicrobial activity, which could not be determined. To decrease the interaction between bacteria and PEEK, various approaches have been proposed, including the incorporation of therapeutic and bioactive agents into the PEEK matrix or on its surface, the development of PEEK coatings, and the addition of reinforcing agents to produce nanocomposites or mixtures. However, the kinetics of biofilm formation or its quantification were not determined. The species studied were Escherichia coli and Staphylococcus aureus [13].

Escherichia coli and Pseudomonas aeruginosa were selected as representative bacterial models due to their clinical relevance and well-established ability to form biofilms on abiotic surfaces. Both species are among the most frequently isolated pathogens in nosocomial infections associated with medical devices, including catheters, prosthetic implants, and ventilators. P. aeruginosa, a Gram-negative opportunistic pathogen, is known for its robust ability to form complex, multidrug-resistant biofilms, especially in immunocompromised patients. Similarly, E. coli, particularly the uropathogenic and extraintestinal strains, is a leading cause of device-associated infections such as urinary tract and bloodstream infections. Their distinct adhesion mechanisms, biofilm maturation pathways, and tolerance to antimicrobial agents make them ideal models for evaluating how surface properties of biomaterials influence microbial colonization and biofilm development. Using these two organisms enables a comprehensive assessment of the interaction between clinically essential pathogens and polymeric materials commonly used in the manufacture of medical devices.

This study experimentally assessed biofilm formation on polymeric surfaces commonly used in medical devices. Insights into how P. aeruginosa and E. coli interact with PEEK and PA12 enhance the understanding of how surface properties influence microbial adhesion and biofilm growth.

2. Materials and Methods

Strains and growth conditions: Escherichia coli W3110 [F− lam-In (rrnD–rrnE)1 rph-1] and Pseudomonas aeruginosa PAO1 were used throughout this study. For long-term storage, glycerol stocks were generated from freshly grown cultures and kept at −80 °C. Microorganisms were cultivated in Luria–Bertani (LB) broth (OXOID Becton & Dickinson, Franklin Lakes, NJ, USA) [14]. Cells were grown at 37 °C [15]. For overnight (ON) cultures, an isolated colony from an LB agar plate (OXOID Becton & Dickinson, Franklin Lakes, NJ, USA) was transferred into 3 mL of LB medium and incubated at 37 °C [16].

Biofilm determinations: Biofilm formation was assessed using 96-well polyvinyl chloride (PVC) microplates (Costar, Cambridge, MA, USA) filled with 150 µL of LB medium and inoculated with overnight cultures standardized to OD₆₀₀ = 0.01. Plates were incubated at 37 °C in a sealed plastic container to prevent evaporation [17,18,19,20,21]. Following incubation, wells were gently washed, stained with 1% crystal violet (Merck, Darmstadt, Germany) for 20 min, rinsed with distilled water, and air-dried. The retained dye was solubilized in 150 µL of 40% acetic acid (Merck, Darmstadt, Germany) for quantification at OD₅₅₀. Comparable results were obtained using either distilled water or fresh LB for rinsing. All assays were performed in triplicate across three independent experiments, with standard deviation consistently below 20%. For biofilm kinetics, specific wells were sampled and stained at defined time points, while others remained at 37 °C. Before use, PVC plates were UV-sterilized for 15 min, and uninoculated wells served as controls for contamination.

To assess biofilm formation on PEEK and PA12 (VESTAKEEP^® (https://products.evonik.com/assets/35/91/VESTAKEEP_Compounds_EN_EN_243591.pdf, accessed on 29 June 2025) and VESTAMID^® PA12 (https://www.vestamid.com/en, accessed on 29 June 2025), 1 cm³ polymer granules were UV-sterilized for 15 min and confirmed to be non-toxic to E. coli and P. aeruginosa, as no growth differences were observed in their presence (Figure 3). Each well received one granule and 200 µL of a 1:5 dilution of the overnight culture. Controls included wells without granules, LB with granules, and LB alone. At 2 h intervals, granules were washed, stained with crystal violet (15 min), rinsed, dried, and the stain eluted in 150 µL of 40% acetic acid. OD₅₅₀ was measured and expressed in arbitrary units, normalizing time zero to 1. Experiments were done in triplicate, yielding consistent results, and the SD was recorded. For validation, PA12 and PEEK catheter segments (1.5 cm) were aseptically cut, placed in 24-well PVC plates, and analyzed using the same biofilm quantification protocol.

