Next Article in Journal
Ability of Essential Oil Vapours to Reduce Numbers of Culturable Aerosolised Coronavirus, Bacteria and Fungi
Previous Article in Journal
Lagoon, Anaerobic Digestion, and Composting of Animal Manure Treatments Impact on Tetracycline Resistance Genes
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Antibiotics
Volume 11
Issue 3
10.3390/antibiotics11030392
antibiotics-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Evaluation of Bacteriophage-Antibiotic Combination Therapy for Biofilm-Embedded MDR Enterococcus faecium

by
Katherine Lev
1,
Ashlan J. Kunz Coyne
1,
Razieh Kebriaei
1,
Taylor Morrisette
2,
Kyle Stamper
1,
Dana J. Holger
1,
Gregory S. Canfield
3,4,
Breck A. Duerkop
3,
Cesar A. Arias
5,6 and
Michael J. Rybak
1,7,*
1
Anti-Infective Research Laboratory, College of Pharmacy and Health Sciences, Wayne State University, Detroit, MI 48201, USA
2
Department of Pharmacy and Clinical Services, Medical University of South Carolina College of Pharmacy, Charleston, SC 29208, USA
3
Department of Immunology and Microbiology, University of Colorado School of Medicine, Aurora, CO 80045, USA
4
Department of Infectious Diseases, University of Colorado School of Medicine, Aurora, CO 80045, USA
5
Division of Infectious Diseases, Houston Methodist Hospital, Houston, TX 77030, USA
6
Center for Infectious Diseases Research, Houston Methodist Research Institute, Houston, TX 77030, USA
7
School of Medicine, Wayne State University, Detroit, MI 48201, USA
*
Author to whom correspondence should be addressed.
Antibiotics 2022, 11(3), 392; https://doi.org/10.3390/antibiotics11030392
Submission received: 24 January 2022 / Revised: 22 February 2022 / Accepted: 5 March 2022 / Published: 15 March 2022
Browse Figures
Versions Notes

Abstract

:
Multidrug-resistant (MDR) Enterococcus faecium is a challenging pathogen known to cause biofilm-mediated infections with limited effective therapeutic options. Lytic bacteriophages target, infect, and lyse specific bacterial cells and have anti-biofilm activity, making them a possible treatment option. Here, we examine two biofilm-producing clinical E. faecium strains, daptomycin (DAP)-resistant R497 and DAP-susceptible dose-dependent (SDD) HOU503, with initial susceptibility to E. faecium bacteriophage 113 (ATCC 19950-B1). An initial synergy screening was performed with modified checkerboard MIC assays developed by our laboratory to efficiently screen for antibiotic and phage synergy, including at very low phage multiplicity of infection (MOI). The data were compared by one-way ANOVA and Tukey (HSD) tests. In 24 h time kill analyses (TKA), combinations with phage-DAP-ampicillin (AMP), phage-DAP-ceftaroline (CPT), and phage-DAP-ertapenem (ERT) were synergistic and bactericidal compared to any single agent (ANOVA range of mean differences 3.34 to 3.84 log10 CFU/mL; p < 0.001). Furthermore, phage-DAP-AMP and phage-DAP-CPT prevented the emergence of DAP and phage resistance. With HOU503, the combination of phage-DAP-AMP showed the best killing effect, followed closely by phage-DAP-CPT; both showed bactericidal and synergistic effects compared to any single agent (ANOVA range of mean differences 3.99 to 4.08 log10 CFU/mL; p < 0.001).

1. Introduction

Vancomycin-resistant enterococci (VRE) pose major therapeutic challenges due to their high prevalence in nosocomial infections, coupled with their propensity to develop resistance to standard-of-care (SOC) antibiotics [1]. Infections caused by VRE are endemic to hospitals, carry a high degree of morbidity and mortality, and prolong hospital length of stay, making them a rising public health threat [2,3,4,5]. Of note, E. faecium infections exhibit higher rates of antibiotic resistance, have increased mortality rates, and result in significantly higher hospital costs compared to E. faecalis [3,6]. Clearly, there is an urgent need for innovative treatment options for serious VRE infections [3,6,7].
A biofilm is a multilayer, three-dimensional aggregate of microbial cells, extracellular DNA, and proteins, bound by a polysaccharide matrix, which provides biofilm stability and serves as a bacterial shield against antimicrobials and immune modulators [8,9,10,11,12]. This bacterial structure complicates therapeutic decisions, since the selected treatment should be effective within the bacterial biofilm. E. faecium strains have a predisposition to colonize indwelling medical devices and form biofilms, which contribute to their ability to withstand antibiotics and evade the host immune system, allowing their persistence during infection and, worse, treatment failure [12,13,14].
Daptomycin (DAP) is a concentration-dependent cyclic lipopeptide with bactericidal activity against VRE in vitro by disrupting cytoplasmic membranes and peptidoglycan synthesis [15,16,17]. Furthermore, DAP can rapidly penetrate inside biofilms, as previously reported with fluorescent visualization [18]. However, the recent emergence of DAP-resistant VRE isolates due to substitutions in the liaFSR regulatory system points to an urgent need for viable options given the limited therapeutic alternatives [19,20,21,22,23,24,25].
Lytic bacteriophages (phages) are viruses that may serve as a novel therapy against biofilm-mediated infections given their ability to target specific host organisms and replicate within bacteria leading to an abundance of phage at the site of infection [26,27,28,29]. Furthermore, some phages have demonstrated success against biofilms formed by other bacterial pathogens implicated in biofilm-mediated infections [9,30,31,32,33]. The adjunctive use of phage and antibiotic therapies may be more effective than either one alone due to enhanced bactericidal activity and the potential to decrease/eliminate/delay the emergence of antibiotic and/or phage resistance or to re-sensitize resistant strains [34,35,36]. The additional benefit of phage-antibiotic combinations compared to the use of either agent alone has been demonstrated previously with promising results, whereas the use of phage monotherapy resulted in the emergence of phage resistance, which required either adjusting the concentration of the administered phage or the use of phage and antibiotic combinations [30,37,38,39,40,41]. E. faecium-specific phage 113 has demonstrated activity against multiple antibiotic-resistant isolates in the planktonic state [39]. This observation led us to investigate the hypothesis that this phage, in combination with antibiotics, has promising activity against E. faecium biofilms. The work described here determines: (i) the activity of bacteriophage 113 in combination with DAP and various β-lactams (e.g., ampicillin (AMP), ceftaroline (CPT), or ertapenem (ERT)) against biofilm-producing E. faecium isolates; and (ii) the impact of phage presence on both antibiotic and phage resistance in the biofilm state.

