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Antibiofilm Activity and Synergistic Inhibition of Staphylococcus aureus Biofilms by Bactericidal Protein P128 in Combination with Antibiotics Sandhya Nair, Srividya Desai, Nethravathi Poonacha, Aradhana Vipra, Umender Sharma GangaGen Biotechnologies Pvt. Ltd., Yeshwantpur, Bangalore, India

P128 is an antistaphylococcal protein, comprising a cell wall-degrading enzymatic region and a Staphylococcus-specific binding region, which possesses specific and potent bactericidal activity against sensitive and drug-resistant strains of Staphylococcus aureus. To explore P128’s ability to kill S. aureus in a range of environments relevant to clinical infection, we investigated the anti-S. aureus activity of P128 alone and in combination with standard-of-care antibiotics on planktonic and biofilm-embedded cells. P128 was found to have potent antibiofilm activity on preformed S. aureus biofilms as detected by CFU reduction and a colorimetric minimum biofilm inhibitory concentration (MBIC) assay. Scanning electron microscopic images of biofilms formed on the surfaces of microtiter plates and on catheters showed that P128 at low concentrations could destroy the biofilm structure and lyse the cells. When it was tested in combination with antibiotics which are known to be poor inhibitors of S. aureus in biofilms, such as vancomycin, gentamicin, ciprofloxacin, linezolid, and daptomycin, P128 showed highly synergistic antibiofilm activity that resulted in much reduced MBIC values for P128 and the individual antibiotics. The synergistic effect was seen for both sensitive and resistant isolates of S. aureus. Additionally, in an in vitro mixed-biofilm model mimicking the wound infection environment, P128 was able to prevent biofilm formation by virtue of its anti-Staphylococcus activity. The potent S. aureus biofilm-inhibiting activity of P128 both alone and in combination with antibiotics is an encouraging sign for the development of P128 for treatment of complicated S. aureus infections involving biofilms.

S

taphylococcus aureus is responsible for causing a variety of community-acquired and hospital-acquired infections in humans all over the world. A significant number of clinical isolates of S. aureus have evolved to become resistant to commonly used antibiotics. The emergence of hospital- and community-associated methicillin-resistant S. aureus (MRSA) has worsened the situation further (1). Resistance has also been reported against both recently introduced and last-resort drugs used for treatment of S. aureus infection, such as vancomycin, daptomycin, and linezolid (2, 3). Thus, there is an urgent need to develop new therapies against this important human pathogen. S. aureus is known to form biofilms, and the bacteria residing in biofilms have been shown to be highly resistant to the action of antibiotics (4). Thus, under clinical conditions where biofilms play a role in pathogenesis, including wounds in diabetic patients and endocarditis, treatment failures are frequent despite the long duration of many treatments (5). The phenotypic resistance of bacteria residing in biofilms has been attributed to multiple factors, including both the biofilm matrix acting as a permeability barrier and the presence of slow-growing bacteria with poor metabolic rates, known as persisters (6–8). The majority of conventional drugs have been shown to have poor antipersister activity (9). Thus, drugs which show potent bactericidal activity on nonreplicating or slowly replicating persisters are expected to be more effective at eradicating biofilms (7). To this end, alternate approaches for killing bacteria in biofilms are being investigated (10). Included among these are bacteriophages and phage-derived proteins (enzybiotics) which have been found to kill bacteria in biofilms, thus offering a viable alternative (11). An S. aureus-specific bacteriophage has been shown to be efficacious in an in vivo animal infection model involving biofilms (12). Many enzybiotics do not require metabolically active bacteria for inhibitory action

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and thus can effectively kill nonreplicating bacteria (13, 14). In fact, one of the most commonly used assays for measuring the bactericidal activity of enzybiotics is the optical density (OD) fall assay, which measures lysis of bacteria in a buffer solution (15). Lysostaphin, the earliest known cell wall hydrolase, was shown to eradicate the biofilms formed by either S. aureus or S. epidermidis strains which were resistant to the standard-of-care (SoC) antibiotics oxacillin and vancomycin (16). A number of phage-derived lysins have shown potent bactericidal activity on S. aureus biofilms (17). Despite demonstrations of efficacy in vitro and in various animal models, only a few phage lysins have progressed to evaluation in clinical trials (18). P128, which incorporates a phage tail-associated muralytic enzyme (TAME) possessing antistaphylococcal activity, is currently under testing in a clinical trial (ClinicalTrials.gov identifier NCT01746654) for clearance of S. aureus from the nasal surfaces of patients, including chronic kidney disease patients who carry S. aureus in the nares. P128 possesses potent antistaphylococcal activity against sensitive and drug-resistant strains of S. aureus grow-

Received 26 May 2016 Returned for modification 20 June 2016 Accepted 18 September 2016 Accepted manuscript posted online 26 September 2016 Citation Nair S, Desai S, Poonacha N, Vipra A, Sharma U. 2016. Antibiofilm activity and synergistic inhibition of Staphylococcus aureus biofilms by bactericidal protein P128 in combination with antibiotics. Antimicrob Agents Chemother 60:7280 –7289. doi:10.1128/AAC.01118-16. Address correspondence to Umender Sharma, [email protected]. Supplemental material for this article may be found at http://dx.doi.org/10.1128 /AAC.01118-16. Copyright © 2016, American Society for Microbiology. All Rights Reserved.

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Antibiofilm Activity of P128

TABLE 1 Strains used in this study Strain

Source

MRSA status

S. aureus BK1

Public Health Research Institute (PHRI), NJ Gulbarga University, India ATCC ATCC (ATCC 700699) PHRI, NJ Kamineni Hospital, Hyderabad, India ATCC ATCC ATCC (ATCC 29212) ATCC (ATCC BAA-47)

MRSA

S. aureus B9241 S. aureus ATCC 29213 S. aureus Mu50 S. aureus MW2 (BK31) S. aureus B9356 S. aureus ATCC 8325-4 S. epidermidis ATCC 12228 Enterococcus faecalis V583 Pseudomonas aeruginosa PAO1 a

GMRSA MSSA MRSA MRSA MRSA MSSA MSSEa

Methicillin-susceptible S. epidermidis.