3. Results and Discussion

3.1. Validation of the Protocol for Measuring Biofilm Formation

Biofilm formation by the laboratory strains was first validated under the experimental conditions described (see Section 2) [16,20,22]. Biofilm formation was quantified in arbitrary units, taking 1 unit as the amount of biofilm formed at time zero. It is essential to note that cell growth in the culture followed the expected sigmoidal curve (Figure 3). The results are consistent with previously reported findings, demonstrating that E. coli and P. aeruginosa strains exhibit a pattern of maximum biofilm formation during the exponential growth phase and the early stage of the stationary phase [17]. Later, there is a phase of little biofilm formation and detachment of the already formed biofilm, which implies that it is not cell density that encourages biofilm formation, but a complex system of molecular signaling [20]. To verify whether PEEK or PA12 beads had any effect on bacterial growth in LB, the development of E. coli and P. aeruginosa cultures was monitored in the presence or absence of the beads. As shown in Figure 3, the growth curves in the presence or absence of the beads are virtually indistinguishable, indicating that the presence of either PEEK or PA12 does not affect the growth of E. coli or P. aeruginosa cultures.

3.2. Biofilm Formation on PEEK or PA12 Beads

E. coli and P. aeruginosa formed biofilms on PEEK and PA12 surfaces, as confirmed by crystal violet staining and OD₅₅₀ quantification. No growth was detected in negative controls. The kinetics of biofilm development differed slightly between polymers and species but followed reproducible patterns, enabling comparative analysis of biofilm formation on each material.

As an illustrative example, a representative experiment with E. coli is presented in Figure 4. Cultures were incubated in PVC microplate wells containing PA12 and PEEK beads to evaluate biofilm formation. At designated time points, the culture medium was removed, and the PA12 and PEEK beads were thoroughly rinsed with running water to eliminate non-adherent, planktonic cells. Although the beads appeared visually unaltered following rinsing, subsequent staining with crystal violet, a dye that binds to biofilm matrix components and adherent cells, revealed varying degrees of biofilm retention. Differential dye accumulation on the bead surfaces indicated heterogeneity in biofilm formation. The retained dye was eluted using 40% acetic acid to obtain a semi-quantitative measure of biofilm biomass, and the OD₅₅₀ was determined using a spectrophotometer.

This method provided an indirect yet reliable assessment of the extent of biofilm development on the PA12 and PEEK surfaces. It is essential to note that the polymer beads are irregularly shaped, with an average volume of approximately 1 cm³. Therefore, the experiments were performed at least three times in triplicate. The trend was very similar, as the standard deviation never exceeded 20%.

The kinetics of biofilm formation by P. aeruginosa and E. coli over 48 h on the two polymeric substrates, PA12 and PEEK, were determined (Figure 5). As a positive control, biofilm development was also measured in standard PVC wells. For E. coli, biofilm formation was seen on both PA12 and PEEK surfaces, reaching maximum levels at 12 h on PA12 and at 24 h on PEEK. The formation pattern in PVC wells closely resembled that on PEEK. Notably, a decrease in biofilm biomass was seen after 24 h, consistent with previously reported trends in PVC, likely reflecting the start of dispersal mechanisms [17]. In contrast, P. aeruginosa showed much less biofilm formation on PA12, suggesting that specific surface properties of this material may inhibit adhesion or biofilm development. On PEEK, however, a peak in biofilm formation was observed at 24 h, followed by a decrease at 48 h, a pattern similar to that seen with E. coli. These results indicate that PEEK supports more substantial biofilm formation than PA12 under the tested conditions, possibly due to differences in surface traits such as hydrophobicity, roughness, or chemical makeup. Additionally, the patterns over time suggest that biofilm maturation and the start of dispersal processes may occur after 24 h of incubation.

3.3. Biofilm Formation on the Surface of a Medical Device Made of PEEK or PA12

Catheters are essential medical devices that significantly improve patient care. Their use has expanded due to their vital roles in drug delivery, nutritional support, blood sampling, and dialysis, especially in patients with critical or chronic conditions [21,23,24].

An estimated 250 million intravascular catheters are used annually in Mexico and the United States [25,26]. However, roughly 3.5% of these devices become colonized by bacterial or fungal pathogens, leading to catheter-related bloodstream infections (CRBSIs), which are both severe and costly [27,28]. CRBSIs pose a significant concern in intensive care units, contributing to increased patient morbidity and mortality, with fatality rates reported between 19% and 34% [26]. These infections often originate from microbial colonization of the catheter surface during insertion, leading to the formation of biofilm, a key factor in the persistence and resistance of infections [29,30,31]. To prevent such complications, catheters often need to be removed and replaced. The most cost-effective way to reduce CRBSIs is to prevent bacterial adhesion and subsequent biofilm development on the catheter surfaces [29]. To explore this, two commercially available catheters made from different materials, including nylon, polyethylene, polyurethane, and PA12, as well as a capillary catheter containing PEEK, were used in further studies [30,31]. An ideal catheter should be flexible, disposable, radiopaque, and stretchable without losing structural integrity. After testing the raw polymer materials based on previous findings, bacterial adhesion and biofilm formation were evaluated on the catheter materials ready for use, to analyze the biofilm development kinetics under controlled experimental conditions (Figure 6).

