https://orcid.org/0000-0001-5580-1292
Figures
Abstract
Biofilm-impaired tissue is a significant factor in chronic wounds such as diabetic foot ulcers. Most, if not all, anti-biotics in clinical use have been optimized against planktonic phenotypes. In this study, an in vitro assessment was performed to determine the potential efficacy of a first-in-class series of antibiofilm antibiotics and compare outcomes to current clinical standards of care. The agent, CZ-01179, was formulated into a hydrogel and tested against mature biofilms of a clinical isolate of methicillin-resistant Staphylococcus aureus and Pseudomonas aeruginosa ATCC 27853 using two separate methods. In the first method, biofilms were grown on cellulose discs on an agar surface. Topical agents were spread on gauze and placed over the biofilms for 24 h. Biofilms were quantified and imaged with confocal and scanning electron microscopy. In the second method, biofilms were grown on bioabsorbable collagen coupons in a modified CDC biofilm reactor. Coupons were immersed in treatment for 24 h. The first method was limited in its ability to assess efficacy. Efficacy profiles against biofilms grown on collagen were more definitive, with CZ-01179 gel eradicating well-established biofilms to a greater degree compared to clinical standards. In conclusion, CZ-01179 may be a promising topical agent that targets the biofilm phenotype. Pre-clinical work is currently being performed to determine the translatable potential of CZ-01179 gel.
Citation: Miller M, Rogers JC, Badham MA, Cadenas L, Brightwell E, Adams J, et al. (2020) Examination of a first-in-class bis-dialkylnorspermidine-terphenyl antibiotic in topical formulation against mono and polymicrobial biofilms. PLoS ONE 15(10): e0234832. https://doi.org/10.1371/journal.pone.0234832
Editor: Noreen J. Hickok, Thomas Jefferson University, UNITED STATES
Received: May 28, 2020; Accepted: September 17, 2020; Published: October 19, 2020
This is an open access article, free of all copyright, and may be freely reproduced, distributed, transmitted, modified, built upon, or otherwise used by anyone for any lawful purpose. The work is made available under the Creative Commons CC0 public domain dedication.
Data Availability: All relevant data are within the manuscript and its Supporting Information files.
Funding: DLW received a Merit Review award from the Department of Veterans Affairs that supported this work. Award number is 1I01RX002287-01. The grant submission was peer reviewed, but the funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript. The funder (Curza Global) provided support in the form of salaries for authors (REL, PRS, HRKR, JCR, TJH, DLW), but did not have any additional role in the study design, data collection and analysis, decision to publish, or preparation of the manuscript. The specific roles of these authors are articulated in the ‘author contributions’ section. Authors DLW, PRS, REL and TJH have financial interest in Curza Global, LLC.
Competing interests: I have read the journal's policy and the authors of this manuscript have the following competing interests: There are multiple patents secured by Curza Global on which multiple authors (DLW, REL, PRS, TJH, HRKR) are listed as inventors. This does not alter our adherence to PLOS ONE policies on sharing data and materials.
Introduction
The Centers for Disease Control and Prevention (CDC) label the rapid global growth of drug-resistant pathogens “one of our most serious health threats” [1]. The World Health Organization (WHO) also warns that “without urgent, coordinated action by many stakeholders, the world is headed for a post-antibiotic era, in which common infections and minor injuries which have been treatable for decades can once again kill” [2]. Despite this global public health need, the pipeline for new antibiotics, in particular those that display activity against biofilms, is thin [3, 4].
In 2008 the interagency Antimicrobial Availability Task Force, overseen by the Infectious Disease Society of America (IDSA), identified Pseudomonas aeruginosa and methicillin-resistant Staphylococcus aureus (MRSA), along with four other pathogens, as essential targets to combat antibacterial resistance; these are “ESKAPE” pathogens [4–6]. ESKAPE pathogens represent the paradigms of pathogenesis, transmission, and resistance, and as such, the National Institute of Allergy and Infectious Disease (NIAID) and the IDSA, in coordination with other health organizations, catalyzed initiatives for drug development and research into gene transfer and resistance [2, 7, 8]. These initiatives and promises of funding focus heavily on planktonic-based outcomes, leaving clinicians with minimal alternative options to treat and prevent biofilm-impaired wounds; current selections still constitute topical therapies from the first and second world wars, such as Dakin’s solution and colistin, as treatments for chronic wounds [4, 9–11]. Since 2008 only 7 drugs have been approved by the FDA for treatment of “ESKAPE” pathogens, and none are for treatment of P. aeruginosa [12–22].
Multiple studies show S. aureus to be the most frequent organism responsible for chronic wounds [23–27], but recent studies of chronic wound bacterial profiles provide further evidence supporting the necessity for broad spectrum antibiotic topicals for effective chronic wound therapy [28–30].
An 8-week study at the Wound Healing Center in Copenhagen investigated the bacterial profile of chronic leg ulcers [28]. During the monitoring period, 2 or more bacterial species were identified in 94.4% of the wounds, 4–6 bacterial species were present in 50% of the wounds, and 39% of wounds had more than 6 bacterial species present. The most abundant isolate identified was S. aureus, being present in 93.5% of wounds, while P. aeruginosa was present in 52.2% of wounds. The abundance of bacterial species per wound can contribute to antimicrobial resistance, and more complex polymicrobial biofilms, making treatment less effective.
These results are supported by studies performed with molecular analysis and imaging on chronic and acute wounds from 123 patients [29, 30]. Biofilm prevalence is similar to other wounds studied, with the majority of chronic wounds containing an abundance of diverse biofilms, while acute wounds have few and less diverse biofilms [28, 29]. Molecular analysis provides greater detail of biofilm ecology within chronic wounds, including the presence of strictly anaerobic bacteria, not seen in growth cultures [29, 30].
Topical therapies are valuable as practitioners consider the inherent characteristics of biofilms. Biofilms are well known to be impervious to systemic antibiotic treatments due in part to reduced activity against persister cells in the anaerobic core of the community [31–35]. In addition, achieving sufficient serum concentrations to eradicate biofilms is often impossible [35]. These clinical paradigms and global scenarios warrant two areas of focus: 1) Development of topical antimicrobial technologies that target and eradicate biofilms. Topical products provide local, high doses of antibiotics that can be applied regularly to sustain antimicrobial delivery. Topical delivery also helps maintain a moist wound bed, which facilitates the prevention of tissue dehydration, accelerates angiogenesis, assists in the breakdown of necrotic tissue and/or fibrin, and provides for the transport of cytokines and growth factors [36, 37]. 2) Development of novel antimicrobial agents that address the current global threat of antibiotic resistance.