ing as planktonic cells or in biofilms (14, 19, 20). The mechanism of killing of staphylococci by P128 involves cleavage of the pentaglycine cross bridge of peptidoglycan (21). The absence of a pentaglycine in species other than staphylococci makes P128 inactive on other bacteria. P128 is equally active on bacteria growing in media and bacteria under conditions of nonreplication and nutrient starvation. The lack of inhibitory activity on bacteria other than staphylococci and on eukaryotic cells (14, 22) predicts it to be a safe drug candidate for treating human infections involving staphylococci. In order to further explore the utility of P128 to treat serious, difficult-to-treat infections caused by S. aureus, such as bacteremia, infective endocarditis, catheter-associated infections, and chronic diabetic wounds, we tested the antistaphylococcal activity of P128 in combination with SoC drugs on planktonic cells and biofilms. P128 was found to kill staphylococci in biofilms in a rapid manner, and importantly, it was highly synergistic with antibiotics in killing S. aureus in biofilms. P128 could also prevent biofilm formation in a multispecies model mimicking biofilm formation in chronic wounds (23). The potent antibiofilm activity of P128 and its synergy with SoC antibiotics make it a good candidate for further development for treatment of biofilm-associated S. aureus infections. MATERIALS AND METHODS Bacterial strains and culture conditions. The strains used in this study are listed in Table 1. S. aureus cultures were routinely grown in Trypticase soy broth (TSB) or LB broth or agar at 37°C. MIC and drug combination studies by checkerboard assays. MICs were determined using a modified Clinical and Laboratory Standards Institute (CLSI) broth microdilution procedure as described earlier (19, 24). In order to determine the effects of combinations of P128 with antibiotics, combinations of various dilutions of P128 and a second drug were tested for growth inhibition by use of a microdilution checkerboard technique (25). Briefly, an S. aureus culture at a final concentration of 5 ⫻ 105 CFU/ml was added to wells of 96-well microtiter plates (precoated with 0.5% bovine serum albumin [BSA]) containing 2-fold dilutions of P128 and the second drug in cation-adjusted Mueller-Hinton broth (CAMHB). The plates were incubated at 35°C for 24 h, and the individual MICs and combination MICs were read. The fractional inhibitory concentration index (FICI) was determined using the following equation: FICI ⫽ (MIC of drug A in the combination/MIC of drug A alone) ⫹ (MIC of drug B in the combination/MIC of drug B alone). The combination was considered to be synergistic when the FICI was ⱕ0.5, additive when the FICI was 0.5 to 1.0, indifferent when the FICI was 1 to 4, and antagonistic when the

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FICI was ⱖ4. The experiments were performed in triplicate and repeated twice. SEM of P128-treated biofilms. MRSA strain BK1 or MW2 was used to study the biofilm eradication activity of P128 by scanning electron microscopy (SEM). For this purpose, biofilms formed in microtiter plates or on the surfaces of catheters were treated with increasing concentrations of P128 or antibiotics and then subjected to SEM. (i) Biofilm formation and drug treatment of biofilms in microtiter plates. S. aureus BK1 or MW2 was revived on blood agar plates, and the culture was grown overnight in TSB supplemented with 2% glucose. The overnight culture was diluted 1:50 in TSB with 2% glucose to achieve an optical density at 600 nm (OD600) of ⬃0.1. Two hundred microliters of the diluted culture was aliquoted into each well of a 96-well plate and incubated at 37°C under static conditions for 18 h. After incubation, the contents were aspirated and washed twice with 200 ␮l of phosphate-buffered saline (PBS), 100 ␮l of medium and 100 ␮l of drug were added to each well, and the plate was incubated again at 37°C for 18 h. The contents were aspirated, washed with 200 ␮l of PBS, and allowed to dry at 37°C for 15 min. The individual wells were cut out, and biofilms were visualized by SEM. (ii) Biofilm formation and drug treatment of biofilms on catheters. To study the efficacy of P128 on biofilms on catheters, an overnight culture of S. aureus MW2 was diluted 1:40 in TSB containing 5% rabbit plasma. Catheter (JMS Infusion set) pieces of 1 to 2 cm were cut, sliced into two halves, and added to the culture. The cultures with catheter pieces were incubated at 37°C with shaking at 100 rpm for 24 h. Postincubation, the catheters were removed and rinsed twice in PBS to remove the adhering planktonic cells. The biofilms on catheters were challenged with 1 and 8 ␮g/ml of P128, 90 ␮g/ml of vancomycin, or 10 ␮g/ml of daptomycin (with 50 ␮g/ml CaCl2) by transferring the catheter pieces to tubes containing the drugs. The tubes were incubated at 37°C under static conditions for 18 h. The catheters were then removed from the tubes, rinsed once in PBS, and immersed in 0.1% safranin for 5 min. The stained catheters were washed once in PBS and allowed to dry. After drying, samples were fixed on aluminum stubs with double-sided carbon adhesive tape, coated with 5- to 7-nm-thick gold by use of a sputter-coating system (Q150T; Quorum Technologies), and examined by SEM (Neon 40; Carl Zeiss) for the presence of biofilm structures. For CFU enumeration, the catheters were removed from the tubes, rinsed twice in PBS, and placed in Eppendorf tubes containing 1 ml PBS. To release the adhered biofilm into PBS, the catheters were scraped with an inoculation loop. The samples were vortexed thoroughly and plated on LB agar plates. Crystal violet staining method to determine the effect of P128 on S. aureus biofilms. The protocol described previously by Schuch et al. (15) was followed. Briefly, S. aureus strains grown on blood agar plates were inoculated into TSB with 2% glucose, and the cultures were grown overnight. The cultures were diluted 1:50 in TSB to achieve an OD600 of ⬃0.1. Two hundred microliters of the diluted culture was added to 1.8 ml of TSB aliquoted into each well of a 24-well plate (⬃5 ⫻ 105 CFU). The plates were kept at 37°C under static conditions for 18 h, the contents were aspirated, the wells were washed twice with 2.0 ml of 1⫻ PBS, and 1 ml of medium plus 1 ml of the drug was added to each well and incubated at 37°C for 0, 2, 4, and 24 h. TSB supplemented with 50 ␮g/ml CaCl2 was used for daptomycin treatment wells. The contents were aspirated out at the stipulated time points, and the wells were washed with 2 ml of 1⫻ PBS. The wells were allowed to dry at 37°C for 15 min and stained with 1 ml of 1% crystal violet (CV) for 5 min. The wells were washed with 1 ml of 1⫻ PBS, air dried, and observed for the intensity of the blue color. MBIC and synergy of P128 with antibiotics. The minimum biofilm inhibitory concentration (MBIC) assay was optimized using the standard S. aureus strain ATCC 29213. An overnight culture of the strain was diluted 1:40 in LB broth. Two hundred microliters of diluted culture was aliquoted into each well of a microtiter plate. Microtiter plates were placed in a shaker-incubator set to 37°C and 100 rpm for 24 h, followed by 48 h of incubation under static conditions at 37°C. The contents of one set of four

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TABLE 2 Synergy of P128 in combination with SoC antibiotics on planktonic cells of S. aureus MIC (␮g/ml)

MIC (␮g/ml)

MIC (␮g/ml)

S. aureus straina

P128

Vancomycin

P128 ⫹ vancomycin

FICI

P128

Ciprofloxacin

P128 ⫹ ciprofloxacin

FICI

P128

Gentamicin

ATCC 29213

4

1

4 ⫹ 0.09

1.09

4

0.4

NDb

NDb

4

0.48

BK1

4

1

4 ⫹ 0.18

1.18

4

32

0.25 ⫹ 8

0.31

4

0.48

B9241

8

1

4 ⫹ 0.18

0.68

8

8

2 ⫹ 0.25

0.28

8

62

P128 ⫹ gentamicin 1.00 ⫹ 0.48 1.00 ⫹ 0.48 2 ⫹ 31.25

FICI 1.25 1.25 0.75

a

ATCC 29213 is susceptible to vancomycin, ciprofloxacin, and gentamicin; BK1 is susceptible to vancomycin and gentamicin and resistant to ciprofloxacin; and B9241 is susceptible to vancomycin and resistant to gentamicin and ciprofloxacin. b ND, not determined.