2. Results

2.1. Bacterial Isolates

E. faecium clinical isolates, R497 (DAP-resistant, MBIC = 16 µg/mL) and HOU503 (vancomycin-resistant; DAP-susceptible dose dependent (SDD), MBIC = 2 µg/mL), were evaluated in this study due to their initial susceptibility to phage 113 and relatively high biofilm production compared to the other isolates determined by the modified small drop agar method and crystal violet microtiter plate biofilm quantification assay, respectively (Table 1) [42,43,44].

2.2. Checkerboard Analyses

DAP in combination with AMP against DAP-resistant R497 in biofilm was additive, with a fractional inhibitory concentration (FIC) index of 1 (Figure 1A), while DAP plus AMP in the presence of subinhibitory phage (MOI of 10−8) was synergistic, with an FIC index of 0.5 (Figure 1B). Notably, DAP or AMP in the presence of low phage (MOI of 10−8) showed a 64-fold reduction in comparison to DAP or AMP without phage (Figure 1C,D). Furthermore, the inhibition of R497 growth was greater with the combination of phage plus AMP compared to phage plus DAP. For DAP-SDD HOU503, both phage-DAP-AMP and DAP-AMP combinations were additive, but not synergistic, despite increased killing at lower antibiotic concentrations with the addition of phage (Figure 2A,B; MOI of 10−8).

2.3. Time-Kill Analyses

In 24 h TKA against DAP-resistant R497, all the combinations of phage-DAP-AMP, phage-DAP-CPT, and phage-DAP-ERT were bactericidal, as compared to any single agent, and exhibited detection-level killing by 24 h (Figure 3A–C). Of note, combination therapy with phage-AMP also showed killing to detection limit (2 log10 CFU/mL) at 24 h (Figure 3A). In DAP-SDD HOU503, the combination of phage-DAP-AMP showed the best killing effect with a 3.61 log10 CFU/mL reduction from the initial inoculum (Figure 3D), followed closely by phage-DAP-CPT, with a 3.10 log10 CFU/mL reduction (Figure 3E), and both were again bactericidal and synergistic compared to the single-agent antibiotic or phage.

2.4. Bacteriophage and Antibiotic Resistance Testing

Bacteriophage treatment-emergent resistance was observed for R497 in treatments with phage alone and phage-ERT (Table 2) In HOU503, we observed resistance in all combinations of phage plus single antibiotic, as well as in phage-DAP-ERT, which is consistent with the results of the TKA, which showed minimal killing activity in this combination. However, combinations of phage-DAP-AMP and phage-DAP-CPT prevented the emergence of phage resistance in HOU503. There was no emergence of DAP resistance noted in any of the treatments for R497 or HOU503.

3. Discussion

We developed an effective checkerboard MIC method modified from standard MIC testing to screen for antibiotic and phage synergy and identify optimal concentrations of each to best prevent bacterial growth, including at very-low-phage MOI. While the modified checkerboard MIC method provides information pertaining to the inhibition of bacterial growth, it is not meant to measure bacterial killing, and it does provide information on antibiotic effects over multiple time points within a 24-hour period, as seen with TKAs. However, our use of the modified checkerboard MIC method in this study facilitated the accurate and higher throughput screening of effective phage and antibiotic combinations at specific concentrations and phage MOI to then use in TKAs. The data from our modified checkerboard MICs were successful at predicting the TKA results, as demonstrated by multiple phage and antibiotic combinations. For example, in modified checkerboard MIC against R497, DAP-AMP displayed additive activity with an FIC index of 1, and phage-DAP-AMP demonstrated synergy with an FIC index of ≤0.5. In TKA, both DAP-AMP and phage-DAP-AMP combinations demonstrated synergistic activity; however, phage-DAP-AMP achieved killing to the detection limit, while DAP-AMP did not, aligning with the additional killing demonstrated in modified checkerboard MIC by phage-DAP-AMP compared to DAP-AMP. The close alignment between the modified checkerboard MIC data and the TKA results was also demonstrated by other combinations, including phage-AMP and phage-DAP combinations against R497, further validating the use of this method to screen for antibiotic and phage killing effects and possible synergy.
Additionally, we described conserved killing across several different DAP-β-lactam-phage combinations, even when the DAP-β-lactam combinations alone failed against DAP-resistant and DAP-SDD clinical E. faecium strains, R497 and HOU503, respectively. As shown previously, DAP plus β-lactam combinations have demonstrated bactericidal activity for some E. faecium strains while at the same time demonstrating no activity against others. Thus, our goal was to explore the fundamental questions of whether adjunctive phage administration results in antibiotic enhancement. Here, we demonstrated that in 24-hour biofilm TKA against R497, all the combinations of phage-DAP-AMP, phage-DAP-CPT, and phage-DAP-ERT were synergistic and bactericidal compared to any single agent. We previously demonstrated the significant activity of DAP plus AMP against R497 in the planktonic state, while DAP plus ERT or CPT previously showed no activity [45,46]. Notably, the addition of subinhibitory concentrations of phage to AMP had significant activity against DAP-resistant R497 compared to either phage or AMP alone, thus highlighting this combination for further studies. For HOU503, the combination of phage-DAP-AMP and DAP-AMP was additive, but not synergistic, despite increased killing at lower antibiotic concentrations with the addition of phage at an MOI of 10−8. This led to the hypothesis that phage 113 may enhance antibiotic activity in HOU503, but to a lesser extent than in R497.
Furthermore, combinations of phage-DAP-AMP and phage-DAP-CPT prevented treatment-emergent antibiotic and phage resistance in R497, while bacteriophage resistance was observed with R497 in treatments with phage alone and phage-ERT. These data indicate that the addition of DAP, CPT, or AMP to bacteriophage 113 prevents treatment-emergent bacteriophage resistance. As shown previously in the planktonic state [36], the bacterial strain with greater phage susceptibility (R497) showed less emergence of bacteriophage resistance compared to a strain with lower phage susceptibility (HOU503), although an analysis of the emergence of resistance over a longer period may be necessary to confirm these initial findings. The role phage cocktails might play in circumventing phage resistance in the presence of antibiotics warrants further study. Furthermore, the genetic analysis of additional E. faecium strains, as well as bacteriophages, would provide further insight into the trends we report here.
In summary, we show that in instances of multidrug-resistant E. faecium, bacteriophages in combination with DAP and β-lactams are a promising option for eradicating biofilm-mediated infections and preventing the emergence of antibiotic and phage resistance. Additional complementary studies are indicated to examine the addition of multiple phages in a cocktail, as well as further analysis of phage-antibiotic interactions for their potential for in vivo use.