We tested the in vitro efficacy of a topical formulation, the active component of which is a compound synthesized as part of a first-in-class series of antibiofilm agents (referred to as CZ compounds). More specifically, CZs are designed and synthesized to specifically eradicate, and in some cases disperse, biofilms via non-specific, global, and rapid disruption of Gram-positive and -negative bacteria [38]. CZs, like other polyamine based antibiotics [39], are considered to exert their effect via membrane permeability and disruption of the lipopolysaccharide (LPS) layer. CZs are synthesized by a straightforward and scalable approach and formulate well with polymer and other carrier agents. We formulated CZ-01179 in a gel and tested its ability to eradicate well-established biofilms of MRSA and P. aeruginosa.
Experiments were performed using two separate methods. First, biofilms were grown on cellulose discs following a previously established method by Hammond et al. [40]. Second, biofilms were grown on bioabsorbable collagen in a modified CDC biofilm reactor (Fig 1), then exposed to topical products in a multi-well plate system. We hypothesized that CZ-01179 formulated in a carrier gel would have greater efficacy against biofilms of MRSA and P. aeruginosa in monomicrobial and polymicrobial phenotypes than antibiotic-based clinical standards of care.
(A) Schematic of how a CDC biofilm reactor is set up in general. Source: BioSurface Technologies (B) Image of a fully assembled CDC biofilm reactor. (C) Reactor rods were modified to hold collagen coupons (left panel). A snapshot of the process is shown for removing a collagen coupon on which mature biofilm was grown (right panel). Coupons were rinsed in conical tubes, then used for subsequent analysis. This figure was reused from Williams et al. [41] under the Creative Commons Attribute (CC BY) license.
Materials and methods
Bacterial strains
A clinical isolate of MRSA with known pathogenicity was used; it was isolated from a knee-related infection in a patient and produced positive signals of infection in multiple animal models [42, 43]. P. aeruginosa ATCC 27853 was purchased from the American Type Culture Collection (ATCC). Each organism was passaged and maintained in Luria-Bertani (LB) broth or on Columbia blood agar at 37° C prior to experimentation.
Supplies and reagents
General supplies, reagents, and growth media were purchased from Fisher Scientific (Hampton, NH). An 8-ply 100% cotton gauze was purchased from Kendall Curity®; Coviden (Mansfield, MA). Five clinically-relevant topical products were purchased via the pharmacy at the Department of Veterans Affairs in Salt Lake City: gentamicin sulfate ointment USP, 0.1% (Perrigo Company, Allegan, MI), mupirocin ointment USP, 2% (Glenmark Pharmaceuticals, Mahwah, NJ), silver sulfadiazine cream, USP 1% (Ascend Laboratories, Montvale, NJ), Neosporin® (400 U Bacitracin Zinc– 3.5mg Neomycin Sulfate– 5,000 U Polymixin B Sulfate; Johnson and Johnson, New Brunswick, NJ), Altabax® (retapamulin ointment) 1%, (GlaxoSmithKline, Barnard Castle, County Durham, United Kingdom). Hyaluronic acid (HA; 1.01 MDa– 1.8 MDa) was purchased from Lifecore Biomedical (Chaska, MN; catalog #HA15M-5). This HA is a bacterial fermentation product of Streptococcus pyogenes. Reagents and chemicals for synthesizing CZ-01179 were purchased from Sigma Aldrich (St. Louis, MO). Cellulose discs (6 mm) were purchased from BD (Sparks, MD), and collagen coupons were cut from HeliPlug® Collagen Wound Dressing (Integra LifeSciences, Plainsboro, NJ). BacLight™ Bacterial Viability kits were purchased from Molecular Probes (Eugene, OR). The Nunc™ Lab-Tek™ chamber slide system was purchased from ThermoScientific™ (Waltham, MA).
CZ-01179 synthesis and gel formulation
CZ-01179 was synthesized (Fig 2) by the following method: to a stirring solution of a dicarbaldehyde (5'-(tert-butyl)-[1,1':3',1''-terphenyl]-4,4''-dicarbaldehyde: 2.12 g, 6.22 mmol, 1 equiv.) in MeOH (100 mL) and DCE (25 mL) at 0°C was added the diamine (N1-(3-aminopropyl)-N3-(2-ethylbutyl)propane-1,3-diamine: 3.61 g, 16.8 mmol, 2.7 equiv.) portion-wise over 20 min. The solution was stirred for 16 h. NaBH4 (0.95 g, 24.9, 1 equiv.) was added portion-wise over 20 min and the reaction stirred for an additional 1 h. The solvent was evaporated, and the crude solid partitioned between EtOAc (500 ml) and 10% NaOH (250 ml). The NaOH phase was washed with EtOAc (500 ml), and the combined organics were dried over Na2SO4. Column chromatography was performed using gradient conditions starting at (300:16:1 CH2Cl2:MeOH:NH4OH). The free base was acidified with HCl in MeOH (100 ml) and cooled to 0°C for 1 h. The resulting precipitate was filtered and dried to afford the HCl salt as a white solid (25–52%). Recrystallization with H2O (solvent) and iPrOH (anti-solvent) delivered analytically pure material. 1H NMR (500 MHz, D2O) δ ppm 7.78–7.69 (m, 7H), 7.61 (bs, 4H), 4.38 (s, 4H), 3.26–3.20 (m, 16H), 3.01 (s, 4H), 2.17 (bs, 8H), 1.67 (bs, 2H), 1.38 (bs, 17H), 0.88 (s, 12H). 13C NMR (125 MHz, D2O) δ ppm 153.4, 141.8, 140.4, 130.4, 129.7, 127.8, 123.8, 122.9, 50.9, 50.9, 44.9, 44.6, 43.9, 37.6, 34.4, 30.5, 22.6, 22.4, 22.4, 9.4. IR (neat): 3334 (bs), 2963, 2766, 1457 (all s) cm-1. mp decomposition (180–184°C). LRMS Calculated for C48H80N6 m/z 741.6 [M+H]+, Obsd. 370.7 [M+H]+/2.