wells were aspirated and discarded. The wells were washed twice with 1⫻ PBS, and the presence of biofilm in the wells at the end of 72 h was determined by a metabolic dye reduction assay using MTT [3-(4,5-dimethyl-2-thiazolyl)-2,5-diphenyl-2H-tetrazolium bromide; Himedia]. In this assay, live cells reduce the dye, leading to color formation which can be read at 570 nm, and the intensity of the color can be correlated to the number of live cells. One hundred microliters of PBS was added to each well along with 10 ␮l of MTT solution. The plate was incubated for 2 h in the dark. After this, 110 ␮l of solvent solution (solubilizing agent) was added, and the plate was incubated for 15 min at ambient temperature with gentle agitation. The absorbance was read at 570 nm in a microplate reader. Another set of four wells was processed for harvesting of the biofilm and determination of the number of CFU present by plating on solid medium. For biofilm inhibitory studies, the wells were washed twice with 1⫻ PBS, challenged with various concentrations of P128 or other antibiotic drugs in LB, and incubated for 6 or 24 h at 37°C. LB supplemented with 50 ␮g/ml CaCl2 was used for daptomycin treatment wells. The contents of the wells were aspirated out and discarded. The biofilms adhered to the wells were quantified by MTT assay as described above. The MBIC was defined as the minimum concentration of P128 or other drug showing no color development. To test if P128 showed synergy with other drugs, combinations of P128 and antibiotics were tested by a previously described checkerboard method (25). In each experiment, in addition to the combination MBIC, the MBIC of each drug was also determined individually. The fractional MBICs were determined by the MTT dye method as described above. The FICI and synergy were also calculated in a similar manner. Inhibition of multispecies biofilm formation. Lubbock chronic wound pathogenic biofilm (LCWPB) medium (Bolton broth [Oxoid Ltd.] supplemented with 50% bovine plasma and 5% hemolyzed horse blood) was used for multispecies biofilm formation according to a procedure described earlier (23). Briefly, Pseudomonas aeruginosa PAO1, Enterococcus faecalis ATCC 29212, and S. aureus ATCC 700699 grown on TSB agar plates were inoculated into TSB and grown at 37°C in a shaker for 16 h. The cultures were individually diluted to 1 ⫻ 106 CFU/ml and mixed in equal volumes, and 10 ␮l was added to 3 ml of LCWPB medium containing a sterile pipette tip. For biofilm formation, either two (P. aeruginosa PAO1 and S. aureus ATCC 700699) or all three (P. aeruginosa PAO1, E. faecalis ATCC 29212, and S. aureus ATCC 700699) bacterial species were inoculated into LCWPB medium. In this model, the pipette tip acts as a surface for biofilm formation. To test the ability of P128 to prevent biofilm formation, P128 at 10, 50, or 250 ␮g/ml was added to the tubes, and the tubes were incubated at 37°C in a shaker for 24 h with shaking at 150 rpm. Upon completion of incubation, the tips were removed from the tubes and placed on petri plates for observation. In the absence of P128, a confluent and thick mass of biofilm could be seen. The mass of the biofilm was greater for culture tubes with 3 species than for those with 2 species. For enumeration of bacteria in biofilms, the biofilms formed on the tips were washed twice in PBS, transferred to clean test tubes, and again washed twice with PBS. The washed biofilm mass was

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then transferred to a 50-ml conical polypropylene tube, and the biofilm was macerated with sterile scissors. In situations where biofilm formation was not visible, the tips alone were processed as described above. The contents were vortexed thoroughly, diluted, and plated on Trypticase soy agar (TSA) plates. The plates were incubated for 24 h at 37°C followed by incubation at ambient temperature for 24 h to enhance pigment production.

RESULTS

Effects of combinations of P128 and antibiotics on planktonic cells. In order to find out whether P128 can act in a synergistic manner with commonly used antibiotics, we performed MICbased growth inhibition assays. MIC-based synergy was studied by the checkerboard method by determining the FICI by use of a previously reported method (25). The synergistic potential of P128 with vancomycin, gentamicin, and ciprofloxacin to inhibit planktonic S. aureus cells was tested on one sensitive S. aureus strain (ATCC 29213) and two resistant S. aureus strains, namely, BK1 (MRSA) and B9241 (gentamicin-resistant MRSA [GMRSA]). The MICs of P128 on these strains were found to range from 4 to 8 ␮g/ml, while the MIC of vancomycin was 1 ␮g/ml (Table 2). Similarly, the MIC of ciprofloxacin was 0.4 ␮g/ml on the sensitive strains and 8 to 32 ␮g/ml on fluoroquinolone-resistant strains. Gentamicin showed MICs of 0.4 and 62 ␮g/ml on sensitive and resistant strains, respectively. The combination of P128 and vancomycin or gentamicin showed an additive or indifferent effect, as the FICIs for the three strains were found to be between 0.68 and 1.25. In contrast to vancomycin and gentamicin, ciprofloxacin in combination with P128 showed a clear synergistic effect, with FICIs ranging from 0.28 to 0.31. The combination MIC of ciprofloxacin on the BK1 and B9241 strains dropped to 8 and 0.25 ␮g/ml, in contrast to the individual MICs of 32 ␮g/ml and 8 ␮g/ml, respectively (Table 2). In summary, when they were tested on planktonic S. aureus bacteria, vancomycin and gentamicin showed an additive or indifferent effect in combination with P128 in inhibiting both methicillin-susceptible S. aureus (MSSA) and MRSA, while ciprofloxacin showed synergy in combination with P128. Standardization of biofilm formation conditions. Culture conditions were optimized for reproducibly obtaining robust biofilms of S. aureus ATCC 29213 in microtiter plates. For this purpose, biofilms were generated in microtiter plates, and the surfaceadhered cultures remaining after washing off the planktonic cells were analyzed at the end of 48 and 72 h by an MTT dye assay. In order to understand the relationship between CFU, visible color formation, and OD570 in the MTT assay, increasing numbers of

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Antibiofilm Activity of P128

TABLE 3 Synergy of P128 and SoC drugs in inhibiting S. aureus biofilmsa MBIC of P128 (␮g/ml)b,c Strain

6h

S. aureus ATCC 29213 6.04 ⫾ 2.89

S. aureus BK1

S. aureus B9241

MBIC of antibiotic (␮g/ml)b,c 24 h 200

Antibiotic

Gentamicin Vancomycin Ciprofloxacin 28.95 ⫾17.53 400 Gentamicin Vancomycin Ciprofloxacin 105.15 ⫾ 34.28 ⬎1,000 Gentamicin Vancomycin Ciprofloxacin

MBIC of P128 and antibiotic (␮g/ml) in combination (24 h)b

MBIC of P128 and antibiotic (␮g/ml) in combination (6 h)b

6h

24 h

P128

Antibiotic

FICI P128

Antibiotic

FICI

⬎1,000 7.8 ⬎250 ⬎1,000 31.25 ⬎250 ⬎1,000 31.25 ⬎250

⬎1,000 218.75 ⫾ 62.5 6.5 ⫾ 2.2 ⬎1,000 250 ⬎250 ⬎1,000 125 250

0.78 2.07 ⫾ 0.8 0.78 12.5 20.83 ⫾ 7.2 12.5 16.6 ⫾ 7.2 16.6 ⫾ 7.2 25

3.9 3.9 0.9 0.45 1.9 0.48 26.03 ⫾ 9.0 15.6 41.6 ⫾ 18

0.13 0.84 0.13 0.43 0.77 0.43 0.18 0.65 0.40

3.1 39.06 ⫾ 15.6 0.48 109.35 ⫾ 31.2 7.8 27.33 ⫾ 7.8 109.35 ⫾ 31.2 39.06 ⫾ 15.6 13.65 ⫾ 3.9

