Successful treatment of biofilm infections using shock waves combined with antibiotic therapy

Abstract

Many bacteria secrete a highly hydrated framework of extracellular polymer matrix on suitable substrates and embed within the matrix to form a biofilm. Bacterial biofilms are observed on many medical devices, endocarditis, periodontitis and lung infections in cystic fibrosis patients. Bacteria in biofilm are protected from antibiotics and >1,000 times of the minimum inhibitory concentration may be required to treat biofilm infections. Here, we demonstrated that shock waves could be used to remove Salmonella, Pseudomonas and Staphylococcus biofilms in urinary catheters. The studies were extended to a Pseudomonas chronic pneumonia lung infection and Staphylococcus skin suture infection model in mice. The biofilm infections in mice, treated with shock waves became susceptible to antibiotics, unlike untreated biofilms. Mice exposed to shock waves responded to ciprofloxacin treatment, while ciprofloxacin alone was ineffective in treating the infection. These results demonstrate for the first time that, shock waves, combined with antibiotic treatment can be used to treat biofilm infection on medical devices as well as in situ infections.

Figure 1. Biofilm formation on urinary catheters.

(a–d) Scanning electron micrographs (SEM) of bacterial biofilms on sections of urinary catheter. The micrographs were taken at the same magnification and the scale bar in (d) is 10 μm. (a) Untreated catheter surface. (b) Catheter surface with S. Typhimurium biofilm. (c) P. aeruginosa biofilm. (d) S. aureus biofilm. (e) Crystal violet staining of biofilms caused by the different bacteria on equal amounts of catheter substrate. Statistical significance was calculated using One-way ANOVA. Asterisks indicate statistical significance as follows: (***p < 0.001). Error bar–mean ± SD. (f) Photograph of hand held shock wave generator. (g) Schematic of hand held shock wave generator. The igniter is used for the ignition of the polymer tube by a spark generated by electrodes, explosive coating undergoing combustion and the combustion flame front travelling at 2000 m/s, and shock waves emanating from the open end of the polymer tube.

Figure 2. Effect of shock waves on biofilm formation.

(a–f) SEM of bacterial biofilms grown on plastic microfuge tubes. (a) S. Typhimurium biofilm. (b) P. aeruginosa biofilm. (c) S. aureus biofilm. (d) S. Typhimurium biofilm after shock wave. (e) P. aeruginosa biofilm after shock wave. (f) S. aureus biofilm after shock wave. (g) Crystal violet staining of biofilms before and after shock wave treatment. (h–k) SEM of biofilms on urinary catheters before and after shock waves. (h) P. aeruginosa biofilm. (i) S. aureus biofilm. (j) P. aeruginosa biofilm after shock wave. (k) S. aureus biofilm after shock wave. (l) The area covered by the biofilm estimated from examining 50 SEM fields. Scale bar in 3f & 2k–10 μm. Statistical significance was calculated using One-way ANOVA. Asterisks indicate statistical significance as follows: (***p < 0.001). Error bar–mean ± SD.

Figure 3. Antibiotic sensitivity of bacteria released by shock wave treatment.

Catheter sections with biofilms of P. aeruginosa or S. aureus formed in bovine (a) or human (b) urine were washed and placed in PBS. After shock wave exposure, the PBS was plated to check the release of the bacteria from the biofilm. The control histograms were not treated with a shock wave. Catheter sections with biofilms formed in bovine (c) or human (d) urine were also treated with a shock wave, or left untreated, then incubated with 4 μg/ml ciprofloxacin. After 6 h, the catheter samples was washed, sonicated in a bath sonicator to release all bacteria, and plated in LA to estimate the number of viable bacteria. Control samples in c and d were exposed to neither shock waves nor ciprofloxacin. Statistical significance was calculated using One-way ANOVA. Asterisks indicate statistical significance as follows: (***p < 0.001). Error bar–mean ± SD.

Figure 4. The diaphragmless shock wave generator and the pressure profile.

(a) Photograph of diaphragmless shock wave generator/Diaphragmless shock tube (DST). The length of the driven section is 6.064 m. The L- bend which attenuated the shock wave by 30%. The mice were kept in the perforated chamber (insert) during shock wave treatment. (b) Schematic and working principle of DST. Pressure signal at the end of the shock tube (c,e) and inside the perforated chamber (d,f) was measured by mounting a pressure transducer at the respective places.

Figure 5. Use of shock wave adjunct therapy in the treatment of murine P. aeruginosa lung infection.

BALB/c mice were infected with agarose beads coated with P. aeruginosa for 3 days before treatment with a daily shock wave, with and without daily ciprofloxacin therapy for a further 3 days. (a–e) The mice were killed and the lungs examined by SEM. (a) Uninfected mice. (b) Mice infected with P. aeruginosa. (c) Mice infected with P. aeruginosa and treated with ciprofloxacin (2.5 mg/kg, via intravenous delivery). (d) Mice infected with P. aeruginosa and treated with shock wave therapy alone. (e) Mice infected with P. aeruginosa and treated with a shock waves and ciprofloxacin. Scale bar in 4E–10 μm. (f) The number of bacteria in the lungs were determined by plating the homogenized lung tissue after 3 days of treatment. The Statistical significance was calculated using Mann-Whitney test. Asterisks indicate statistical significance as follows: (*p < 0.01), (***p < 0.001), ns–not significant. Error bar–mean ± SD. (g) The survival of mice infected with P. aeruginosa for 3 days, then treated with ciprofloxacin with and without shock waves for 5 days (day 0 in g). The animals in g were examined twice daily. The Statistical significance was calculated using Log-rank test. Asterisks indicate statistical significance as follows: (***p < 0.001)

Figure 6. Use of shock wave adjunct therapy in the treatment of murine suture associated S. aureus infections.

BALB/c mice were provided abdominal sutures after 3 days the sutures were removed. SEM of untreated sutures revealed that sterile sutures did not develop obvious biofilms (a). (b) SEM of Sterile suture treated with shock wave. Mice were also given sutures where the surgical wire was pre-incubated with S. aureus (c-f). After 3 days the mice were killed and the sutures examined by SEM. The excised S. aureus biofilm-coated sutures were left untreated (c), treated ex vivo with ciprofloxacin alone (4 μg/ml for 6 h) (d), shock wave therapy alone (e) or a combination of ciprofloxacin and shock wave (f). Scale bar in e–10 μm. (g) The bacteria were enumerated in the suture with or without any treatment by homogenizing by ultrasonication and plating. Statistical significance was calculated using One-way ANOVA. Asterisks indicate statistical significance as follows: (**p < 0.005), ( p < 0.001). Error bar–mean ± SD.

Figure 7. Use of shock wave adjunct therapy in the treatment of murine suture-associated S. aureus infections.

BALB/c mice were provided abdominal sutures and the sutures were subsequently infected with S. aureus. The mice were left untreated (control), treated with shock wave therapy alone, ciprofloxacin alone (i.v. 2.5 mg/kg daily) or a combination of ciprofloxacin and shock wave. After 3 days of treatment, the sutures (a) and surrounding skin (b) were removed independently and homogenised and the bacteria were enumerated by viable count. The Statistical significance was calculated using Mann-Whitney test. Asterisks indicate statistical significance as follows: (**p < 0.005). Error bar–mean ± SD.