CZ-01179 was formulated into a gel following synthesis. CZ-01179 powder was added to sterile PBS; three separate solutions were made with final concentrations of 0.5% (5 mg/mL), 1% (10 mg/mL) or 2% (20 mg/mL). After the CZ-01179 dissolved, HA was added to a final concentration of 1.5% (15 mg/mL). Each gel had the same HA concentration. The gel was mixed thoroughly and allowed to settle for a minimum of 24 h at room temperature before being used for experimentation. At a 2% concentration, pH of the gel was approximately 7.0. Gels with CZ-01179 at 0.5% and 1.0% were initially at pH 4 and were adjusted with NaOH to a pH of approximately 7.0 prior to experimentation. The pH of clinically-relevant gels was not adjusted so as to reflect clinical state.
Cytotoxicity
A minimal essential media (MEM) elution assay was conducted by Nelson Laboratories to determine cytotoxicity profiles (ISO 10993–5) of the 2% CZ-01179 gel and clinical standards. Test articles and controls were extracted in 1x minimal essential media (MEM) with 5% bovine serum for 24–25 h at 37 ± 1°C with agitation. Multiple well cell culture plates were seeded with a verified quantity of industry standard L-929 cells (ATCC CCL-1) and incubated until ~80% confluent. The test articles were held at room temperature for less than four h before testing. The extract fluids were not filtered, centrifuged or manipulated in any way following the extraction process. The test extracts were added to the cell monolayers in triplicate. The cells were incubated at 37 ± 1°C with 5 ± 1% CO2 for 48 ± 3 h.
Cell monolayers were examined and scored (0–4) based on the degree of cellular destruction. Specifically, Grade 0 = No reactivity, no cell lysis; Grade 1 = Slight reactivity, ≤20% rounding, occasional lysis; Grade 2 = Mild reactivity, 20% ≤ 50% rounding, no extensive cell lysis; Grade 3 = Moderate reactivity, 50% ≤ 70% rounding and lysed cells; Grade 4 = Severe reactivity, nearly complete destruction of cell layers. Testing was performed in compliance with US FDA goods and manufacturing practice (GMP) regulations 21 CFR Parts 210, 211 and 820.
Cellulose disc assay
A modified protocol of Hammond et al. was used to grow biofilms on cellulose discs and test efficacy of topical antibiotic products [40]. Isolates were grown in LB broth overnight (~24 h) at 37°C. A 1 mL aliquot was placed into a microcentrifuge tube, pelleted at 12,000 rpm for 5 min, washed, and resuspended in 1 mL of fresh LB as a stock culture. A baseline of colony forming units (CFU)/mL in stock cultures was determined for each experiment using a 10-fold dilution series. The stock concentration of MRSA was ~3.6 x 109 CFU/mL, and for P. aeruginosa was ~4.7 x 109 CFU/mL.
An n = 8 cellulose discs were sterilely and equidistantly placed on the surface of a single LB agar plate. Fifty μL of bacterial suspension were pipetted onto the surface of each disc. Only one isolate was inoculated/plate so as not to have cross contamination. Plates were incubated at 37°C for 48 h to allow biofilms to form on the cellulose material. Notably, this incubation time differed from the Hammond et al. method, which suggested an incubation time of 24 h [40]. The rationale for a longer incubation period was to form more robust biofilms; minimal and immature biofilms were observed by scanning electron microscopy (SEM) with 24 h growth. A more significant bioburden challenge was desired for topical assessment, thus biofilms were grown for 48 h.
Approximately 800 mg of topical agent were spread in a thin layer, i.e., “buttered” on sterile 2” x 2” cotton gauze. The “buttered” side of the gauze pad was placed in contact with the discs such that all n = 8 discs were covered completely. Three additional gauze pads were placed on top of the “buttered” gauze pad; the rationale was to increase the gauze height so that the lid of the Petri dish compressed the stack and simulated pressure of a bandage over a wound (recommended by Hammond et al.) [40]. Masking tape held the Petri dish lid in place and it was incubated for 24 h at 37°C. Biofilms of MRSA were treated with CZ-01179, mupirocin, gentamicin, silver sulfadiazine, retapamulin, or Neosporin®. Biofilms of P. aeruginosa were treated with CZ-01179, gentamicin, silver sulfadiazine, or Neosporin® (retapamulin and mupirocin are indicated for Gram-positive organisms).
Cellulose discs were sterilely removed and placed individually into 1 mL of PBS. Samples were vortexed for 1 min, sonicated for 10 min at 42 kHz and plated using a 10-fold dilution series to quantify the CFU/disc that remained after treatment. Positive controls of growth (n = 8) with no treatment were also quantified for comparison.
The same growth protocol as outlined above was used to test efficacy of topical products against polymicrobial biofilms. However, inocula concentrations were varied to grow MRSA and P. aeruginosa as polymicrobial biofilms. When the two isolates were inoculated at 1:1 or even 1:1,000 ratio, P. aeruginosa overwhelmed the MRSA isolate. As such, a 1:10,000 ratio was used; MRSA was inoculated at a concentration 10,000 times higher than P. aeruginosa. Each isolate was suspended to a turbidity of 10% using a nephelometer (concentration equated to ~1 x 109 CFU/mL). MRSA was diluted 1:1,000 (~1 x 106 CFU/mL) and P. aeruginosa was diluted 1:10,000,000 (~1 x 102 CFU/mL) using a 10-fold dilution series. Twenty-five μL of each solution were pipetted onto cellulose discs for a total of 50 μL per sample. Polymicrobial biofilm growth was quantified as described above to obtain a baseline of CFU/disc.
Biofilms were observed qualitatively on cellulose discs using confocal laser scanning microscopy (CLSM); a BacLight™ Bacterial Viability kit was used in low light. Treated and untreated cellulose discs (n = 8 from each test) were removed from the agar, stained and fixed in a Nunc™ Lab-Tek™ chamber slide system. Using the chamber slide allowed for staining and fixation all in one chamber so as to preserve the structure of the biofilm prior to evaluation with CLSM. Manufacturer instructions were followed: 3 μL of SYTO 9 3.34 mM and 3 μL of propidium iodide 20 mM per 1 mL of sterile water. A 100 μL volume submerged each cellulose disc in an individual chamber. Chamber slides were covered in aluminum foil to shield from light for 15 min in order for the stain to integrate with the bacterial cells. Stain was decanted from each chamber and samples were washed with sterile PBS. Each sample was fixed with 100 μL of 10% neutral buffered formalin for 30 min, then washed again with 100 μL of PBS. Samples were air dried in the same dark environment and viewed by CLSM.