0.06 0.23 0.13 0.14 0.53 0.16 0.30 0.33 0.07

12.5 10.93 ⫾ 3.15 12.5 12.5 200 21.87 ⫾ 6.2 200 25 21.87 ⫾ 6.2

a

The activities of P128 and antibiotics on preformed biofilms were determined individually and in combination. MBIC values presented here are averages of three values ⫾ standard deviations (SD). In cases where the same value was obtained thrice, no SD value is shown. c In cases where the MBIC values were higher than the highest value tested (for P128 or individual drugs, not those in combination), we considered the highest concentration for the sake of calculation. This gives an underestimation of synergy, and hence the actual synergy is even higher (i.e., lower FICI) than that shown here. b

CFU (105 to 108) were exposed to MTT dye, and visible color formation and the OD570 were noted. As shown in Table S1 in the supplemental material, color formation in this assay could be observed only with ⬎3 ⫻ 106 CFU, which corresponded to an OD570 of 0.16. A linear increase in OD from 0.16 to 1.8 was observed upon an increase in CFU from 106 to 108 (see Fig. S1). In S. aureus ATCC 29213 biofilms at the end of 48 h, an average OD570 of 0.08 was observed, and approximately 106 CFU could be recovered from the wells. On further incubation, the bacterial counts increased to 108 CFU (OD570 ⫽ 2.0) at the end of 72 h. Based on these results, all the cultures were incubated for up to 72 h to allow formation of a thick biofilm. The OD570 values obtained at the end of 72 h with the various Staphylococcus strains used in this study are shown in Table S2. The 72-h biofilms of various S. aureus strains contained roughly 108 CFU per well of a microtiter plate. P128 inhibits growth of S. aureus in established biofilms. The antibiofilm activity of P128 on preformed biofilms of S. aureus was measured by determining the minimum biofilm inhibition concentration (MBIC), defined as the minimum concentration showing a visible lack of color development in an MTT-based assay. The assay measures the reduction in viability and/or inhibition of growth in preformed biofilms. Since P128 kills Staphylococcus planktonic cultures rapidly (14), we expected it to show a rapid effect on biofilms as well. Hence, we determined the MBIC values for this protein on various staphylococci after exposure times ranging from 2 to 24 h. As shown in Table S3 in the supplemental material, P128 caused a rapid reduction in cellular viability in biofilms, as the MBIC values obtained in 2 h were comparable to its MIC values on planktonic cells (Table 2). We observed that P128 inhibited cellular growth in biofilms of both MRSA and MSSA strains, with equal efficiencies. There were small increases in MBIC values for up to 8 h; however, the MBIC values of P128 increased drastically after incubation periods beyond 8 h (see Table S3). The reasons for the increase in MBIC values upon prolonged incubation are not currently understood. Based on these results, we decided to investigate the synergy of P128 and antibiotics against various MSSA and MRSA strains after 6 and 24 h of exposure (see below). As shown in Table 3, gentamicin did not inhibit biofilm formation even at the highest concentration tested (1,000 ␮g/ml), while vancomycin and ciprofloxacin had MBIC values considerably higher than their MICs.

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Bactericidal activity and biofilm eradication activity of P128. In order to determine whether P128 could also act as a bactericidal agent in biofilms, we monitored the CFU reduction in S. aureus ATCC 29213 72-h preformed biofilms treated with various concentrations of P128 for 6 h. As shown in Fig. 1A, treatment of S. aureus ATCC 29213 biofilms with P128 showed dose-dependent killing of S. aureus cells. During the incubation period, there was very slow growth of bacteria, resulting in an approximately 1-log CFU increase in untreated cultures in 6 h. P128 at 7.8 ␮g/ml killed ⬎90% of the cells, while exposure of biofilms to P128 concentrations of ⬎31 ␮g/ml led to 99.9% killing of S. aureus cells. The bactericidal activity of P128 was further confirmed on biofilms formed on the surfaces of catheters (see below). In the crystal violet staining assay, daptomycin, vancomycin, and linezolid showed insignificant activity on 24-h preformed biofilms of MSSA and MRSA strains even after treatment at a high concentration (250 ␮g/ml) for 24 h (Fig. 1B). This observation is in agreement with the findings of earlier investigators (15). In contrast, P128 at 1⫻ MIC (8 ␮g/ml) was able to eliminate the biofilms of the four strains tested within 2 h of treatment (Fig. 1B; see Fig. S2 in the supplemental material). This confirmed that P128 can eradicate an established biofilm of a MRSA strain in a rapid manner. In contrast to the results seen for the MBIC assay, P128 at 8 ␮g/ml was found to be effective in the crystal violet staining-based assay even after treatment for up to 24 h, with the only exception being strain B9241, as P128 was effective for up to 4 h but biofilm growth could be seen at the 24-h time point for this strain (see Fig. S2C). It should be noted that biofilms in the MBIC assay were grown for 72 h, while in the case of microscopy-based or crystal violet staining assays the biofilms were grown for 18 to 24 h. The biofilm eradication activity of P128 in biofilms was further confirmed by observing the treated biofilms by scanning electron microscopy (SEM). Observations of 18-h-old biofilms by SEM showed that S. aureus BK1 formed thick biofilms on the surfaces of the microtiter plates (Fig. 1C). Gentamicin at 50 ␮g/ml (⬎100 times higher than the planktonic cell MIC) did not have any appreciable effect on the appearance of the biofilms, and both the matrix and the embedded bacterial cells were seen to be intact. In contrast, P128 at the lowest concentration tested (12.5 ␮g/ml) destroyed the biofilm structure and lysed the bacterial cells com-

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FIG 1 Antibiofilm activity of P128 on MSSA and MRSA biofilms. (A) Seventy-two-hour biofilms of S. aureus ATCC 29213 in microtiter plates were treated with the indicated concentrations of P128 for 6 h, and cell viability was determined by plating on TSB agar plates. (B) Crystal violet staining of S. aureus BK1 biofilms in microtiter plates treated with daptomycin (Dapto), vancomycin (vanco), linezolid (Lin), or P128 at the indicated concentrations for 2, 4, and 24 h. CC, untreated control culture. (C) Scanning electron micrographs of S. aureus BK1 24-h biofilms on the surfaces of microtiter plates treated with the indicated concentrations of P128 or gentamicin.

