Influence of polymerized siloxane coating on growth and biofilm formation of aerobic grown nosocomial bacteria

1 Introduction

Health care-associated infections (HAI’s), or infections acquired in health-care settings are the most frequent adverse event in health-care delivery worldwide. In developed countries 7% of hospitalized patients at any given time, will acquire at least one HAI. In developing countries, case frequency increases onto 10% in hospitalized patients [1]. The main group of HAI’s are those related to the use of invasive devices. 50–70% of the HAI’s reported by the Centers for Disease Control in U.S. are caused by biofilm formation on indwelling medical devices [2, 3]. For this reason, the development of biomaterials and material coating substrates that prevent the formation of biofilms is of great importance to lower the risk of bacterial attachment and growth during and after implantation of a medical device. Recently an aqueous emulsion of polydimethylsiloxane (PDMS), which is a widely used silicone oil, was used as a coating substrate on stainless steel, polycarbonate and soda-lime glass. This coating proved to be non-cytotoxic to eukaryotic cells (fibroblasts), but significantly reduced adherence of aerobic nosocomial bacteria (Escherichia coli and Staphylococcus epidermidis) [4]. However, PDMS provides a hydrophobic surface, which is described to be less resistant to bacterial adhesion than hydrophilic surfaces [5, 6]. For this reason, a study was conducted to clarify whether PDMS can reduce the growth of nosocomial bacteria in a planktonic culture assay. Additionally biofilm growth was studied due to the different behavior of bacteria in planktonic culture and in biofilms [7].

2 Materials and methods

3 Results

3.1 Effects of PDMS on bacterial growth in planktonic cultures

In planktonic cultures with and without PDMS, the growth curves showed different phases of growth. E. coli almost reached the end of exponential growth or log phase in the examination time period of 300 min. For all bacteria tested, initial growth was retarded due to lag phase, followed by exponential growth. S.  epidermidis and especially P. aeruginosa had a slower growth rate than E. coli. After 300 min growth P. aeruginosa was still in the early exponential phase, while S. epidermidis has onward progressed in this phase. None of the bacteria reached the stationary phase. This allowed us to investigate the influence of PDMS on growing cultures and a comparison of effects during growth in a statistical end point analysis.

The impact of PDMS on the growth rate differed between the bacterial species. A dose-dependent effect of PDMS could be shown on E. coli and S. epidermidis as well as for 0.2 to 5 vol-% PDMS on P.  aeruginosa. Most commonly, higher concentrations inhibited growth stronger than lower concentrations, except for P. aeruginosa with 7 vol-% PDMS, which reduced bacterial growth less than 5 vol-%.

A 2 vol-% PDMS supplementation to liquid growth medium significantly reduced the growth rate of E. coli, P. aeruginosa and S. epidermidis (p < 0.001; see Fig. 1). Due to the reduced growth of every tested bacterial strain by the used siloxane concentration, we defined 2 vol-% as a sufficient and sensible effective concentration for further tests. After 300 min of incubation in presence of 2 vol-% PDMS the OD of the P. aeruginosa culture was reduced by 89.8% compared to the OD without PDMS (p < 0.001). S. epidermidis showed an OD impairment and growth reduction of 71.5% (p < 0.001) and E. coli exhibited an impairment of 35.0% (p < 0.001). In general, growth of P. aeruginosa was reduced the most, closely followed by S. epidermidis and followed by distance to both by E. coli.

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Fig.1

Growth of planktonic bacteria cultures in glucose nutrient broth with 0 vol-% (x), 0.2 vol-% (•), 2 vol-% (♦), 5 vol-% (▴) and 7 vol-% (■) PDMS supplementation; means of three individual tests with triple measurements, each.

With 7 vol-% supplementation of siloxane maximum growth reduction for S. epidermidis (97.4 %; p < 0.001) and E. coli (45.7%; p < 0.001). For P. aeruginosa we found a maximum growth reduction with 5 vol-% supplemented PDMS (125 %; p < 0.001), whereas bacterial growth with 7 vol-% was less reduced (94.7 %; p < 0.001).

We calculated negative values for optical density for P. aeruginosa and S. epidermidis cultures with 2, 5, and 7 vol-% PDMS, because of blanking against OD of equivalent pre-incubated broth-PDMS mixtures.