SEM images were also collected to observe surface morphologies of monomicrobial and polymicrobial growth on cellulose discs. Separate cellulose discs (not used in Live/Dead assays) were fixed for a minimum of 2 h in modified Karnovsky’s fixative (2.5% glutaraldehyde, 2% paraformaldehyde in PBS buffered to pH 7.2–7.4). Samples were dehydrated in 100% ethanol for at least 2 h, air dried, sputter coated with gold, then imaged in a JEOL JSM-6610 SEM.
Collagen coupon assay
Monomicrobial and polymicrobial biofilms of MRSA and P. aeruginosa were also grown on bioabsorbable collagen and exposed to topical therapies. The rationale was two-fold: 1) to perform experiments using a material that would more closely model a physiological substrate, and 2) as will be shown, the Hammond et al. method had important limitations that led to inconsistent results—motivating a secondary analysis [40].
Biofilms were grown on collagen using a modified CDC biofilm reactor (Fig 1A and 1B) as described previously [41]. Rather than using standard coupon rods, blank polypropylene rods were purchased and 4 holes (8 mm diameter) were drilled into the bottom portion. The holes were drilled half way through the rod (Fig 1C). HeliPlug™ collagen coupons were sterilely cut to size (5 mm x 10 mm), and pressed into each bored-out cavity of a rod (Fig 1C). Assembly was performed in a biosafety cabinet to maintain sterility.
Five-hundred mL of brain heart infusion (BHI) broth were added to the reactor after it was assembled. The broth was aseptically inoculated with 105 CFU/mL (adjusted from 0.5 McFarland standard) of MRSA or P. aeruginosa for monomicrobial biofilm growth. The reactor was set on a hot plate at 34°C and a baffle rotation of 130 rpm. Bacteria were grown in batch phase for 24 h, after which a 10% solution of BHI was flowed through the reactor at a rate of 6.94 mL/min using a peristaltic pump (MasterFlex L/S Microbore, Cole Palmer, Vernon Hills, IL) for an additional 24 h (Fig 1A).
The inoculation protocol to grow polymicrobial biofilms on collagen was similar to cellulose; each isolate was suspended to a 0.5 McFarland Standard (~1 x 108 CFU/mL). MRSA was diluted 1:1,000 (~1 x 105 CFU/mL) and P. aeruginosa was diluted 1:10,000,000 (~1 x 10 CFU/mL). However, we determined experimentally that polymicrobial biofilms grew more successfully if the reactor was inoculated multiple times with bacterial solution (one isolate would typically outcompete the other with a single inoculation). The reactor was inoculated five times total: at setup, then 15 min, 45 min, 1 h 45 min, and 3 h 45 min after the initial inoculation. Polymicrobial biofilms were otherwise grown as described for monomicrobial growth.
The efficacy of each topical product was determined against biofilms on collagen by placing 1g of product into a well of a 24-well plate. Collagen coupons were sterilely removed from the reactor and placed individually atop the topical product. Each coupon was then covered with an additional 1g of product, and the lid of the plate replaced. This procedure exposed biofilms on all surfaces of the collagen to a topical product, which was an important difference compared to the Hammond et al. method [40]; in the Hammond et al. method, only the top side of a cellulose disc was exposed to topical treatment, with the underside remaining untreated against the agar surface.
Samples were incubated with their respective topical agents (same as tested against cellulose discs) for 24 h at 37°C. Collagen coupons were removed, rinsed 3x in PBS and placed into 2 mL of PBS. Each sample was vortexed for 1 min, sonicated at 42 kHz for 10 min, vortexed again and plated in triplicate on trypticase soy agar (TSA) using a 10-fold dilution series. Selective agar was made to resolve growth between S. aureus and P. aeruginosa; TSA + triclosan 0.625 μg/mL for P. aeruginosa, and TSA + 7.5% NaCl for MRSA. Selective agar protocols were adapted from previous methods and confirmed with polymicrobial biofilm and planktonic quantification [44–47]. Agar plates were incubated at 37°C and colonies counted at 24 and 48 h. Positive controls of growth (not exposed to antibiotic treatments) were quantified for baseline comparison. All data were collected with n = 8 repeats.
Results
Cytotoxicity
MEM elution tests showed that CZ-01179 had equivalent cytotoxicity outcomes compared to clinical topical products. Mupirocin, gentamicin, retapamulin, and silver sulfadiazine received failing scores; all n = 3 samples of clinically-relevant topicals scored 4 on a scale of 0–4 (score of 3–4 being considered failure and 0–2 considered passing). All n = 3 samples of CZ-01179 also had a score of 4. Neosporin® was the only topical product to receive passing scores of 1, 2, and 1 for the three samples tested.
Cellulose disc assay
The 48-h biofilm growth protocol for cellulose produced well-established, mature monomicrobial and polymicrobial biofilms of both MRSA and P. aeruginosa (Fig 3A–3C). However, quantification outcomes following efficacy analyses were highly variable, in particular with the clinically-relevant products. We established a sub-hypothesis after observing the inconsistent outcomes: we hypothesized that topical treatments failed to reach the biofilms that formed on the underside of the cellulose disc (immediately adjacent to the surface of the agar), and the lack of exposure in that region led to highly variable quantification data.
(A) MRSA biofilms on cellulose (gray substrate) after 48 h of growth. (B) P. aeruginosa biofilms on cellulose after 48 h of growth. The sheet-like growth of P. aeruginosa biofilms covered the substrate completely. (C) Polymicrobial biofilms of MRSA and P. aeruginosa on cellulose after 48 h of growth. P. aeruginosa grew in sheet-like structures with MRSA clusters observed throughout. (D) MRSA biofilm on collagen (gray substrate) after 48 h of growth. (E) P. aeruginosa biofilm on collagen after 48 h of growth. (F) Polymicrobial biofilms on collagen after 48 h of growth. Morphology was similar to that of cellulose with P. aeruginosa growing in sheet-like structures with MRSA clusters interspersed throughout.