4. Materials and Methods

4.1. Bacterial Isolates

A panel of five clinical E. faecium strains isolated from patients with bacteremia and selected from the Anti-Infective Research Laboratory library, including those with well-characterized liaFSR mutations, were evaluated for further experiments.

4.2. Antimicrobial Agents and Media

DAP, AMP, and ERT were obtained commercially from Sigma Chemical Company (St. Louis, MO, USA), and CPT analytical powder was obtained from Allergan Pharmaceuticals (Parsippany, NJ, USA). Tryptic soy broth supplemented with 1% glucose (GSTSB) was used for each 24-hour incubation phase immediately prior to the start of experiments. Brain heart infusion (BHI) broth (Difco, Detroit, MI, USA) with 50 mg/L calcium and 12.5 mg/L magnesium was used for susceptibility testing and time-kill analyses (TKAs). To prepare 0.5 and 1.5% BHI agar, respective weight percentages of agar (Oxoid, Lenexa, KS, USA) were added to the broth. For all experiments with DAP, an additional 25 mg/L of calcium was added to the broth due to dependency of DAP on calcium for antimicrobial activity [47].

4.3. Bacteriophage Source and Propagation

E. faecium bacteriophage 113 (ATCC 19950-B1) and propagating organism E. faecium (ATCC 19950) were purchased commercially from ATCC (Manassas, VA, USA). Phage propagation was completed to yield high-titer stocks for use in resistance testing and TKA. An underlay of BHI agar (1.5%) was poured into square petri plates. Next, a 5 mL overlay of 0.5% BHI agar was briefly combined with 20 µL of an overnight host E. faecium bacterial culture containing approximately 109 CFU/mL and poured atop the underlay layer. After the overlay was solidified, 500 µL of purified liquid bacteriophage was spread over top and incubated at 37 °C overnight. The overlay agar was scraped into 3 mL of phosphate buffered saline (PBS) + 10 mM magnesium sulfate and centrifuged at 1000 rpm for 25 min at 4 °C. The supernatant was filtered and stored, covered, at 2–8 °C for experimental use [48,49].

4.4. Biofilm Quantification Assay

E. faecium strains were proven to produce biofilm by biofilm quantification techniques similar to those previously described [43,44]. In brief, E. faecium biofilm was formed in a 96-well flat-bottom microtiter tray by incubation overnight at 37 °C in a shaker incubator. The following day, the tray was rinsed using sterile H2O and the biofilm was fixed to the tray overnight. Following this, the samples were treated with crystal violet solution (0.2%) for 30 min to allow the dye to penetrate the biofilm. The excess crystal violet was subsequently removed, samples were washed, and the plates were treated with glacial acetic acid (33%) to re-solubilize the biofilm. Biofilm production was read at OD560 and samples were analyzed for production compared to wells containing media alone, which was used as a negative control. Biofilm quantification was corrected by subtracting the average OD560 of the media containing only wells that had been treated the same as the wells with biofilm [50]. High, medium, and low categories were determined in relation to the negative control, where high biofilm production was defined as a 3-fold increase from the negative control, medium biofilm production was defined as a 2-fold increase from the negative control, and low biofilm production was defined as a 0.5-fold or lower increase from the negative control.

4.5. Phage Sensitivity Assay

Susceptibility of five E. faecium strains against Phage 113 was tested using spot testing, where 10-fold serial dilutions of phage were spotted onto 0.5% BHI overlay plates containing overnight culture of the target bacteria. Plates contained 5 mL of overlay agar, which was briefly mixed with 20 microliters of purified phage onto the bacterial lawn. The plates were incubated overnight following drop testing, and the plaque counts were read the following day [42]. Results were qualified as clear, turbid, or no effect of phage (resistant). Susceptible strains were further evaluated via plaque assay to determine efficiency of plating (EOP) as a ratio of plaque forming units (PFU) of the susceptible organism to PFU’s of the reference organism (E. faecium strain ATCC 19950) [42]. Phage susceptibility was classified as high, medium, or low based on plaque-forming unit (PFU) counts, where >107 was defined as high, between 103 and 107 was defined as medium, and <103 PFU/mL was defined as low susceptibility. Phage nonsusceptibility was defined as no visual detection of individual phage plaques and/or no bacterial lawn clearance.

4.6. Antibiotic Susceptibility Testing

Minimum biofilm inhibitory concentration (MBIC) values were determined in duplicate using the pin-lid method with similar steps to those described previously using the Calgary Biofilm Device (CBD) [45,51,52]. Briefly, a starting inoculum of 1 × 106 CFU/mL in glucose-supplemented TSB (GSTSB) was placed in a 96-well microtiter plate, covered with a p6-peg lid, then incubated for 18–24 h at 37 °C for E. faecium biofilm formation on the pegs. Next, the 96-peg lid was removed, rinsed with sterile PBS to remove planktonic bacteria, and transferred to a separate 96-well microtiter plate containing serial dilutions of a single antibiotic using a broth microdilution (BMD) technique, as described by the Clinical and Laboratory Standards Institute (CLSI), except that brain heart infusion (BHI) was used for BMD instead of Mueller–Hinton broth reports and incubated for 24 h at 37 °C [53,54,55]. Following incubation, the peg lid was removed and the MBIC recorded and defined as the column of wells with the lowest antibiotic concentration and without bacterial growth.

4.7. Modified Checkerboard for Antibiotic and Bacteriophage Synergy Screening

Antibiotic and bacteriophage synergy in R497 and HOU503 was determined by analyzing the activity of antibiotics and phage 113 in combination compared to their individual activities using a modified checkerboard assay. Initially, a single antibiotic was serially diluted 2-fold through a 96-well round-bottom microtiter tray as was performed for MIC testing, with a starting concentration of 2xMIC for each antibiotic. A second antibiotic was serially diluted 2-fold in the perpendicular direction in a secondary 96-well tray. Antibiotic dilutions in checkerboards containing phage were applied in the same manner as previously described; however, phage dilutions were performed 10-fold rather than 2-fold to achieve a wide range of MOI. Once dilutions were complete, 50 µL from each dilution well of the secondary tray was deposited into the corresponding wells of the original tray. To the original tray, 50 µL each of broth and isolate bacterial stock was added and the plate was incubated at 37 °C for 18–24 h, and then read at OD600. In triple-therapy checkerboards assessing synergy of DAP plus AMP in the presence of constant phage, phage 113 was added to each well, excluding controls, at a subinhibitory concentration of 10−8 PFU/mL. Synergy, additive activity, and antagonism were defined as a calculated FIC index of ≤0.5, FIC of 1–4, and >4, respectively [37,56].