pletely, and no intact biofilm was visible in a number of fields analyzed (Fig. 1C). P128 eliminates preformed biofilms from the surfaces of catheters. In order to simulate the in vivo conditions for biofilm formation in device-associated infections, we allowed S. aureus to form biofilms on the surfaces of catheters. Because the conditions used for S. aureus biofilm formation in microtiter plates did not yield any biofilms on the catheter surface, 5% hemolyzed plasma, a blood component used to obtain luxuriant biofilms in vitro (23), was added to the culture medium. This modification led to the formation of robust biofilms by the MRSA MW2 strain as detected by safranin staining and by SEM (Fig. 2A and B). Treatment of biofilms with P128 at 1⫻ MIC (8 ␮g/ml) led to their eradication, as no biofilm was visible upon safranin staining (Fig. 2A). Similar observations were made by visualization of P128-treated biofilms by SEM, wherein it was observed that P128 used at 1⫻ MIC eradicated biofilms from the surfaces of catheters, whereas vancomycin at 90 ␮g/ml (90⫻ MIC) had a minimal effect on the structure of the biofilms (Fig. 2B). Biofilms treated with daptomycin at 10⫻ MIC (10 ␮g/ml) showed significant reductions in biofilm mass, though some intact patches of biofilm could be visualized (Fig. 2B). Thus, both safranin staining assays and SEM observations

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confirmed that P128 possesses potent antibiofilm activity on MRSA biofilms growing on catheters. The bactericidal effect of 8 ␮g/ml (1⫻ MIC) of P128 on S. aureus cells growing in biofilms on the surfaces of catheters led to a 3-log reduction in CFU counts (Fig. 2C). Under similar conditions, daptomycin at 10 ␮g/ml (10⫻ MIC) showed a 1-log CFU reduction, while vancomycin at 15 ␮g/ml (15⫻ MIC) showed no significant effect on S. aureus viability. P128 shows strong synergy with antibiotics in inhibiting S. aureus in biofilms. In order to determine whether P128 could synergize with known anti-Staphylococcus drugs to inhibit S. aureus in biofilms, we determined the MBIC values of gentamicin, vancomycin, and ciprofloxacin in combination with P128 on three S. aureus strains of different antibiotic sensitivities, namely, ATCC 29213 (sensitive strain), BK1 (MRSA; resistant to ciprofloxacin), and B9241 (MRSA; resistant to gentamicin and ciprofloxacin). The assays were performed in a 96-well checkerboard format, using various concentrations of P128 and one of the antibiotics. As seen in Table 3, the MBIC values of P128 and the antibiotics in combination were much reduced compared to the individual MBIC values for all three strains of S. aureus. The maximum re-

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FIG 2 Antibiofilm activity of P128 on preformed MRSA biofilms on catheters. (A) Safranin-stained images of S. aureus MW2 biofilms on the surfaces of catheters treated with P128. (B) Scanning electron micrographs of S. aureus MW2 biofilms on the surfaces of catheters treated with P128, vancomycin, or daptomycin as described in Materials and Methods. (C) Viable cells remaining on the catheter surface after treatment with P128 or antibiotics. The experiment was repeated twice with similar results, and CFU data from one of the experiments are presented here.

duction in MBIC for a combination was observed in the case of gentamicin (⬎250-fold), especially with the two gentamicin-sensitive strains (ATCC 29213 and BK1). In the majority of cases (except the combination of P128 and vancomycin in the 6-h assay), the FICIs of P128 combinations with the antibiotics ranged from 0.06 to 0.53, suggesting a strong synergistic mechanism of inhibition. The synergistic effect of P128 could be seen even for drug combinations on strains which were resistant to the particular drugs: for example, P128 and ciprofloxacin showed synergy on the ciprofloxacin-resistant BK1 strain (FICIs of 0.43 and 0.16 in

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6-h and 24-h assays, respectively). Similarly, on S. aureus B9241, which is resistant to ciprofloxacin and gentamicin, a combination of P128 with gentamicin or ciprofloxacin showed synergistic inhibition, with FICI values ranging from 0.07 to 0.39. Isobolograms showing sub-MBIC values of P128 plotted against the sub-MBIC values of one of the three drugs in combination on B9241 are depicted in Fig. 3A. The shape of the curves clearly shows that P128 acts synergistically with these drugs. Isobolograms showing the synergy of P128 with the drugs used on ATCC 29213 and BK1 are depicted in Fig. S3 in the supplemental material. With the

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FIG 3 Synergy of P128 with antibiotics on S. aureus biofilms, represented in the form of isobolograms generated by plotting fractional MBIC values of P128 against fractional MBIC values of drugs used in combination. The individual MBIC values of P128 and the drugs are joined by solid lines, while the MBIC values obtained for P128-drug combinations are joined by dotted lines. (A) Isobolograms for activity of P128 in combination with gentamicin, vancomycin, or ciprofloxacin on S. aureus B9241. (B) Isobolograms for activity of P128 in combination with vancomycin, linezolid, or daptomycin on S. aureus MW2.

exception of isobolograms depicting P128 and vancomycin combinations on ATCC 29213 and BK1 showing an additive effect, the other graphs point toward a synergistic inhibition of biofilms. Interestingly, despite an increase in the MBIC value of P128 over 24 h, the combinations of P128 with all three drugs were found to be highly synergistic in inhibiting growth of S. aureus in biofilms even after 24 h (Table 3, last column). The shapes of the curves in the isobolograms shown in Fig. S4 demonstrate that vancomycin, gentamicin, and ciprofloxacin showed high levels of synergy with P128 on all three strains of S. aureus in the 24-h assay. Furthermore, in order to find out if P128 would show synergy with SoC drugs possessing other mechanisms of action, we tested the synergy of P128 with linezolid and daptomycin on S. aureus MW2. As seen in Table S4, vancomycin and linezolid were poorly effective on biofilms, whereas daptomycin showed more activity. However, with all three drugs there was a huge reduction in MBIC (8- to ⬎128-fold) in the presence of P128, resulting in FICI values ranging from 0.24 to 0.36. The synergistic effect of P128 on the three drugs is clearly visible in isobolograms plotted using sub-MBIC values of various drugs against sub-MBIC values of P128 (Fig. 3B). Thus, P128 showed synergy with different classes of antibiotics against various strains of S. aureus by virtue of its ability to reduce the MBIC values severalfold. Prevention of biofilm formation by P128 in a mixed-culture model simulating chronic wounds in vitro. Chronic wounds, such as venous leg ulcers, are often infected with multiple species of Gram-positive and Gram-negative bacteria residing in a biofilm (26, 27). An in vitro model which uses plasma and laked horse

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blood for the growth of S. aureus, P. aeruginosa, and E. faecalis, either singly or in mixed cultures allowing biofilm formation using a solid support, has been described previously (23). This model is supposed to mimic the wound environment under in vitro conditions. The ability of P128 to prevent biofilm formation in the mixed-culture biofilm model was tested by the procedure described in Materials and Methods. The combination of either S. aureus and P. aeruginosa or S. aureus, P. aeruginosa, and E. faecalis cultures led to the formation of a thick and leathery biofilm (Fig. 4) carrying approximately equal numbers (107 to 108 CFU/ml) of all the organisms (Table 4). P128 at a concentration as low as 1⫻ MIC (10 ␮g/ml) prevented the formation of biofilms in this model. The lack of biofilm formation was reflected in very low bacterial counts of P. aeruginosa, E. faecalis, and S. aureus obtained after processing the pipette tips used for growth of biofilms. At 50 and 250 ␮g/ml of P128, there was a further reduction in S. aureus counts, whereas the counts of P. aeruginosa and E. faecalis remained at 104 to 105 CFU/ml. Since P128 does not inhibit the growth of E. faecalis and P. aeruginosa (14), these results suggest that inhibition of S. aureus growth alone in this model is sufficient to prevent biofilm formation even by P. aeruginosa and E. faecalis. This is consistent with the results of earlier studies involving inhibition of S. aureus and P. aeruginosa by various biofilm inhibitors in the same model of in vitro biofilm formation (28). DISCUSSION