To test the sub-hypothesis, biofilms were grown on cellulose following the same growth protocol outlined above. SEM and CLSM imaging was performed to determine: 1) if biofilms formed on the underside of the cellulose fiber network that was in apposition to the agar surface, and 2) if those biofilms on the underside of cellulose discs were still viable following the topical product delivery protocol.
SEM imaging confirmed the presence of biofilms on, within, and between the interstices of the fibers on the underside of cellulose discs (Figs 3 and 4). Live/Dead imaging also indicated that biofilms were viable on all surfaces of cellulose discs, but only surfaces in direct contact with topical agents showed cell death; bacteria on the underside and center of cellulose discs stained green (living), supporting our sub-hypothesis that bacteria on untreated surfaces were still viable and were not exposed to topical product treatments (Figs 5 and 6) [41].
Biofilm growth can be seen within chasms and voids between cellulose fibers.
Cellulose fibers stained green along with live cells. (A) MRSA biofilm on the bottom (non-treated) side of a cellulose disc. The majority of cells stained green (living), suggesting limited activity against the biofilms in that region. (B) MRSA biofilm on the top (treated) side of a cellulose disc. The predominance of red indicated that there was significant antimicrobial activity against biofilms in that region. (C) P. aeruginosa biofilm on the bottom (non-treated) side of a cellulose disc. The predominance of green stain suggested very little, if any activity was present against the biofilms. (D) Biofilm of P. aeruginosa on the top (treated) side of a cellulose disc. The majority of cells stained red, suggesting significant antimicrobial activity, in particular compared to the untreated side. (E) Polymicrobial biofilms on the bottom (non-treated) side of a cellulose disc. The lack of red/yellow indicated little to no antimicrobial activity had occurred. (F) Polymicrobial biofilms on the top (treated) side of a disc. The majority of cells stained red, suggesting that CZ-01179 was effective at eradicating polymicrobial biofilms on the top portions of cellulose discs.
Images were collected with BacLight™ Live/Dead stain. (A) MRSA biofilm treated with gentamicin (0.1%). The predominance of green (living cells) indicated there was limited antimicrobial activity against well-established biofilms. (B) MRSA biofilm treated with CZ-01179 (2%). The antimicrobial gel was highly effective at eradicating well-established biofilms. (C) P. aeruginosa biofilm treated with gentamicin (0.1%). The significant amount of living cells post treatment of gentamicin, demonstrates the limitations of the Hammond et al. method, as P. aeruginosa biofilms grown on collagen treated with gentamicin showed complete eradication (43). (D) P. aeruginosa biofilm treated with CZ-01179 (2%). The antibiofilm agent was able to disrupt the sheet-like structures of the biofilm. (E) Polymicrobial biofilms treated with silver sulfadiazine showed minimal efficacy. (F) Deep and widespread antimicrobial activity was observed within the matrix of the polymicrobial biofilm treated with CZ-01179 (2%) gel.
Despite the limitation of this method, Live/Dead staining provided some useful information on topical efficacy. Confocal imaging and staining indicated that CZ-01179 was highly effective against well-established biofilms that were exposed to the formulated gel, whereas clinical products had limited efficacy (Fig 6). These outcomes provided rationale for performing analysis on collagen coupons.
Collagen coupon assay
Biofilms on collagen grew to maturity (Fig 3), and SEM images indicated more robust biofilm formation compared to cellulose discs. Quantification of positive controls supported this observation with ~1 log10 more CFU/coupon compared to cellulose discs for both isolates.
Quantification data from efficacy testing against biofilms on collagen are reported in Table 1. Outcomes indicated that of the clinical standards of care, gentamicin was most effective against both monomicrobial and polymicrobial biofilms of MRSA and of P. aeruginosa (Fig 7 and Table 1). Gentamicin showed a log10 reduction of 3.56 CFU/collagen in monomicrobial biofilms of MRSA, and against polymicrobial biofilms it was effective against MRSA with a log10 reduction of 5.21 CFU/collagen. Against both monomicrobial and polymicrobial biofilms, gentamicin showed complete eradication of P. aeruginosa, with no detectable growth (Table 1). At all three concentrations (0.5%, 1%, 2%) CZ-01179 reduced all monomicrobial and polymicrobial biofilms to below detectable levels (Fig 7 and Table 1; p = 0.001 or less in all cases compared to controls).
(A) Efficacy of topical agents against MRSA biofilms on collagen. (B) Efficacy of topical agents against P. aeruginosa biofilms on collagen. (C) Efficacy of topical agents against polymicrobial biofilm growth on collagen.
Each collagen coupon received 2g of topical agent applied in the specified concentrations.
Mupirocin and retapamulin were only tested against monomicrobial biofilms of MRSA, as these topicals are FDA approved for staphylococcal species. Mupirocin-treated MRSA biofilms showed a log10 reduction of 3.17 CFU/collagen, indicating similar efficacy as gentamicin against MRSA (Table 1). Retapamulin-treated MRSA biofilms had a log10 reduction of 1.62 CFU/collagen (Table 1).
Data were compared statistically using an independent samples t test with alpha set at 0.05. Tests were run with n = 8 samples/group.
Silver sulfadiazine and Neosporin® are sold as broad spectrum antimicrobial topicals, and were tested against monomicrobial and polymicrobial biofilms of MRSA and P. aeruginosa. Against monomicrobial biofilms of MRSA, collagen treated with silver sulfadiazine had a log10 reduction of 2.97, while collagen treated with Neosporin® had a log10 reduction of 2.46 (Table 1). Similar efficacy was seen when applied to polymicrobial biofilms with MRSA log10 reductions of 4.20 and 2.42 CFU/collagen (Table 1), respectively. Both topicals were less effective against P. aeruginosa in monomicrobial biofilms, with 1.55 and 0.87 log10 reductions, respectively. The same was observed for polymicrobial biofilms; silver sulfadiazine showed 3.74 log10 reductions and Neosporin® showed no reduction (Table 1 and Fig 7). These data are consistent with current literature on the efficacy of Neosporin® and silver sulfadiazine [48].