4.8. Time Kill Analyses

TKA were performed in microwell plates to evaluate antibiotic and phage synergy against biofilm-producing organisms, as previously described [57]. In brief, four 3-millimeter polyurethane beads were placed in each well with 2 mL of 1% GSTSB, inoculated with the test organism, and incubated at 37 °C, allowing biofilm formation. Targeted bacterial starting inoculum for both E. faecium strains was 6.5–7 log10 CFU/mL. After 24 h of incubation, GSTSB was aspirated from each well and replaced with calcium-supplemented BHI broth. Antimicrobials were added at 0.5× the MBIC values (DAP, ERT) or free physiological peak concentration (AMP free peak = 72 µg/mL, CPT free peak = 13.2 µg/mL), whichever was lower, to simulate sub-inhibitory concentrations. Phage dosing was optimized to a multiplicity of infection (MOI) ratio of 1, which represents the ratio of phage to target organism. Beads were removed with sterile forceps at 0 (prior to adding antibiotic and/or phage), 4, 8, and 24 h to create a growth curve. Beads were washed with 1 mL of 0.9% sodium chloride and processed for 6 min alternating in 1-minute intervals each of vortexing and sonication at 20 Hz (Bransonic 12 Branson Ultrasonic Corporation) to disrupt the biofilm structure and recover bacterial cells. Samples were appropriately diluted with 0.9% sodium chloride to eliminate antibiotic carryover [58]. Samples containing phage were then centrifuged and filtered to separate phage and bacteria for counting. The collected bacterial samples were serially diluted appropriately, plated on BHI agar (easySpiral, Interscience for Microbiology, Saint Nom la Breteche, France) with a detection limit of 102 CFU/mL, and incubated for 24 h at 37 °C; bacterial colonies were counted using a laser colony counter (Scan 1200, Interscience for Microbiology, Saint Nom la Breteche, France). Synergy was defined as a  ≥2 log10 CFU/mLkill compared to the most effective agent (or double-combination regimen) alone at 24 h. Bactericidal activity was defined as a  ≥3 log10 CFU/mL reduction from baseline.
Combinations that resulted in ≥1 log10 bacterial growth in comparison to the least active single agent were considered antagonistic. Single drug/phage exposures in biofilm TKA included DAP, AMP, CPT, ERT, and phage 113. Additionally, combination evaluations were performed with DAP plus each of the non-DAP antimicrobials. Statistical analysis was carried out using SPSS version 21.0 (IBM Corp., Armonk, NY, USA) software. One-way analysis of variance (ANOVA) with Tukey’s multiple comparison test was used to compare changes in CFU/mL between regimens, where significance was considered at p < 0.05.

4.9. Resistance Testing

Each 24 h TKA liquid sample was tested for the emergence of antibiotic and phage resistance, as previously described [22,42,48,59]. The modified BIM (bacteriophage insensitive mutants) test was used to determine the frequency of spontaneous resistance (FOR) [48]. First, BHI agar underlay was poured into 90-millimeter round plates. Following this, 10 µL of each sample was mixed with 100 µL high-titer phage (at ≥109 PFU/mL) and incubated for 10 min at 37 °C. After incubation, this was briefly mixed with 3 mL of 0.5% BHI agar overlay and poured onto the prepared underlay plates. Plates were then incubated for 24 h at 37 °C and bacterial colonies were counted. After counting, plates were left out at room temperature overnight and counted again at 48 h. FOR was calculated as the number of colonies on each plate divided by the number of bacteria in the replicate for both 24- and 48-hour time points. After 48 hours, a single colony was picked from each plate if available, and sub-cultured in BHI liquid culture for 6–7 h at 37 °C [48].
The double-drop method was used after completion of the culture to verify phage sensitivity [42,48]. Using standard agar plates, 10 µL of high titer phage (109 PFU/mL) was dropped onto 5 µL of sub-cultured bacterial samples. Plates were incubated for 24 h at 37 °C and spots were compared to the negative control (made using PBS instead of high-titer phage). Comparatively, samples were labeled as R (resistant, no visible difference from negative control spot), turbid (plaques formed with EOP of 0.001–0.01; T), or S (clear plaques with EOP of 0.1–1; C) [48].

4.10. Phage Quantification

Bacteriophages were quantified using a modified small-drop agar overlay method [42]. A 20 µL sample of reference organism cultured for 16–18 h overnight was briefly suspended with 5 mL of 0.5% BHI agar and poured over a 1.5% BHI agar underlay. This was left to set for 10 min. Following this, 5 µL of 10× serial dilutions of filtered phage sample was spotted over top of the bacterial lawn and incubated overnight at 37 °C. Plaques were counted the following day after 16–18 h of incubation.

Author Contributions

K.L., A.J.K.C., R.K., T.M., K.S., D.J.H., G.S.C., B.A.D., C.A.A. and M.J.R. were involved in the conceptualization, methodology, validation, formal analysis, investigation, writing—original draft preparation, and writing—review and editing — of this manuscript. All authors have read and agreed to the published version of the manuscript.

Funding

This research received no external funding. M.J.R. is supported by NIH grants R21 AI163726. C.A.A. is supported by NIH grants K24AI121296, R01AI134637, R01AI48342, and P01AI152999.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

Not applicable.

Acknowledgments

We would like to acknowledge Allergan (Abbvie) Pharmaceuticals for providing analytical ceftaroline powder for our experiments.

Conflicts of Interest

K.L., A.J.K.C., R.K., T.M., K.C.S., D.J.H., G.S.C., C.A.A. and M.J.R. have nothing to declare. M.J.R. has received grant support from and has consulted or spoken on behalf of Allergan, Melinta, Merck, Paratek, Shionogi, Spero, and Tetraphase. C.A.A. has received grant support from Merck and MeMed Diagnostics, and is a co-founder of Entasis Therapeutics and shareholder in Ancilia Biosciences, C.