3.2 Effects of PDMS on bacterial single species biofilm formation and growth

Even without PDMS supplementation cell mass production of the used bacteria differed strongly. While for planktonic cultures the growth rate was the main determining factor, other biofilm-specific factors became important in this assay. Without PDMS exposition S. epidermidis showed the strongest biofilm growth within the time period of testing (18 hrs) followed by P. aeruginosa. Intensity of PDMS-free biofilm growth of E. coli and B. subtilis was similar and much weaker compared to S. epidermidis and P. aeruginosa (see Fig. 2).

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Fig.2

Growth of B. subtilis subsp. spizizenii, E. coli, P. aeruginosa and S. epidermidis after 18 hrs of biofilm culture on PDMS-coated/uncoated polystyrene, n = 8.

With PDMS exposition, biofilm growth of S. epidermidis and P. aeruginosa was again stronger compared to E. coli and B. subtilis. The impact of PDMS on biofilm production was significant and strongly dependent on the used bacteria. E. coli and P. aeruginosa showed increased biofilm formation on PDMS coated polystyrene, while S. epidermidis and B. subtilis in contrast showed significantly reduced biofilm formation. The bacterial cell mass in biofilms formed by S. epidermidis and B. subtilis was significant reduced by 64.7% (S. epidermidis, p < 0.001) and 23.1% (B. subtilis, p < 0.001) on the PDMS-coated surface, whereas cell mass in biofilms of P. aeruginosa and E. coli was 17.5% (P. aeruginosa, p = 0.011) and 10.1% (E. coli, p = 0.033) higher than without surface coating.

Surface coating for bacterial adherence and growth reduction [3, 10–12] is a promising approach to address the problem of health care-associated infections (HAI’s) which are related to invasive medical devices. A coating based on polymerized siloxanes and establishing a hydrophobic and positively charged surface already proved to reduce the number of adherent aerobic nosocomial bacteria on stainless steel, polycarbonate, and soda-lime glass [4]. The study showed that PDMS also affects the growth rate of aerobic nosocomial bacteria, which were grown planktonically, or in biofilms. This confirms results of a study from Yan and Li [13] which exposed S.  epidermidis and P. aeruginosa to silicon oil on blood agar and found PDMS to be antimicrobial. Comparable results were found with anaerobic bacterial strains (Propionibacterium acnes, Peptostreptococcus spp., Bacteroides fragilis, Fusobacterium spp., and Clostridium tertium) [14]. However, in our study different effects of PDMS on bacterial growth in the different culture assays were obvious.

Less intensive growth in tested planktonic bacteria strengthens the growth inhibition due to PDMS. The inhibitory effect of PDMS on bacterial growth seems to be strongly depended on growth rate. P. aeruginosa growth rate is typically slow and we could investigate the strongest inhibitory effect in planktonic cultures onto this bacterial culture. In contrast, growth of E. coli with (a typically) shorter generation time compared to the other bacteria used in this study could not be influenced that intensely.

In planktonic culture with addition of PDMS in different concentrations, even the weakest tested concentration of 0.2% PDMS led to a significant growth reduction of E. coli and P. aeruginosa. Starting from 2 vol-% PDMS growth of S. epidermidis was also significantly inhibited. Effect strength increased with increasing PDMS concentration, implying a dose dependent effect. An exception was P. aeruginosa incubated with 7 vol-% PDMS, because maximum effect for P. aeruginosa was found at 5 vol-% PDMS supplementation.

We calculated negative OD values for some initial points of time in PDMS supplemented cultures of P. aeruginosa and S. epidermidis. This was due to blanking against pre-incubated PDMS-broth mixtures of the same PDMS concentrations. Negative values are based on three additive factors: Slow growing cultures, cytotoxic effects, and putative changing of refraction index due to PDMS and bacterial cell interactions. Temporarily decline of growth curves can be an indicator for loss of bacterial cell integrity. PDMS seems to induce cell death due to cytotoxic effects, delays reproduction and reduces maximum growth rate respectively, as it was shown for silver nanoparticles in E. coli [15]. For P. aeruginosa a cytotoxic effect based on silver that diffused from coating to bacteria was presumed [16].

Biofilms and planktonic cultures of P. aeruginosa differ in gene expression and therefore metabolism like other bacteria, too [17]. Due to optimal growth and reproduction conditions for single cells, metabolism of planktonic log phase cultures is increased compared to biofilms. The physiological state of cells can lead to considerable variation in the receptiveness of bacteria to bactericides [18]. Reduction of viability seems to take place during active metabolic states like growth and cell division. Biofilm-associated tolerance is different from resistance displayed by planktonic cultures of bacteria, because of modified gene expression, which reduces metabolism and growth rate [19]. Therefore, it can be supposed that biofilm specific factors influenced the effect of PDMS-coating on bacterial growth. It is relevant to mention, that we only have examined single-species biofilms. Presumably, combinations of various species like in natural biofilm communities can lead to improved resistance [20].