Discussion
Chronic wounds affect millions of patients annually. They lead to significant morbidity and burden healthcare systems with cost and complexity. When affected by biofilm-related infection, treatment becomes even more challenging. Quality of life can be affected and may lead to amputation. Developing a topical therapy that targets the biofilm phenotype and reduces rates of infection would be an important advancement in clinical care of these wound types. This study assessed the in vitro activity of an antibiofilm gel against MRSA and P. aeruginosa in monomicrobial and polymicrobial biofilms in two separate growth systems as part of a translational process toward this objective.
The efficacy profiles of CZ-01179 gel and clinical products were first collected following a method published by Hammond et al. [40]. Biofilm formation throughout cellulose discs was confirmed by SEM, and CLSM indicated that bacteria on the underside of discs were not eradicated as treatments were only administered to the topside of a disc. Given variable diffusion of topical products this led to variable outcomes, complicating data interpretation. We conclude that this method may not be ideal for assessing efficacy of topical products unless it is modified to control for the lack of exposure to biofilms that are adjacent to the agar surface.
CZ-01179 gels (all three concentrations) had equal efficacy against P. aeruginosa biofilms as gentamicin in the collagen test (Fig 7). CZ-01179 gels were more efficacious at eradicating biofilms in all other cases when compared to the standard of care topicals in the collagen tests (Fig 7). Gentamicin had the greatest log10 reduction against monomicrobial biofilms of P. aeruginosa and polymicrobial biofilms amongst the clinical standards of care. These data were promising, but broader-scale consideration is given in clinical context; aminoglycosides (including gentamicin) are susceptible to both intrinsic and imported resistance mechanisms by P. aeruginosa [10]. These antibacterial resistant strains of P. aeruginosa result in ~6,700 infections per year in the USA alone [1]. These and other limitations motivate and warrant development of additional topical products such as CZ gel that are active against Gram-positive and -negative bacteria provides. Additional product options can help relieve the selective pressures leading to multidrug resistant pathogens.
Mupirocin reduced MRSA CFU by 3.17 log10 units on collagen. Mupirocin is a commonly deployed antibiotic against Gram-positive bacteria such as S. aureus including MRSA, however, there are some limitations to be considered when targeting biofilms. GlaxoSmithKline explains that mupirocin, under the trade name Bactroban®, has limited activity towards anaerobic organisms [49]. A prominent characteristic of mature biofilms is the oxygen gradient that is present at various levels throughout the structure, resulting in a predominantly anaerobic core [35, 50]. This anaerobic core allows for polymicrobial diversity in biofilms, demonstrated by the presence of several strictly anaerobic bacteria within patient chronic wounds, and unaffected by many standard of care topicals [29]. Because S. aureus is a well-known facultative anaerobic organism, it can thrive at all levels along the oxygen gradient in varying metabolic stages. As many standards of care have limited activity on the anaerobic core of a biofilm, the diversity provided by anaerobes and anaerobic phenotypes contributes to prolonged chronicity of biofilm-impaired wounds such as diabetic foot ulcers. Furthermore, like many antibiotics, mupirocin has also been implicated in resistance development by strains of S. aureus [51–53]. Prolonged clinical use of mupirocin is generally not recommended, especially in settings of endemic MRSA colonization [54].
The 2 log10 difference in MRSA reduction between monomicrobial and polymicrobial biofilms can likely be attributed to selective pressures of P. aeruginosa against S. aureus; specifically, excretion of LasA protease and 4-hydroxy-2-heptylquinoline-N-oxide (HQNO), known anti-staphylococcal agents [45–47, 55–58]. It is possible that a longer study would show an initial decrease in MRSA CFUs in polymicrobial biofilms, but then a greater resurgence, as the gene giving resistance to HQNO also provides protection against aminoglycosides, including gentamicin [10, 32, 45, 58]. The HQNO-induced S. aureus strain is easily identified by its small size and slow growth [45, 56, 58]. Due to natural limitations of in vitro studies, the slow growth of this mutated S. aureus strain might conceal the population of surviving cells. Current in vivo studies with a monitoring period post treatment, are being conducted by our laboratory which will provide a more accurate assessment of CZ efficacy against polymicrobial biofilms.
While our quantification data showed that samples treated with CZ-01179 had no detectable surviving bacteria, Live/Dead staining would provide further evidence for its efficacy. Unfortunately, the porous and fibrous nature of the collagen HeliPlug® make it unsuitable for confocal imaging. Finding an additional method for imaging would help us in confirming topical efficacy and biofilm disruption.
Efficacy data resulted in p values that were equivalent across experimental groups when compared against controls (Table 1). While statistical significance is an important parameter for comparing data sets, clinical significance should also be considered. All of the clinically-relevant topical products but gentamicin had a bioburden that remained above 105 CFU. Though challenged, a 105 level is still used as a clinical rule of thumb to indicate a level of bioburden that may cause infection [59]. We seek to develop antibiofilm agents that can effectively reduce biofilm levels to below the 105 level. These in vitro experiments indicated that CZ-01179 achieved that, yet in vivo data is needed to make a more definitive conclusion. We recently published the first in vivo data set of CZ-01179 against biofilms of Acinetobacter baumannii and showed greater than a 105 reduction [60]. In vivo work is currently ongoing with MRSA and P. aeruginosa.
When compared against clinical standards in a MEM elution test, CZ-01179 gel was found to have similar cytotoxicity profiles at therapeutic concentrations. The balance of cytotoxicity is an important consideration, recognizing that infection is also toxic. Promisingly, recent in vivo data indicate that cytotoxicity in topical applications are not seen [60].
Antimicrobial delivery by topical therapy provides the ability to achieve high antimicrobial doses necessary for eradication of a biofilm that would otherwise be unachievable with systemic therapy. CZs constitute a promising class of compounds as they have reduced risk of resistance, formulate well with polymers, are highly soluble and stable (beneficial for hydrogel materials), and are not limited in efficacy by the metabolic state of bacteria in a biofilm. Taken together, data indicated that CZ-01179 is a candidate for advancement toward in vivo testing to assess its ability to treat and/or prevent biofilm-related wound infections caused by monomicrobial or polymicrobial biofilms.
Supporting information
S2 Table. Raw microbiological data for P. aeruginosa (ATCC 27853).
https://doi.org/10.1371/journal.pone.0234832.s002
(PDF)
S3 Table. Raw microbiological data for polymicrobial biofilms composed of MRSA and P. aeruginosa (ATCC 27853).
https://doi.org/10.1371/journal.pone.0234832.s003
(PDF)
References
- 1.