Biofilms are surface-adhered phenotypically heterogenous communities of microorganisms (29) found both in vitro and in vivo in

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FIG 4 Prevention of multispecies biofilm formation by P128 by virtue of its ability to kill S. aureus. The first three tubes contain pipette tips transferred from cultures treated with increasing concentrations of P128 (10, 50, and 250 ␮g/ml), while the last tube contains a tip from an untreated culture. Biofilm formation could be seen only in the last tube.

infected tissues. S. aureus is known to form biofilms under a variety of clinical conditions, such as osteomyelitis, indwelling medical device-associated infections, endocarditis, chronic wound infection, chronic rhinosinusitis, and ocular infections (30). In a chronic wound environment, bacterial contamination leads to colonization of bacteria followed by formation of bacterial biofilms on the surfaces of dead cells in the wounds (29, 31, 32). The established biofilms are highly recalcitrant to antibiotics and can evade the immune response (4, 7). Biofilms can act as reservoirs of infection and are difficult to eradicate, leading to both treatment failure and recurrent episodes of the disease. It has been proven that one of the major reasons for treatment failure in the case of chronic wounds is phenotypic resistance of bacteria present in a biofilm to antimicrobials and to the immune system (31). There is physiological heterogeneity among cells in biofilms (33), and many characteristics of biofilms contribute to their resistance to antibacterials and immunity, including a protective barrier in the form of the biofilm matrix, expression of specific proteins, low metabolic activity, and induction of a persister state in which bacterial resistance to antimicrobial treatment increases (7, 34). Thus, an ideal antibiofilm agent should be able to destroy and penetrate the biofilm matrix and should be bactericidal to slowly replicating and persister cell populations within the biofilm. Bacteriophages and phage-derived proteins are emerging as viable alternatives for treatment of drug-resistant infections caused by biofilm-forming bacteria (11, 17). In this study, we examined the antibacterial properties of P128 on S. aureus biofilms. P128 at low concentrations showed strong inhibition of S. aureus cells growing in biofilms. P128 was equally efficient at eliminating MSSA and MRSA biofilms from the surfaces of both microtiter plates and catheters. These results are consistent with the previously reported inhibitory activity of P128 seen on MSSA and MRSA strains in imaging and alamarBlue-based viability assays

TABLE 4 Prevention of multispecies biofilm formation by P128 by inhibition of S. aureus growtha Biofilm members

P128 concn (␮g/ml)

Isolate

No. of CFU/ml

Biofilm formation

P. aeruginosa PAO1, S. aureus ATCC 700699

0 (cell control)

P. aeruginosa S. aureus P. aeruginosa S. aureus P. aeruginosa S. aureus P. aeruginosa S. aureus

2.4 ⫻ 108 2.1 ⫻ 107 2 ⫻ 106 2.2 ⫻ 105 1.5 ⫻ 105 1.8 ⫻ 105 8 ⫻ 105 2 ⫻ 103



P. aeruginosa

7 ⫻ 108



S. aureus E. faecalis P. aeruginosa S. aureus E. faecalis P. aeruginosa S. aureus E. faecalis P. aeruginosa S. aureus E. faecalis

1 ⫻ 108 3 ⫻ 107 2.2 ⫻ 105 2 ⫻ 104 9 ⫻ 105 7 ⫻ 104 2 ⫻ 103 1.8 ⫻ 104 1.7 ⫻ 105 ⬍10 1.6 ⫻ 104

10 50 250

P. aeruginosa PAO1, S. aureus ATCC 700699, E. faecalis ATCC 29212

0 (cell control)

10

50

250

⫺ ⫺ ⫺

⫺ ⫺ ⫺

a The cultures were treated with the indicated concentrations of P128 and incubated for 24 h. Subsequently, pipette tips with or without biofilms were processed as described in Materials and Methods, and the CFU counts were recorded. ⫹ and ⫺, presence and absence, respectively, of a visible biofilm on the surface of the pipette tip. The experiment was repeated three times with similar results, and CFU counts from one of the experiments are shown here.

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(20). The higher MBIC values seen with 24-h MBIC assays in the current study can possibly be attributed to the thicker biofilms formed over a period of 72 h in this assay, in contrast to 18- to 24-h-old biofilms in other assays. It has been shown (35) that the constitutions of biofilms formed by MSSA and MRSA are different; P128’s ability to act equally well on both is thus an important finding. The ability to eradicate biofilms from the surfaces of catheters suggests that P128 has the potential to control biofilms in device-associated infections caused by S. aureus. Similar to its rapid activity on planktonic cells, P128 could kill or inhibit the growth of S. aureus in biofilms in a rapid manner, as demonstrated by the low MBIC values seen in a 2-h assay using sensitive and resistant strains of S. aureus. The MBIC values were only 1- to 4-fold higher than the planktonic MICs, demonstrating that P128 has potent activity on S. aureus biofilms. The ability of P128 to destroy the biofilm structure of S. aureus as evidenced by SEM suggests that the biofilm matrix might not be a major barrier for the entry of P128. In addition, P128 can kill cells which are metabolically inactive (e.g., in buffers), and this property may play a crucial role in killing poorly metabolizing cells trapped inside biofilms. The antibiofilm activity of P128 observed with various media, surfaces, and strains of S. aureus demonstrates that P128 can eliminate biofilms formed under a variety of physiological conditions. Because eradication of biofilms by single antimicrobial agents is extremely difficult, the discovery of agents showing synergy in inhibiting bacteria in biofilms should lead to better clinical treatment outcomes for S. aureus infections involving biofilms. In addition, combination therapy of serious infections can prevent the emergence of drug resistance and can also help in reducing the duration of therapy. Earlier studies have shown synergistic activity of lysostaphin or phage-derived lysins with antibiotics on planktonic cells of S. epidermidis and S. aureus (36, 37). However, to the best of our knowledge, combinations of lysins and antibiotics have not been tested on S. aureus biofilms. Although the combinations of P128 and antibiotics showed only an additive effect or a modest synergy on planktonic cells of S. aureus, the same combinations showed dramatic synergistic effects on biofilms. The huge lowering of the MBICs of antibiotics of various classes, including newer ones, such as linezolid and daptomycin, resulting in low FICI values for combinations of P128 with antibiotics, suggests that P128 can greatly potentiate the effects of antibiotics on biofilms. Because P128 kills bacteria by disrupting the peptidoglycan of the bacterial cell wall, the strong synergy seen with antibiotics may result from an increase in permeability of the cells to the antibiotics. The ability of P128 to prevent biofilm formation in a mixedculture biofilm model by inhibiting growth of S. aureus suggests that S. aureus plays a major role in biofilm formation in this setting. Our findings are also supported by the recently reported excellent clinical efficacy of Staphylococcus phage Sb-1 in diabetic foot ulcer patients (38) who had not responded to antibiotic treatment earlier and had open wounds, which are often polymicrobial in nature (27). Based on these results, it is reasonable to suggest that P128 might help in controlling biofilms in chronic wounds which are infected with multiple bacterial species (26, 27). Though we have not tested all the strains used in this study in the various assays described, we did not observe any difference in the antibiofilm activity of P128 on various strains or in different assays mea-