To evaluate the ability of E. coli and P. aeruginosa to form biofilms on clinical catheter materials, a time-course experiment was performed using commercially available catheters made of PA12 or PEEK. Segments of the delivery tubing (which typically comes into contact with the human body and is maintained for extended periods) were cut and placed into the wells of a PVC microplate (Figure 6).

The results, shown in Figure 6, reveal a rapid rise in biofilm biomass during the initial incubation, reaching a peak around 24 h, except for the E. coli culture on the PA12 catheter material, where the highest biofilm formation was observed at 12 h. This early stage likely corresponds to bacterial adhesion and the formation of microcolonies. Afterwards, all curves plateau, indicating that the biofilm has reached a mature, stable state with limited further growth. Interestingly, a consistent decrease in biofilm detection amounts is observed at 36 h, possibly due to nutrient depletion, biofilm dispersal, or cell death within the aging biofilm. These findings indicate that there is a critical period within the first 12 h, during which catheter surfaces become colonized by biofilm-forming pathogens, such as E. coli or P. aeruginosa. This highlights the importance of preventing early adhesion in reducing catheter-associated infections. Additionally, this model provides a reproducible and measurable method for assessing anti-biofilm coatings or disinfection protocols under controlled conditions. It is noteworthy that the biofilm-forming behavior of P. aeruginosa was very different when using pure PA12 polymer (Figure 5) than when using the ready-to-use catheter made of the same material. This implies that other polymers used in the PA12 catheter may increase bacterial adhesion. This catheter is composed of nylon, polyethylene, polyurethane, and PA12.

The findings presented have significant implications for designing and applying polymeric materials in medical devices. The observation that E. coli and P. aeruginosa form more robust biofilms on PEEK compared to PA12 suggests that specific surface characteristics of PEEK, such as hydrophobicity, surface energy, or microtexture, may promote microbial adhesion and extracellular polymeric substance (EPS) accumulation. These results underscore the significance of material selection in clinical settings where the risk of biofilm-related infections is heightened, such as with catheters, implants, and tubing systems. Furthermore, they emphasize the necessity of modifying these materials or applying specialized coatings to inhibit microbial colonization.

This study also established a foundation for future investigations. Subsequent research could evaluate the impact of surface treatments, such as sulfonation, plasma etching, or the integration of antimicrobial agents, on biofilm development on PEEK and PA12. Conducting experiments under clinically relevant conditions, such as using human serum, mixed-species biofilms, or within dynamic flow systems, would enhance translational applicability. Expanding the analysis to include additional clinically applicable pathogens, such as Staphylococcus aureus or Candida albicans, may offer a more comprehensive understanding of biofilm dynamics on medical-grade surfaces.

A notable limitation of the current study is that assays were conducted under static laboratory conditions, which may not accurately represent the dynamic physiological environments encountered in vivo. Nevertheless, the biofilm quantification method demonstrates strong consistency and reproducibility, rendering it a valuable tool for assessing material susceptibility to microbial colonization. The outcomes underscore the critical importance of detecting and managing biofilm formation on medical devices, given the substantial role these microbial communities play in persistent infections, device malfunction, and increased healthcare expenditures. Advancements in early detection methods and surface engineering strategies will be crucial in mitigating biofilm-associated risks in clinical settings. Finally, a significant finding is the difference in P. aeruginosa biofilm formation on pure PA12 granules compared to catheters already infused with other polymers. This initial study will need to be complemented in the future by analyzing each component of the already manufactured catheters.

4. Conclusions

This study demonstrates that E. coli and P. aeruginosa, two clinically relevant pathogens associated with nosocomial infections, can form biofilms on medical-grade polymers PA12 and PEEK. Using a modified crystal violet staining method and spectrophotometric quantification, we were able to track the kinetics of biofilm development on these surfaces over time. Our results reveal distinct patterns of biofilm formation for each organism and material, highlighting that PEEK supports significantly higher biofilm accumulation than PA12 under the tested conditions. This aligns with earlier observations that reported PEEK as a surface prone to biofilm formation. A study in that context reported a bacteria-resistant coating consisting of a hydroxyapatite film on which ionic silver is immobilized via inositol hexaphosphate chelation, achieved through a series of immersion and drying steps performed at low temperatures [32].

The ability to detect and quantify biofilm formation on clinically used polymer surfaces provides critical insights into the potential risk of infection associated with medical devices made from these materials. Given the reproducibility and sensitivity of our experimental protocol, this approach can be further utilized to investigate the interactions between bacteria and various biomaterials, ultimately contributing to the development of infection-resistant medical devices and improved patient outcomes in hospital settings. Future experiments will include testing other species of bacteria and conducting experiments under conditions more closely resembling those found in nosocomial settings.