CDC (2013) Antibiotic resistance threats in the United States, 2013.
- 2.
WHO (2016) Global antimicrobial resistance surveillance system (GLASS) report.
- 3. IDSA (2010) The 10 x '20 initiative: pursuing a global commitment to develop 10 new antibacterial drugs by 2020. Clin Infect Dis 50: 1081–1083. pmid:20214473
- 4. Boucher HW, Talbot GH, Bradley JS, Edwards JE, Gilbert D, et al. (2009) Bad Bugs, No Drugs: No ESKAPE! An Update from the Infectious Diseases Society of America. Clin Infect Dis 48: 1–12. pmid:19035777
- 5. Rice LB (2008) Federal funding for the study of antimicrobial resistance in nosocomial pathogens: no ESKAPE. J Infect Dis 197: 1079–1081. pmid:18419525
- 6.
CDC (2011) A public health action plan to combat antimicrobial resistance.
- 7. Huttner A, Harbarth S, Carlet J, Cosgrove S, Goossens H, et al. (2013) Antimicrobial resistance: a global view from the 2013 World Healthcare-Associated Infections Forum. Antimicrob Resist Infect Control 2: 31. pmid:24237856
- 8. Peters NK, Dixon DM, Holland SM, Fauci AS (2008) The research agenda of the National Institute of Allergy and Infectious Diseases for antimicrobial resistance. J Infect Dis 197: 1087–1093. pmid:18419527
- 9. Duarte B, Cabete J, Formiga A, Neves J (2017) Dakin's solution: is there a place for it in the 21st century? Int Wound J 14: 918–920. pmid:28198104
- 10. Bassetti M, Vena A, Croxatto A, Righi E, Guery B (2018) How to manage Pseudomonas aeruginosa infections. Drugs Context 7: 212527. pmid:29872449
- 11. Mangum LC, Franklin NA, Garcia GR, Akers KS, Wenke JC (2018) Rapid degradation and non-selectivity of Dakin's solution prevents effectiveness in contaminated musculoskeletal wound models. Injury 49: 1763–1773. pmid:30104015
- 12.
FDA (2009) Novel drug approvals for 2009.
- 13.
FDA (2010) Novel drug approvals for 2010.
- 14.
FDA (2011) Novel drug approvals for 2011.
- 15.
FDA (2012) Novel drug approvals for 2012.
- 16.
FDA (2013) Novel drug approvals for 2013.
- 17.
FDA (2014) Novel drug approvals for 2014.
- 18.
FDA (2015) Novel drug approvals for 2015. https://www.fda.gov/drugs/new-drugs-fda-cders-new-molecular-entities-and-new-therapeutic-biological-products/novel-drug-approvals-2015.
- 19.
FDA (2016) Novel drug approvals for 2016. https://www.fda.gov/media/102618/download.
- 20.
FDA (2017) Novel drug approvals for 2017. https://www.fda.gov/drugs/new-drugs-fda-cders-new-molecular-entities-and-new-therapeutic-biological-products/novel-drug-approvals-2017.
- 21.
FDA (2018) Novel drug approvals for 2018. https://www.fda.gov/drugs/new-drugs-fda-cders-new-molecular-entities-and-new-therapeutic-biological-products/novel-drug-approvals-2018.
- 22.
FDA (2019) Novel drug approvals for 2019. https://www.fda.gov/drugs/new-drugs-fda-cders-new-molecular-entities-and-new-therapeutic-biological-products/novel-drug-approvals-2019.
- 23. Kirketerp-Møller K, Jensen P, Fazli M, Madsen K, Pedersen J, et al. (2008) Distribution, organization, and ecology of bacteria in chronic wounds. J Clin Microbiol 46: 2717–2722. pmid:18508940
- 24. Aragon-Sanchez FJ, Cabrera-Galvan JJ, Quintana-Marrero Y, Hernandez-Herrero MJ, Lazaro-Martinez JL, et al. (2008) Outcomes of surgical treatment of diabetic foot osteomyelitis: a series of 185 patients with histopathological confirmation of bone involvement. Diabetologia 51: 1962–1970. pmid:18719880
- 25. Wheat J (1985) Diagnostic strategies in osteomyelitis. Am J Med 78: 218–224. pmid:2990206
- 26. Senneville E, Melliez H, Beltrand E, Legout L, Valette M, et al. (2006) Culture of percutaneous bone biopsy specimens for diagnosis of diabetic foot osteomyelitis: concordance with ulcer swab cultures. Clin Infect Dis 42: 57–62. pmid:16323092
- 27. Lesens O, Desbiez F, Vidal M, Robin F, Descamps S, et al. (2011) Culture of per-wound bone specimens: a simplified approach for the medical management of diabetic foot osteomyelitis. Clin Microbiol Infect 17: 285–291. pmid:20156216
- 28. Gjodsbol K, Christensen JJ, Karlsmark T, Jorgensen B, Klein BM, et al. (2006) Multiple bacterial species reside in chronic wounds: a longitudinal study. Int Wound J 3: 225–231. pmid:16984578
- 29. James GA, Swogger E, Wolcott R, Pulcini Ed, Secor P, et al. (2008) Biofilms in chronic wounds. Wound Repair Regen 16: 37–44. pmid:18086294
- 30. Dowd SE, Sun Y, Secor PR, Rhoads DD, Wolcott BM, et al. (2008) Survey of bacterial diversity in chronic wounds using pyrosequencing, DGGE, and full ribosome shotgun sequencing. BMC Microbiol 6: 43.