Traditionally, biofilm formation is divided into at least three steps: Phase of primary attachment, accumulative phase, and disassembly of biofilm structure [21]. Physical forces like van der Waals forces, steric and electrostatic interactions, bacterial appendages like flagella, pili or fimbriae, as well as adhesive surface structures are essential for initial adherence to surfaces and following biofilm formation [22]. S.  epidermidis harbors adhesins like the polysaccharide intercellular adhesin (PIA) which is required for enforced biofilm accumulation [21]. Due to hydrophobic and van der Waals interactions it adheres strongly to hydrophobic surfaces like plastics [23]. Autolysin AtlE, a bacterial peptidoglycan-hydrolase, is also of importance for binding on unmodified polystyrene [21]. These characteristics enabled S. epidermidis to be the most potent biofilm producer on plastic surface in our study, and explain why S. epidermidis is one of the most common cause of biofilm-associated opportunistic infections on indwelling medical devices [24]. However, S.  epidermidis biofilm growth was most severely affected by 2 vol-% PDMS which led to a reduction of growth for more than 60% compared to the growth without PDMS. In contrast, 2 vol-% PDMS caused an increased biofilm growth of E. coli and P. aeruginosa while P. aeruginosa still exhibiting the stronger growth of both. These results are confirmed by the study of Ozdamar et al. [25] which also approved S. epidermidis less resistant to PDMS than P. aeruginosa. They addressed the effect of 10 vol-% silicone oil on different bacteria and after 12 hrs of growth on brain heart infusion agar at 35°C, S.  epidermidis and P. aeruginosa showed an apparent decrease in the number of CFUs, whereby S.  epidermidis was able to survive for only 5–7 days and P. aeruginosa for 7–10 days.

In P. aeruginosa at least three polysaccharides (alginate, Pel and PsI) mainly determinate stability of the biofilm structure [22, 26]. Alginate interacts with water and nutrients and supplies nutrients to cells in biofilm [26]. Additionally it reduces the access of toxins like antibiotics to the bacteria and blocks host immune reactions [27].

Strains of E. coli are genotypic and phenotypic heterogeneous [28]. Biofilm formation is mediated through motility by flagella, attachment and microcolony formation by type 1 fimbriae, curli and exopolysaccharides like PGA and colonic acid, antigen 43 as well as other factors for later biofilm maturation [29]. Increased biofilm growth after PDMS exposition might have been caused by the large number of hydrophobic amino acid residues on E. coli flagella, which makes the surface of E. coli very hydrophobic promoting contact of E.  coli to hydrophobic surfaces like polystyrene and other polymers [30, 31].

B. subtilis is a robust model organism for examination of biofilm formation of Gram-positive bacteria that manifests in highly structured floating pellicles in liquid cultures [32, 33]. However, laboratory ‘domesticated’ strains of B. subtilis are known to form weaker biofilms in contrast to environmental strains [32]. Also in our study, B. subtilis ssp. spizizenii was a weak biofilm producer. PDMS coating reduced biofilm formation significantly but not as powerful as in S. epidermidis. Additionally it is known that in submerged cultures of B. subtilis BL53 PDMS enhances oxygen supply improving cell metabolism and growth [34]. Therefore the activity of PDMS as oxygen carrier might have been advantageous for the resistance of B. subtilis against the adverse effects of PDMS.

5 Conclusion

PDMS had a significant impact on growth of nosocomial bacteria in planktonic and biofilm culture assays. In planktonic cultures, PDMS exposition resulted in a decreased growth of all tested bacteria in a dose-dependent manner in the used 300 min assay. This antimicrobial effect of PDMS was strongly related to the type of bacteria. Biofilm culture resulted in a modified growth behavior and PDMS resistance, whereby the growth of E. coli and P. aeruginosa were not adversely affected any more by PDMS as in the planktonic assay. However, these results are based on tests with single species biofilms. Previous to practical application it is necessary to confirm these results by tests in which different bacteria are able to interact like in natural biofilms, because this can cause modified bacterial growth and resistance [20]. Furthermore, research on the interactions on the molecular level could provide more insight into how antimicrobial effects can be explained and exploited.