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suring viability or biomass removal. The effects of immunogenicity of P128 on efficacy and safety need to be evaluated. However, the recent demonstration of a lack of any influence of antibodies on efficacy and safety in animals immunized with a lysin (39) suggests that immunogenicity may not pose a big challenge in developing P128 as a therapeutic. Strong synergistic killing of biofilm-embedded S. aureus, including MRSA, by P128 in combination with SoC antibiotics makes P128 a potential candidate for clinical development for treatment of serious S. aureus infections, such as chronic wounds, bacteremia, infective endocarditis, and device-associated infections. ACKNOWLEDGMENTS We thank J. Ramachandran, M. Jayasheela, and T. S. Balganesh for encouragement and for providing suggestions during the study. We thank Amy Percy, Vivek Paul, Bharathi Sriram, and Anand Kumar for reviewing the manuscript. We thank J. Raghupatil and B. Rajagopala for assisting with some of the biofilm experiments and the GangaGen DSP team for providing purified P128 preparations. We appreciate the support provided by Shashimohan Keelara in arranging catheters for biofilm studies. We are thankful to Randall Walcott, Wound Care Center, Lubbock, TX, for sharing the multispecies biofilm protocol developed in his lab. We are grateful to Aloysius Daniel, Central Manufacturing Technology Institute, Bangalore, India, for providing the technical expertise for scanning electron microscopy of the biofilm samples. We appreciate the critical inputs provided by anonymous reviewers for improving the manuscript. S.N. and N.P. planned and executed the experiments and did the troubleshooting. S.D. and A.V. designed the experiments and analyzed the data. U.S. analyzed the data and wrote the manuscript. All the authors reviewed the manuscript.

FUNDING INFORMATION This research received no specific grant from any funding agency in the public, commercial, or not-for-profit sectors.

REFERENCES 1. Yayan J, Ghebremedhin B, Rasche K. 2015. No outbreak of vancomycin and linezolid resistance in staphylococcal pneumonia over a 10-year period. PLoS One 10:e0138895. http://dx.doi.org/10.1371/journal.pone .0138895. 2. Kelley PG, Gao W, Ward PB, Howden BP. 2011. Daptomycin nonsusceptibility in vancomycin-intermediate Staphylococcus aureus (VISA) and heterogeneous-VISA (hVISA): implications for therapy after vancomycin treatment failure. J Antimicrob Chemother 66:1057–1060. http: //dx.doi.org/10.1093/jac/dkr066. 3. Gu B, Kelesidis T, Tsiodras S, Hindler J, Humphries RM. 2013. The emerging problem of linezolid-resistant Staphylococcus. J Antimicrob Chemother 68:4 –11. http://dx.doi.org/10.1093/jac/dks354. 4. Otto M. 2008. Staphylococcal biofilms. Curr Top Microbiol Immunol 322:207–228. 5. Thwaites GE, Edgeworth JD, Gkrania-Klotsas E, Kirby A, Tilley R, Török ME, Walker S, Wertheim HF, Wilson P, Llewelyn MJ, UK Clinical Infection Research Group. 2011. Clinical management of Staphylococcus aureus bacteraemia. Lancet Infect Dis 11:208 –222. http://dx.doi .org/10.1016/S1473-3099(10)70285-1. 6. Keren I, Kaldalu N, Spoering A, Wang Y, Lewis K. 2004. Persister cells and tolerance to antimicrobials. FEMS Microbiol Lett 230:13–18. http: //dx.doi.org/10.1016/S0378-1097(03)00856-5. 7. Lewis K. 2008. Multidrug tolerance of biofilms and persister cells. Curr Top Microbiol Immunol 322:107–131. 8. Nguyen D, Joshi-Datar A, Lepine F, Bauerle E, Olakanmi O, Beer K, McKay G, Siehnel R, Schafhauser J, Wang Y, Britigan BE, Singh PK. 2011. Active starvation responses mediate antibiotic tolerance in biofilms and nutrient-limited bacteria. Science 334:982–986. http://dx.doi.org/10 .1126/science.1211037. 9. Rogers GB, Carroll MP, Bruce KD. 2012. Enhancing the utility of exist-

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December 2016 Volume 60 Number 12

Antibiofilm Activity of P128

10.

11.

12.

13.

14.

15.

16.

17.

18.

19.

20.

21.

22.

23.

ing antibiotics by targeting bacterial behaviour? Br J Pharmacol 165:845– 857. http://dx.doi.org/10.1111/j.1476-5381.2011.01643.x. Chung PY, Toh YS. 2014. Anti-biofilm agents: recent breakthrough against multi-drug resistant Staphylococcus aureus. Pathog Dis 70:231– 239. http://dx.doi.org/10.1111/2049-632X.12141. Parasion S, Kwiatek M, Gryko R, Mizak L, Malm A. 2014. Bacteriophages as an alternative strategy for fighting biofilm development. Pol J Microbiol 63:137–145. Seth AK, Geringer MR, Nguyen KT, Agnew SP, Dumanian Z, Galiano RD, Leung KP, Mustoe TA, Hong SJ. 2013. Bacteriophage therapy for Staphylococcus aureus biofilm-infected wounds: a new approach to chronic wound care. Plast Reconstr Surg 131:225–234. http://dx.doi.org /10.1097/PRS.0b013e31827e47cd. Loeffler JM, Nelson D, Fischetti VA. 2001. Rapid killing of Streptococcus pneumoniae with a bacteriophage cell wall hydrolase. Science 294:2170 – 2172. http://dx.doi.org/10.1126/science.1066869. Paul VD, Rajagopalan SS, Sundarrajan S, George SE, Asrani JY, Pillai R, Chikkamadaiah R, Durgaiah M, Sriram B, Padmanabhan S. 2011. A novel bacteriophage tail-associated muralytic enzyme (TAME) from phage K and its development into a potent antistaphylococcal protein. BMC Microbiol 11:226. http://dx.doi.org/10.1186/1471-2180-11-226. Schuch R, Lee HM, Schneider BC, Sauve KL, Law C, Khan BK, Rotolo JA, Horiuchi Y, Couto DE, Raz A, Fischetti VA, Huang DB, Nowinski RC, Wittekind M. 2014. Combination therapy with lysin CF-301 and antibiotic is superior to antibiotic alone for treating methicillin-resistant Staphylococcus aureus-induced murine bacteremia. J Infect Dis 209:1469 – 1478. http://dx.doi.org/10.1093/infdis/jit637. Wu JA, Kusuma C, Mond JJ, Kokai-Kun JF. 2003. Lysostaphin as a potential therapeutic agent for staphylococcal biofilm eradication. Antimicrob Agents Chemother 47:3407–3414. http://dx.doi.org/10.1128/AAC .47.11.3407-3414.2003. Pastagia M, Schuch R, Fischetti VA, Huang DB. 2013. Lysins: the arrival of pathogen-directed anti-infectives. J Med Microbiol 62:1506 –1516. http://dx.doi.org/10.1099/jmm.0.061028-0. Roach DR, Donovan DM. 2015. Antimicrobial bacteriophage-derived proteins and therapeutic applications. Bacteriophage 5:e1062590. http: //dx.doi.org/10.1080/21597081.2015.1062590. Vipra AA, Desai SN, Roy P, Patil R, Raj JM, Narasimhaswamy N, Paul VD, Chikkamadaiah R, Sriram B. 2012. Antistaphylococcal activity of bacteriophage derived chimeric protein P128. BMC Microbiol 12:41. http: //dx.doi.org/10.1186/1471-2180-12-41. Drilling AJ, Cooksley C, Chan C, Wormald PJ, Vreugde S. 2016. Fighting sinus-derived Staphylococcus aureus biofilms in vitro with a bacteriophage-derived muralytic enzyme. Int Forum Allergy Rhinol 6:349 – 355. http://dx.doi.org/10.1002/alr.21680. Sundarrajan S, Raghupatil J, Vipra A, Narasimhaswamy N, Saravanan S, Appaiah C, Poonacha N, Desai S, Nair S, Bhatt RN, Roy P, Chikkamadaiah R, Durgaiah M, Sriram B, Padmanabhan S, Sharma U. 2014. Bacteriophage-derived CHAP domain protein, P128, kills Staphylococcus cells by cleaving interpeptide cross-bridge of peptidoglycan. Microbiology 160:2157–2169. http://dx.doi.org/10.1099/mic.0.079111-0. George SE, Chikkamadaiah R, Durgaiah M, Joshi AA, Thankappan UP, Madhusudhana SN, Sriram B. 2012. Biochemical characterization and evaluation of cytotoxicity of antistaphylococcal chimeric protein P128. BMC Res Notes 5:280. http://dx.doi.org/10.1186/1756-0500-5-280. Sun Y, Dowd SE, Smith E, Rhoads DD, Wolcott RD. 2008. In vitro multispecies Lubbock chronic wound biofilm model. Wound Repair Regen 16:805– 813. http://dx.doi.org/10.1111/j.1524-475X.2008.00434.x.