- 31. Nichols WW, Evans MJ, Slack MP, Walmsley HL (1989) The penetration of antibiotics into aggregates of mucoid and non-mucoid Pseudomonas aeruginosa. J Gen Microbiol 135: 1291–1303. pmid:2516117
- 32. Boles BR, Thoendel M, Singh PK (2004) Self-generated diversity produces "insurance effects" in biofilm communities. Proc Natl Acad Sci U S A 101: 16630–16635. pmid:15546998
- 33. Wolcott RD, Rumbaugh KP, James G, Schultz G, Phillips P, et al. (2010) Biofilm maturity studies indicate sharp debridement opens a time-dependent therapeutic window. J Wound Care 19: 320–328. pmid:20852503
- 34. Mombelli A, Samaranayake LP (2004) Topical and systemic antibiotics in the management of periodontal diseases. Int Dent J 54: 3–14. pmid:15005467
- 35. Costerton JW (1999) Introduction to biofilm. Int J Antimicrob Agents 11: 217–221; discussion 237–219. pmid:10394973
- 36. Chen WY, Rogers AA, Lydon MJ (1992) Characterization of biologic properties of wound fluid collected during early stages of wound healing. J Invest Dermatol 99: 559–564. pmid:1431216
- 37. Field FK, Kerstein MD (1994) Overview of wound healing in a moist environment. Am J Surg 167: 2S–6S. pmid:8109679
- 38. Ashton NN, Allyn G, Porter ST, Haussener TJ, Sebahar PR, et al. (2019) In vitro testing of a first-in-class tri-alkylnorspermidine-biaryl antibiotic in an anti-biofilm silicone coating. Acta Biomater.
- 39. Katsu T, Nakagawa H, Yasuda K (2002) Interaction between polyamines and bacterial outer membranes as investigated with ion-selective electrodes. Antimicrob Agents Chemother 46: 1073–1079. pmid:11897592
- 40. Hammond A, Miller K, Kruczek C, Dertien J, Colmer-Hamood J, et al. (2011) An in vitro biofilm model to examine the effect of antibiotic ointments on biofilms produced by burn wound bacterial isolates. Burns 37: 312–321. pmid:21130579
- 41. Williams DL, Smith SR, Peterson BR, Allyn G, Cadenas L, et al. (2019) Growth substrate may influence biofilm susceptibility to antibiotics. PLoS One 14: e0206774. pmid:30870411
- 42. Williams DL, Haymond BS, Woodbury KL, Beck JP, Moore DE, et al. (2012) Experimental model of biofilm implant-related osteomyelitis to test combination biomaterials using biofilms as initial inocula. J Biomed Mat Res A 100: 1888–1900.
- 43. Williams DL, Haymond BS, Beck JP, Savage PB, Chaudhary V, et al. (2012) In vivo efficacy of a silicone—cationic steroid antimicrobial coating to prevent implant-related infection. Biomaterials 33: 8641–8656. pmid:22940221
- 44. Vandeplassche E, Coenye T, Crabbe A (2017) Developing selective media for quantification of multispecies biofilms following antibiotic treatment. PLoS One 12: e0187540. pmid:29121069
- 45. Hoffman LR, Deziel E, D'Argenio DA, Lepine F, Emerson J, et al. (2006) Selection for Staphylococcus aureus small-colony variants due to growth in the presence of Pseudomonas aeruginosa. Proc Natl Acad Sci U S A 103: 19890–19895. pmid:17172450
- 46. Voggu L, Schlag S, Biswas R, Rosenstein R, Rausch C, et al. (2006) Microevolution of cytochrome bd oxidase in Staphylococci and its implication in resistance to respiratory toxins released by Pseudomonas. J Bacteriol 188: 8079–8086. pmid:17108291
- 47. Biswas L, Biswas R, Schlag M, Bertram R, Gotz F (2009) Small-colony variant selection as a survival strategy for Staphylococcus aureus in the presence of Pseudomonas aeruginosa. Appl Environ Microbiol 75: 6910–6912. pmid:19717621
- 48. Greer N, Foman NA, MacDonald R, Dorrian J, Fitzgerald P, et al. (2013) Advanced wound care therapies for nonhealing diabetic, venous, and arterial ulcers: a systematic review. Ann Intern Med 159: 532–542. pmid:24126647
- 49.
GlaxoSmithKline (2015) Bactroban 2% cream or ointment product information.
- 50. Archer NK, Mazaitis MJ, Costerton JW, Leid JG, Powers ME, et al. (2011) Staphylococcus aureus biofilms: properties, regulation, and roles in human disease. Virulence 2: 445–459. pmid:21921685
- 51. Maple PA, Hamilton-Miller JM, Brumfitt W (1989) World-wide antibiotic resistance in methicillin-resistant Staphylococcus aureus. Lancet 1: 537–540. pmid:2564067
- 52. Cookson BD (1998) The emergence of mupirocin resistance: a challenge to infection control and antibiotic prescribing practice. J Antimicrob Chemother 41: 11–18. pmid:9511032
- 53. Patel JB, Gorwitz RJ, Jernigan JA (2009) Mupirocin resistance. Clin Infect Dis 49: 935–941. pmid:19673644
- 54. Vasquez JE, Walker ES, Franzus BW, Overbay BK, Reagan DR, et al. (2000) The epidemiology of mupirocin resistance among methicillin-resistant Staphylococcus aureus at a Veterans' Affairs hospital. Infect Control Hosp Epidemiol 21: 459–464. pmid:10926396
- 55. Machan ZA, Taylor GW, Pitt TL, Cole PJ, Wilson R (1992) 2-Heptyl-4-hydroxyquinoline N-oxide, an antistaphylococcal agent produced by Pseudomonas aeruginosa. J Antimicrob Chemother 30: 615–623. pmid:1493979
- 56. Mansito TB, Falcon MA, Moreno J, Carnicero A, Gutierrez-Navarro AM (1987) Effects of staphylolytic enzymes from Pseudomonas aeruginosa on the growth and ultrastructure of Staphylococcus aureus. Microbios 49: 55–64. pmid:3104732
- 57. Barequet IS, Ben Simon GJ, Safrin M, Ohman DE, Kessler E (2004) Pseudomonas aeruginosa LasA protease in treatment of experimental staphylococcal keratitis. Antimicrob Agents Chemother 48: 1681–1687. pmid:15105121
- 58. DeLeon S, Clinton A, Fowler H, Everett J, Horswill AR, et al. (2014) Synergistic interactions of Pseudomonas aeruginosa and Staphylococcus aureus in an in vitro wound model. Infect Immun 82: 4718–4728. pmid:25156721
- 59. Bowler PG (2003) The 105 bacterial growth guideline: reassessing its clinical relevance in wound healing. Ostomy Wound Manage 49: 44–53.
- 60. Williams DL, Kawaguchi B, Taylor NB, Allyn G, Badham MA, et al. (2020) In vivo efficacy of a unique first-in-class antibiofilm antibiotic for biofilm-related wound infections caused by Acinetobacter baumannii. Biofilm 2: Epub ahead of print.