References

  1. Arias, C.A.; Murray, B.E. The rise of the Enterococcus: Beyond vancomycin resistance. Nat. Rev. Microbiol. 2012, 10, 266–278. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  2. Carmeli, Y.; Eliopoulos, G.; Mozaffari, E.; Samore, M. Health and Economic Outcomes of Vancomycin-Resistant Enterococci. Arch. Intern. Med. 2002, 162, 2223–2228. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  3. O’Driscoll, T.; Crank, C.W. Vancomycin-resistant enterococcal infections: Epidemiology, clinical manifestations, and optimal management. Infect. Drug Resist. 2015, 8, 217–230. [Google Scholar]
  4. Centers for Disease Control and Prevention (U.S.). Antibiotic Resistance Threats in the United States. 2019. Available online: https://stacks.cdc.gov/view/cdc/82532 (accessed on 3 May 2021).
  5. Chiang, H.-Y.; Perencevich, E.N.; Nair, R.; Nelson, R.E.; Samore, M.; Khader, K.; Chorazy, M.L.; Herwaldt, L.A.; Blevins, A.; Ward, M.A.; et al. Incidence and Outcomes Associated with Infections Caused by Vancomycin-Resistant Enterococci in the United States: Systematic Literature Review and Meta-Analysis. Infect. Control Hosp. Epidemiol. 2017, 38, 203–215. [Google Scholar] [CrossRef] [PubMed]
  6. Jabbari Shiadeh, S.M.; Pormohammad, A.; Hashemi, A.; Lak, P. Global prevalence of antibiotic resistance in blood-isolated Enterococcus faecalis and Enterococcus faecium: A systematic review and meta-analysis. Infect. Drug Resist. 2019, 12, 2713–2725. [Google Scholar] [CrossRef] [Green Version]
  7. Weiner, L.M.; Webb, A.K.; Limbago, B.; Dudeck, M.A.; Patel, J.; Kallen, A.J.; Edwards, J.R.; Sievert, D.M. Antimicrobial-Resistant Pathogens Associated with Healthcare-Associated Infections: Summary of Data Reported to the National Healthcare Safety Network at the Centers for Disease Control and Prevention, 2011–2014. Infect. Control. Hosp. Epidemiol. 2016, 37, 1288–1301. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  8. Paganelli, F.L.; Willems, R.J.L.; Jansen, P.; Hendrickx, A.; Zhang, X.; Bonten, M.J.M.; Leavis, H.L. Enterococcus faecium Biofilm Formation: Identification of Major Autolysin AtlA Efm, Associated Acm Surface Localization, and AtlA Efm-Independent Extracellular DNA Release. MBio 2013, 4, e00154-13. [Google Scholar] [CrossRef] [Green Version]
  9. Taglialegna, A.; Matilla-Cuenca, L.; Dorado-Morales, P.; Navarro, S.; Ventura, S.; Garnett, J.A.; Lasa, I.; Valle, J. The biofilm-associated surface protein Esp of Enterococcus faecalis forms amyloid-like fibers. NPJ Biofilms Microbiomes. 2020, 6, 15. [Google Scholar] [CrossRef] [Green Version]
  10. Ramadhan, A.A.; Hegedus, E. Biofilm formation and esp gene carriage in enterococci. J. Clin. Pathol. 2005, 58, 685–686. [Google Scholar] [CrossRef] [Green Version]
  11. Sutherland, I.W. Biofilm exopolysaccharides: A strong and sticky framework. Microbiology 2001, 147, 3–9. [Google Scholar] [CrossRef] [Green Version]
  12. Mosselhy, D.A.; Assad, M.; Sironen, T.; Elbahri, M. Nanotheranostics: A Possible Solution for Drug-Resistant Staphylococcus aureus and their Biofilms? Nanomaterials 2021, 11, 82. [Google Scholar] [CrossRef] [PubMed]
  13. Kheir, M.M.; Tan, T.L.; Higuera, C.; George, J.; Valle Della, C.J.; Shen, M.; Parvizi, J. Periprosthetic Joint Infections Caused by Enterococci Have Poor Outcomes. J. Arthroplasty 2017, 32, 933–947. [Google Scholar] [CrossRef] [PubMed]
  14. Holmberg, A.; Rasmussen, M. Mature biofilms of Enterococcus faecalis and Enterococcus faecium are highly resistant to antibiotics. Diagn. Microbiol. Infect. Dis. 2016, 84, 19–21. [Google Scholar] [CrossRef] [PubMed]
  15. Mascio, C.T.M.; Alder, J.D.; Silverman, J.A. Bactericidal Action of Daptomycin against Stationary-Phase and Nondividing Staphylococcus aureus Cells. Antimicrob. Agents Chemother. 2007, 51, 4255–4260. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  16. Jahanbakhsh, S.; Singh, N.B.; Yim, J.; Kebriaei, R.; Smith, J.R.; Lev, K.; Tran, T.T.; Rose, W.E.; Arias, C.A.; Rybak, M.J. Impact of Daptomycin Dose Exposure Alone or in Combination with β-Lactams or Rifampin against Vancomycin-Resistant Enterococci in an In Vitro Biofilm Model. Antimicrob. Agents Chemother. 2020, 64, e02074-19. [Google Scholar] [CrossRef] [PubMed]
  17. Müller, A.; Wenzel, M.; Strahl, H.; Grein, F.; Saaki, T.N.V.; Kohl, B.; Siersma, T.; Bandow, J.E.; Sahl, H.-G.; Schneider, T.; et al. Daptomycin inhibits cell envelope synthesis by interfering with fluid membrane microdomains. Proc. Natl. Acad. Sci. USA 2016, 113, E7077–E7086. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  18. Stewart, P.S.; Davison, W.M.; Steenbergen, J.N. Daptomycin Rapidly Penetrates a Staphylococcus epidermidis Biofilm. Antimicrob. Agents Chemother. 2009, 53, 3505–3507. [Google Scholar] [CrossRef] [Green Version]
  19. Lellek, H.; Franke, G.C.; Ruckert, C.; Wolters, M.; Wolschke, C.; Christner, M.; Büttner, H.; Alawi, M.; Kröger, N.; Rohde, H. Emergence of daptomycin non-susceptibility in colonizing vancomycin-resistant Enterococcus faecium isolates during daptomycin therapy. Int. J. Med. Microbiol. 2015, 305, 902–909. [Google Scholar] [CrossRef]
  20. Munita, J.M.; Murray, B.E.; Arias, C.A. Daptomycin for the treatment of bacteraemia due to vancomycin-resistant enterococci. Int. J. Antimicrob. Agents 2014, 44, 387–395. [Google Scholar] [CrossRef] [Green Version]
  21. Kamboj, M.; Cohen, N.; Gilhuley, K.; Babady, N.E.; Seo, S.K.; Sepkowitz, K.A. Emergence of Daptomycin-Resistant VRE: Experience of a Single Institution. Infect. Control Hosp. Epidemiol. 2011, 32, 391–394. [Google Scholar] [CrossRef]
  22. Kebriaei, R.; Rice, S.A.; Singh, K.V.; Stamper, K.C.; Dinh, A.Q.; Rios, R.; Diaz, L.; Murray, B.E.; Munita, J.M.; Tran, T.T.; et al. Influence of Inoculum Effect on the Efficacy of Daptomycin Monotherapy and in Combination with β-Lactams against Daptomycin-Susceptible Enterococcus faecium Harboring LiaSR Substitutions. Antimicrob. Agents Chemother. 2018, 62, e00315-18. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  23. Hall, A.D.; Steed, M.E.; Arias, C.A.; Murray, B.E.; Rybak, M.J. Evaluation of Standard- and High-Dose Daptomycin versus Linezolid against Vancomycin-Resistant Enterococcus Isolates in an In Vitro Pharmacokinetic/Pharmacodynamic Model with Simulated Endocardial Vegetations. Antimicrob. Agents Chemother. 2012, 56, 3174–3180. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  24. Prater, A.G.; Mehta, H.H.; Beabout, K.; Suoandy, A.; Miller, W.R.; Tran, T.T.; Arias, C.A.; Shamoo, Y. Daptomycin Resistance in Enterococcus faecium Can Be Delayed by Disruption of the LiaFSR Stress Response Pathway. Antimicrob. Agents Chemother. 2021, 65, e01317-20. [Google Scholar] [CrossRef] [PubMed]