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24. Clinical and Laboratory Standards Institute. 2012. Methods for dilution antimicrobial susceptibility tests for bacteria that grow aerobically; approved standard. CLSI document M07-A9.9. CLSI, Wayne, PA. 25. Lu X, Yang X, Li X, Lu Y, Ren Z, Zhao L, Hu X, Jiang J, You X. 2013. In vitro activity of sodium new houttuyfonate alone and in combination with oxacillin or netilmicin against methicillin-resistant Staphylococcus aureus. PLoS One 8:e68053. http://dx.doi.org/10.1371/journal .pone.0068053. 26. Burmølle M, Thomsen TR, Fazli M, Dige I, Christensen L, Homøe P, Tvede M, Nyvad B, Tolker-Nielsen T, Givskov M, Moser C, Kirketerp-Møller K, Johansen HK, Høiby N, Jensen PØ, Sørensen SJ, Bjarnsholt T. 2010. Biofilms in chronic infections—a matter of opportunity—monospecies biofilms in multispecies infections. FEMS Immunol Med Microbiol 59:324 –336. http://dx.doi.org/10.1111/j .1574-695X.2010.00714.x. 27. Wolcott R, Costerton JW, Raoult D, Cutler SJ. 2013. The polymicrobial nature of biofilm infection. Clin Microbiol Infect 19:107–112. http://dx .doi.org/10.1111/j.1469-0691.2012.04001.x. 28. Dowd SE, Sun Y, Smith E, Kennedy JP, Jones CE, Wolcott R. 2009. Effects of biofilm treatments on the multi-species Lubbock chronic wound biofilm model. J Wound Care 18:508, 510 –512. 29. Costerton JW, Stewart PS, Greenberg EP. 1999. Bacterial biofilms: a common cause of persistent infections. Science 284:1318 –1322. http://dx .doi.org/10.1126/science.284.5418.1318. 30. Archer NK, Mazaitis MJ, Costerton JW, Leid JG, Powers ME, Shirtliff ME. 2011. Staphylococcus aureus biofilms: properties, regulation, and roles in human disease. Virulence 2:445– 459. http://dx.doi.org/10.4161 /viru.2.5.17724. 31. Donlan RM, Costerton JW. 2002. Biofilms: survival mechanisms of clinically relevant microorganisms. Clin Microbiol Rev 15:167–193. http://dx .doi.org/10.1128/CMR.15.2.167-193.2002. 32. Parsek MR, Singh PK. 2003. Bacterial biofilms: an emerging link to disease pathogenesis. Annu Rev Microbiol 57:677–701. http://dx.doi.org /10.1146/annurev.micro.57.030502.090720. 33. Stewart PS. 2015. Antimicrobial tolerance in biofilms. Microbiol Spectr 3:3. http://dx.doi.org/10.1128/microbiolspec.MB-0010-2014. 34. Fux CA, Costerton JW, Stewart PS, Stoodley P. 2005. Survival strategies of infectious biofilms. Trends Microbiol 13:34 – 40. http://dx.doi.org/10 .1016/j.tim.2004.11.010. 35. McCarthy H, Rudkin JK, Black NS, Gallagher L, O’Neill E, O’Gara JP. 2015. Methicillin resistance and the biofilm phenotype in Staphylococcus aureus. Front Cell Infect Microbiol 5:1. http://dx.doi.org/10.3389/fcimb .2015.00001. 36. Kiri N, Archer G, Climo MW. 2002. Combinations of lysostaphin with beta-lactams are synergistic against oxacillin-resistant Staphylococcus epidermidis. Antimicrob Agents Chemother 46:2017–2020. http://dx.doi.org /10.1128/AAC.46.6.2017-2020.2002. 37. Daniel A, Euler C, Collin M, Chahales P, Gorelick KJ, Fischetti VA. 2010. Synergism between a novel chimeric lysin and oxacillin protects against infection by methicillin-resistant Staphylococcus aureus. Antimicrob Agents Chemother 54:1603–1612. http://dx.doi.org/10.1128/AAC .01625-09. 38. Fish R, Kutter E, Wheat G, Blasdel B, Kutateladze M, Kuhl S. 2016. Bacteriophage treatment of intransigent diabetic toe ulcers: a case series. J Wound Care 7:S27–S33. http://dx.doi.org/10.12968/jowc.2016.25.7.S27. 39. Zhang L, Li D, Li X, Hu L, Cheng M, Xia F, Gong P, Wang B, Ge J, Zhang H, Cai R, Wang Y, Sun C, Feng X, Lei L, Han W, Gu J. 2016. LysGH15 kills Staphylococcus aureus without being affected by the humoral immune response or inducing inflammation. Sci Rep 6:29344. http: //dx.doi.org/10.1038/srep29344.

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Antibiofilm Activity and Synergistic Inhibition of Staphylococcus aureus Biofilms by Bactericidal Protein P128 in Combination with Antibiotics.

P128 is an antistaphylococcal protein, comprising a cell wall-degrading enzymatic region and a Staphylococcus-specific binding region, which possesses...
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