  25. Diaz, L.; Tran, T.T.; Munita, J.M.; Miller, W.R.; Rincon, S.; Carvajal, L.P.; Wollam, A.; Reyes, J.; Panesso, D.; Rojas, N.L.; et al. Whole-Genome Analyses of Enterococcus faecium Isolates with Diverse Daptomycin MICs. Antimicrob. Agents Chemother. 2014, 58, 4527–4534. [Google Scholar] [CrossRef] [Green Version]
  26. Morrisette, T.; Kebriaei, R.; Lev, K.L.; Morales, S.; Rybak, M.J. Bacteriophage Therapeutics: A Primer for Clinicians on Phage-Antibiotic Combinations. Pharmacotherapy 2020, 40, 153–168. [Google Scholar] [CrossRef] [Green Version]
  27. Kumaran, D.; Taha, M.; Yi, Q.; Ramirez-Arcos, S.; Diallo, J.-S.; Carli, A.; Abdelbary, H. Does Treatment Order Matter? Investigating the Ability of Bacteriophage to Augment Antibiotic Activity against Staphylococcus aureus Biofilms. Front. Microbiol. 2018, 9, 127. [Google Scholar] [CrossRef] [Green Version]
  28. Harper, D.R.; Parracho, H.M.R.T.; Walker, J.; Sharp, R.; Hughes, G.; Werthén, M.; Lehman, S.; Morales, S. Bacteriophages and Biofilms. Antibiotics 2014, 3, 270–284. [Google Scholar] [CrossRef]
  29. Parasion, S.; Kwiatek, M.; Gryko, R.; Mizak, L.; Malm, A. Bacteriophages as an alternative strategy for fighting biofilm development. Pol. J. Microbiol. 2014, 63, 137–145. [Google Scholar] [CrossRef]
  30. Tkhilaishvili, T.; Lombardi, L.; Klatt, A.-B.; Trampuz, A.; Luca, M.D. Bacteriophage Sb-1 enhances antibiotic activity against biofilm, degrades exopolysaccharide matrix and targets persisters of Staphylococcus aureus. Int. J. Antimicrob. Agents 2018, 52, 0842–0853. [Google Scholar] [CrossRef]
  31. Azeredo, J.; García, P.; Drulis-Kawa, Z. Targeting biofilms using phages and their enzymes. Curr. Opin. Biotechnol. 2021, 68, 251–261. [Google Scholar] [CrossRef]
  32. Dakheel, K.H.; Abdul Rahim, R.; Al-Obaidi, J.R.; Neela, V.K.; Hun, T.G.; Isa, M.N.M.; Razali, N.; Yusoff, K. Proteomic analysis revealed the biofilm-degradation abilities of the bacteriophage UPMK_1 and UPMK_2 against Methicillin-resistant Staphylococcus aureus. Biotechnol. Lett. 2022, 1–10. [Google Scholar] [CrossRef] [PubMed]
  33. Pires, D.P.; Melo, L.; Vilas Boas, D.; Sillankorva, S.; Azeredo, J. Phage therapy as an alternative or complementary strategy to prevent and control biofilm-related infections. Curr. Opin. Microbiol. 2017, 39, 48–56. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  34. Segall, A.M.; Roach, D.R.; Strathdee, S.A. Stronger together? Perspectives on phage-antibiotic synergy in clinical applications of phage therapy. Curr. Opin. Microbiol 2019, 51, 46–50. [Google Scholar] [CrossRef] [PubMed]
  35. Li, X.; He, Y.; Wang, Z.; Wei, J.; Hu, T.; Si, J.; Tao, G.; Zhang, L.; Xie, L.; Abdalla, A.E.; et al. A combination therapy of Phages and Antibiotics: Two is better than one. Int. J. Biol. Sci. 2021, 17, 3573–3582. [Google Scholar] [CrossRef] [PubMed]
  36. Kebriaei, R.; Lev, K.; Morrisette, T.; Stamper, K.C.; Abdul-Mutakabbir, J.C.; Lehman, S.M.; Morales, S.; Rybak, M.J. Bacteriophage-Antibiotic Combination Strategy: An Alternative against Methicillin-Resistant Phenotypes of Staphylococcus aureus. Antimicrob. Agents Chemother. 2020, 64, e00461-20. [Google Scholar] [CrossRef] [PubMed]
  37. Rodriguez-Gonzalez, R.A.; Leung, C.Y.; Chan, B.K.; Turner, P.E.; Weitz, J.S. Quantitative Models of Phage-Antibiotic Combination Therapy. mSystems 2020, 5, e00756-19. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  38. Torres-Barceló, C.; Hochberg, M.E. Evolutionary Rationale for Phages as Complements of Antibiotics. Trends Microbiol. 2016, 24, 249–256. [Google Scholar] [CrossRef]
  39. Morrisette, T.; Lev, K.L.; Kebriaei, R.; Abdul-Mutakabbir, J.C.; Stamper, K.C.; Morales, S.; Lehman, S.M.; Canfield, G.S.; Duerkop, B.A.; Arias, C.A.; et al. Bacteriophage-Antibiotic Combinations for Enterococcus faecium with Varying Bacteriophage and Daptomycin Susceptibilities. Antimicrob. Agents Chemother. 2020, 64, e00993-20. [Google Scholar] [CrossRef]
  40. Shlezinger, M.; Coppenhagen-Glazer, S.; Gelman, D.; Beyth, N.; Hazan, R. Eradication of Vancomycin-Resistant Enterococci by Combining Phage and Vancomycin. Viruses 2019, 11, 954. [Google Scholar] [CrossRef] [Green Version]
  41. Canfield, G.S.; Chatterjee, A.; Espinosa, J.; Mangalea, M.R.; Sheriff, E.K.; Keidan, M.; McBride, B.D.; Hang, H.C.; Duerkop, B.A. Lytic Bacteriophages Facilitate Antibiotic Sensitization of Enterococcus faecium. Antimicrob. Agents Chemother. 2021, 65, e00143-21. [Google Scholar] [CrossRef]
  42. Mazzocco, A.; Waddell, T.E.; Lingohr, E.; Johnson, R.P. Enumeration of Bacteriophages Using the Small Drop Plaque Assay System. In Bacteriophages; Clokie, M.R.J., Kropinski, A.M., Eds.; Humana Press: Totowa, NJ, USA, 2009; pp. 81–85. Available online: http://link.springer.com/10.1007/978-1-60327-164-6_9 (accessed on 13 January 2022).
  43. Stepanović, S.; Vuković, D.; Dakić, I.; Savić, B.; Švabić-Vlahović, M. A modified microtiter-plate test for quantification of staphylococcal biofilm formation. J. Microbiol. Methods 2000, 40, 175–179. [Google Scholar] [CrossRef]
  44. Christensen, G.D.; Simpson, W.A.; Younger, J.J.; Baddour, L.M.; Barrett, L.M.; Melton, D.M.; Beachey, E.H. Adherence of coagulase-negative staphylococci to plastic tissue culture plates: A quantitative model for the adherence of staphylococci to medical devices. J. Clin. Microbiol. 1985, 22, 996–1006. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  45. Ceri, H.; Olson, M.; Morck, D.; Storey, D.; Read, R.; Olson, B. The MBEC assay system: Multiple equivalent biofilms for antibiotic and biocide susceptibility testing. Methods Enzymol. 2001, 337, 377–385. [Google Scholar] [PubMed]
  46. Kebriaei, R.; Stamper, K.C.; Singh, K.V.; Khan, A.; Rice, S.A.; Dinh, A.Q.; Tran, T.T.; Murray, B.E.; Arias, C.A.; Rybak, M.J. Mechanistic Insights Into the Differential Efficacy of Daptomycin Plus β-Lactam Combinations Against Daptomycin-Resistant Enterococcus faecium. J. Infect. Dis. 2020, 222, 1531–1539. [Google Scholar] [CrossRef] [PubMed]
  47. Grein, F.; Müller, A.; Scherer, K.M.; Liu, X.; Ludwig, K.C.; Klöckner, A.; Strach, M.; Sahl, H.-G.; Kubitscheck, U.; Schneider, T. Ca2+-Daptomycin targets cell wall biosynthesis by forming a tripartite complex with undecaprenyl-coupled intermediates and membrane lipids. Nat. Commun. 2020, 11, 1455. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  48. Lehman, S.; Mearns, G.; Rankin, D.; Cole, R.A.; Smrekar, F.; Branston, S.D.; Morales, S. Design and Preclinical Development of a Phage Product for the Treatment of Antibiotic-Resistant Staphylococcus aureus Infections. Viruses 2019, 11, 88. [Google Scholar]
  49. Bonilla, N.; Rojas, M.I.; Netto Flores Cruz, G.; Hung, S.-H.; Rohwer, F.; Barr, J.J. Phage on tap–a quick and efficient protocol for the preparation of bacteriophage laboratory stocks. Peer J. 2016, 4, e2261. [Google Scholar] [CrossRef] [Green Version]
  50. Bose. J.L.; Lehman, M.K.; Fey, P.D.; Bayles, K.W. Contribution of the Staphylococcus aureus Atl AM and GL Murein Hydrolase Activities in Cell Division, Autolysis, and Biofilm Formation. PLoS ONE 2012, 7, e42244. [Google Scholar]
  51. Ceri, H.; Olson, M.E.; Stremick, C.; Read, R.R.; Morck, D.; Buret, A. The Calgary Biofilm Device: New technology for rapid determination of antibiotic susceptibilities of bacterial biofilms. J. Clin. Microbiol. 1999, 37, 1771–1776. [Google Scholar] [CrossRef] [Green Version]
  52. Ali, L.; Khambaty, F.; Diachenko, G. Investigating the suitability of the Calgary Biofilm Device for assessing the antimicrobial efficacy of new agents. Bioresour. Technol. 2006, 97, 1887–1893. [Google Scholar] [CrossRef]
  53. Clinical and Laboratory Standards Institute. Methods for Dilution Antimicrobial Susceptibility Tests for Bacteria That Grow Aerobically; Approved Standard—Tenth Edition. Available online: https://clsi.org/media/1632/m07a10_sample.pdf (accessed on 13 January 2022).
  54. Humphries, R.M.; Ambler, J.; Mitchell, S.L.; Castanheira, M.; Dingle, T.; Hindler, J.A.; Koeth, L.; Sei, K. CLSI Methods Development and Standardization Working Group Best Practices for Evaluation of Antimicrobial Susceptibility Tests. J. Clin. Microbiol. 2018, 56, e01934-17. [Google Scholar] [CrossRef] [Green Version]
  55. Rice, L.B.; Carias, L.L.; Rudin, S.; Hutton, R.; Marshall, S.; Hassan, M.; Josseaume, N.; Dubost, L.; Marie, A.; Arthur, M. Role of Class A Penicillin-Binding Proteins in the Expression of β-Lactam Resistance in Enterococcus faecium. J. Bacteriol. 2009, 191, 3649–3656. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  56. Zhvania, P.; Hoyle, N.S.; Nadareishvili, L.; Nizharadze, D.; Kutateladze, M. Phage Therapy in a 16-Year-Old Boy with Netherton Syndrome. Front Med. 2017, 4, 94. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  57. Barber, K.E.; Werth, B.J.; McRoberts, J.P.; Rybak, M.J. A Novel Approach Utilizing Biofilm Time-Kill Curves To Assess the Bactericidal Activity of Ceftaroline Combinations against Biofilm-Producing Methicillin-Resistant Staphylococcus aureus. Antimicrob. Agents Chemother. 2014, 58, 2989–2992. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  58. Rybak, M.J.; Hershberger, E.; Moldovan, T.; Grucz, R.G. In Vitro Activities of Daptomycin, Vancomycin, Linezolid, and Quinupristin-Dalfopristin against Staphylococci and Enterococci, Including Vancomycin- Intermediate and -Resistant Strains. Antimicrob. Agents Chemother. 2000, 44, 1062–1066. [Google Scholar] [CrossRef] [Green Version]
  59. O’Flynn, G.; Ross, R.P.; Fitzgerald, G.F.; Coffey, A. Evaluation of a Cocktail of Three Bacteriophages for Biocontrol of Escherichia coli O157:H7. Appl. Environ. Microbiol. 2004, 70, 3417–3424. [Google Scholar] [CrossRef] [Green Version]
Antibiotics 11 00392 g001 550
Figure 1. (A–D) Checkerboard analysis of E. faecium strain R497. Additivity, defined as an FIC index >0.5 but <4, is indicated by the red outline in (A). Synergy, defined as an FIC index ≤ 0.5, is indicated by the red outline in (B). Comparisons are versus growth control and depicted by the blue color gradient as percentage of growth.
Figure 1. (A–D) Checkerboard analysis of E. faecium strain R497. Additivity, defined as an FIC index >0.5 but <4, is indicated by the red outline in (A). Synergy, defined as an FIC index ≤ 0.5, is indicated by the red outline in (B). Comparisons are versus growth control and depicted by the blue color gradient as percentage of growth.
Antibiotics 11 00392 g001
Antibiotics 11 00392 g002 550
Figure 2. (A–D) Checkerboard analysis of E. faecium strain HOU503. Additivity, defined as an FIC index > 0.5 but <4, is indicated by the red outline in Figure 1A,B. Comparisons are versus growth control and depicted by the blue color gradient as percentage of growth.
Figure 2. (A–D) Checkerboard analysis of E. faecium strain HOU503. Additivity, defined as an FIC index > 0.5 but <4, is indicated by the red outline in Figure 1A,B. Comparisons are versus growth control and depicted by the blue color gradient as percentage of growth.
Antibiotics 11 00392 g002
Antibiotics 11 00392 g003 550
Figure 3. (A–F) Time-kill analysis of DAP-resistant and DAP-SDD E. faecium strains R497 and HOU503, respectively. GC, growth control. (A) R497 vs. DAP (0.5xMIC) + AMP (free peak concentration = 72) + phage (MOI = 1). (B) R497 vs. DAP (0.5xMIC) + CPT (free peak concentration = 13.2) + phage (MOI = 1) (C) R497 vs. DAP (0.5xMIC) + ERT (free peak concentration = 15.5) + phage (MOI = 1) (D) HOU503 vs. DAP (0.5xMIC) + AMP (free peak concentration = 72) + phage (MOI = 1) (E) HOU503 vs. DAP (0.5xMIC) + CPT (free peak concentration = 13.2) + phage (MOI = 1) (F) R497 vs. DAP (0.5xMIC) + ERT (free peak concentration = 15.5) + phage (MOI = 1). DAP, daptomycin; AMP, ampicillin; CPT, ceftaroline; ERT, ertapenem.
Figure 3. (A–F) Time-kill analysis of DAP-resistant and DAP-SDD E. faecium strains R497 and HOU503, respectively. GC, growth control. (A) R497 vs. DAP (0.5xMIC) + AMP (free peak concentration = 72) + phage (MOI = 1). (B) R497 vs. DAP (0.5xMIC) + CPT (free peak concentration = 13.2) + phage (MOI = 1) (C) R497 vs. DAP (0.5xMIC) + ERT (free peak concentration = 15.5) + phage (MOI = 1) (D) HOU503 vs. DAP (0.5xMIC) + AMP (free peak concentration = 72) + phage (MOI = 1) (E) HOU503 vs. DAP (0.5xMIC) + CPT (free peak concentration = 13.2) + phage (MOI = 1) (F) R497 vs. DAP (0.5xMIC) + ERT (free peak concentration = 15.5) + phage (MOI = 1). DAP, daptomycin; AMP, ampicillin; CPT, ceftaroline; ERT, ertapenem.
Antibiotics 11 00392 g003
Table
Table 1. High, medium, and low phage susceptibility were defined as phage counts of >107, between 103 and 107, and <103 PFU/mL, respectively.
Table 1. High, medium, and low phage susceptibility were defined as phage counts of >107, between 103 and 107, and <103 PFU/mL, respectively.
OrganismBacteriophage Susceptibility
(PFU/mL Compared to Host) a		Biofilm Quantification
(OD Compared to Control) b
R497HighMedium
HOU503MediumHigh
S447 (55)LowNone
SF12047 (56)LowNone
12311 (56)LowLow
a Modified small-drop agar method results. b Crystal violet microtiter plate assay results. PFU, plaque forming units; OD, optical density.
Table
Table 2. Phage resistance check for bacteriophage and antibiotics at the end of 24 h exposure.
Table 2. Phage resistance check for bacteriophage and antibiotics at the end of 24 h exposure.
Bacteriophage Resistance a
RegimenR497HOU503
PhageRR
Phage-DAPSR
Phage-AMPSR
Phage-CPTSR
Phage-ERTRR
Phage-DAP-AMPSS
Phage-DAP-CPTSS
Phage-DAP-ERTSR
a R, resistant; S, sensitive; DAP, daptomycin; AMP, ampicillin; CPT, ceftaroline; ERT, ertapenem.
Publisher’s Note: MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Share and Cite

MDPI and ACS Style

Lev, K.; Kunz Coyne, A.J.; Kebriaei, R.; Morrisette, T.; Stamper, K.; Holger, D.J.; Canfield, G.S.; Duerkop, B.A.; Arias, C.A.; Rybak, M.J. Evaluation of Bacteriophage-Antibiotic Combination Therapy for Biofilm-Embedded MDR Enterococcus faecium. Antibiotics 2022, 11, 392. https://doi.org/10.3390/antibiotics11030392

AMA Style

Lev K, Kunz Coyne AJ, Kebriaei R, Morrisette T, Stamper K, Holger DJ, Canfield GS, Duerkop BA, Arias CA, Rybak MJ. Evaluation of Bacteriophage-Antibiotic Combination Therapy for Biofilm-Embedded MDR Enterococcus faecium. Antibiotics. 2022; 11(3):392. https://doi.org/10.3390/antibiotics11030392

Chicago/Turabian Style

Lev, Katherine, Ashlan J. Kunz Coyne, Razieh Kebriaei, Taylor Morrisette, Kyle Stamper, Dana J. Holger, Gregory S. Canfield, Breck A. Duerkop, Cesar A. Arias, and Michael J. Rybak. 2022. "Evaluation of Bacteriophage-Antibiotic Combination Therapy for Biofilm-Embedded MDR Enterococcus faecium" Antibiotics 11, no. 3: 392. https://doi.org/10.3390/antibiotics11030392

APA Style

Lev, K., Kunz Coyne, A. J., Kebriaei, R., Morrisette, T., Stamper, K., Holger, D. J., Canfield, G. S., Duerkop, B. A., Arias, C. A., & Rybak, M. J. (2022). Evaluation of Bacteriophage-Antibiotic Combination Therapy for Biofilm-Embedded MDR Enterococcus faecium. Antibiotics, 11(3), 392. https://doi.org/10.3390/antibiotics11030392

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop