Adaptive antimicrobial nanocarriers for the control of infectious biofilms
Liu, Yong
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CHAPTER 2
Surface-Adaptive, Antimicrobially Loaded,
Micellar Nanocarriers with Enhanced Penetration
and Killing Efficiency in Staphylococcal Biofilms
Liu, Y.; Busscher, H. J.; Zhao, B.; Li, Y.; Zhang, Z.; van der Mei, H. C.;
Ren, Y.; Shi, L., ACS Nano 2016, 10 (4), 4779–4789.
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ABSTRACT
Biofilms cause persistent bacterial infections and are extremely recalcitrant to antimicrobials, due in part to reduced penetration of antimicrobials into biofilms that allows bacteria residing in the depth of a biofilm
to survive antimicrobial treatment. Here, we describe the preparation of surface-adaptive, Triclosan-loaded micellar nanocarriers showing (1) enhanced biofilm penetration and accumulation, (2) electrostatic targeting at acidic pH toward negatively charged bacterial cell surfaces in a biofilm, and (3) antimicrobial release due to degradation of the micelle core by bacterial lipases. First, it was established that mixed-shell-polymeric-micelles (MSPM) consisting of a hydrophilic poly(ethylene glycol) (PEG)-shell and pH-responsive poly(β-amino ester) become positively charged at pH 5.0, while being negatively charged at physiological pH. This is opposite to single-shell-polymeric-micelles (SSPM) possessing only a PEG-shell and remaining negatively charged at pH 5.0. The stealth properties of the PEG-shell combined with its surface adaptive charge allow MSPMs to penetrate and accumulate in staphylococcal biofilms, as demonstrated for fluorescent Nile red loaded micelles using confocal-laser-scanning-microscopy. SSPMs, not adapting a positive charge at pH 5.0, could not be demonstrated to penetrate and accumulate in a biofilm. Once micellar nanocarriers are bound to a staphylococcal cell surface, bacterial enzymes degrade the MSPM core to release its antimicrobial content and kill bacteria over the depth of a biofilm. This constitutes a highly effective pathway to control blood-accessible staphylococcal biofilms using antimicrobials, bypassing biofilm recalcitrance to antimicrobial penetration.
KEYWORDS
:
biofilm, antimicrobials, micelles, Triclosan, staphylococci, electrostatic interactions,2
INTRODUCTION
Over 60% of all human infections treated by physicians are due to biofilms,1 examples being oral biofilms and biofilms involved in a variety of pathological conditions like for instance osteomyelitis, chronic otitis media, the infected diabetic foot, chronic bacterial prostatitis or in biomaterialassociated infections.2 In a biofilm,3 microorganisms produce extracellular polymeric substances (EPS) that embed biofilm inhabitants in a protective matrix.2 The biofilm matrix not only provides a microenvironment for microbial growth, catalysis, and communication, but also protects its inhabitants against environmental challenges, such as UV exposure, acids,4 or antimicrobials.5 Biofilms are extremely recalcitrant to antimicrobials and ever since Van Leeuwenhoek in 1684 reported that “the vinegar with which I washt my teeth, kill’d only those animals which were
on the outside of the scurf, but did not pass thro the whole substance of it”,6 little progress has been made in making biofilms more susceptible to antimicrobial treatment.
Cationic tobramycin, for instance, was unable to penetrate Pseudomonas aeruginosa biofilms, even of strains not producing an EPS matrix and was sequestered at the biofilm periphery by electrostatic attraction. Alternatively, a zwitterionic antibiotic, ciprofloxacin, could penetrate biofilms of non-EPS producing P.
aeruginosa strains.7
Most pathogenic strains, however, produce an EPS matrix that hampers penetration of antibiotics into a biofilm. The use of engineered nanoparticles may constitute an advanced and superior way to overcome biofilm recalcitrance to antimicrobial treatment.8−10
Many carbon-based nanomaterials such as carbon nanotubes11
and graphene nanosheets,12−14
as well as some metallic nanocomposites including, but not limited to, silver,15,16
gold,17−19
and magnetic nanoparticles,20,21
possess promising antimicrobial efficacy. Nanoparticles have also been suggested for use as carriers for antimicrobial administration,22,23
especially of poor water-soluble ones,24
such as, for instance, Triclosan.25
Application of polymeric micelles as antimicrobial nanocarriers can tremendously improve the pharmacokinetics and biodistribution of antimicrobials.23,26,27 Micellar nanocarriers excel in their biodegradability and biocompatibility, versatility of synthesis and functional modifications for targeted and stimuliresponsive release of therapeutics.26 Cationic micellar nanocarriers can selectively target microbial membranes over mammalian cell membranes and have been shown to inhibit growth of planktonic, methicillin-resistant Staphylococcus aureus and fungi through strong electrostatic interactions with the negatively charged microbial cell surfaces.28,29 pH-activated nanocarriers can provide not only targeted but also on-demand drug release,30 and local pH changes and presence of bacterial enzymes may trigger antimicrobial release.31,32 However, the antibacterial efficacy of targeted and stimuli-responsive nanocarriers has been mostly evaluated against planktonic bacteria and not against bacteria in their biofilm mode of growth. Moreover, positively charged nanocarriers are short-lived in the blood circulation.33 Whether this should be considered as a disadvantage depends on the goals for their application. Micellar nanocarriers made of cationic poly(dimethylaminoethyl methacrylate) (p(DMAEMA)) coronas and hydrophobic p-(DMAEMA-co-butyl methacrylate-co-propylacrylic acid) cores possess a positive zeta potential ranging from +5 mV at pH 10.2 to +25 mV at pH 3.4 which allows them to target themselves through electrostatic attraction to negatively charged oral surfaces over an extremely wide pH range.27 Adhesion of these cationic pH-activated micelles to negatively charged oral surfaces allows them to remain present in the oral cavity for prolonged periods of time for sustained delivery of farnesol (a quorum quenching molecule) to disrupt oral biofilms, which is an absolute requirement for any therapeutic compound in the oral cavity to prevent rapid wash-out and exert the effects for which they aimed.34 However, a positively charged corona over a wide pH range may bind to almost any surface in the human body, that are nearly all negatively charged, inhibiting the use of these micelles as a blood-born carrier for any drug.
Poly(ethylene glycol) (PEG) has been amply applied to provide stealth properties to materials, making them “invisible” to microorganisms and cells, as well as resistant to protein adsorption.35
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to provide stealth properties to micellar nanocarriers. Mixed-shell polymeric micelles (MSPMs) possess a shell composed of hydrophilic PEG and pH-responsive poly(β-amino ester) (PAE).36,37
Under physiological pH conditions (pH 7.4), the PAE block is hydrophobic and collapsed on the micelle core and the micelles only expose a PEG shell, making them biologically invisible and facilitating long-time circulation in blood and penetration into tumors.36
When in a more acidic, often pathological condition, the PAE block will be protonated and become hydrophilic exposing its positive charge, yielding enhanced accumulation through electrostatic interactions nearby cancerous sites that are generally slightly acidic and comprise negatively charged surfaces.
Here, we hypothesize that MSPMs composed of PEG and pH responsive PAE, will
(1) fully penetrate and accumulate in bacterial biofilms owing to their stealth properties and negative charge under physiological pH conditions
(2) become positively charged under the low pH conditions in the close vicinity of bacterial cell surfaces and there with target themselves to negatively charged bacterial cell surfaces to retain themselves in the biofilm and prevent wash-out and
(3) once interacting with the bacterial cell surface, become hydrolyzed by bacterial lipases to cause release of an encapsulated drug.
Once confirmed, our hypotheses suggest that surface-adaptive, antimicrobially loaded MSPMs will kill bacteria in their biofilm mode of growth better than nonencapsulated antimicrobials or antimicrobials encapsulated by single-shell polymeric micelles (SSPMs, comprised of poly(ethylene glycol)-block-poly(ε- caprolactone) PEG-b-PCL alone). SSPMs likely are biologically equally invisible as MSPMs, but will not surface-adapt to a pathological low pH environment to enable interaction with bacterial cell surfaces to accumulate themselves in a biofilm in bactericidal concentrations.
In this manuscript, we aim to verify the above hypotheses and show that MSPMs loaded with an antimicrobial more effectively kill bacteria in deeper layers of a biofilm than antimicrobially loaded SSPMs or antimicrobials without encapsulation. Zeta potentials of both types of micelles will be compared as a function of pH using particulate microelectrophoresis and binding of both types of micelles to planktonic, green-fluorescent S. aureus determined. Improved penetration and accumulation of MSPMs into green-green-fluorescent
S. aureus biofilms will be demonstrated with respect to SSMPs using fluorescent Nile red loaded micelles.
Micellar nanocarriers are subsequently loaded with a hydrophobic antimicrobial, Triclosan,25
possessing a poor watersolubility. Minimum inhibitory (MIC) and minimum bactericidal (MBC) concentrations of Triclosan in the absence and presence of encapsulation by MSPMs or SSPMs will be compared for planktonic staphylococci, while efficacy against fluorescent S. aureus ATCC12600GFP
and bioluminescent S. aureus Xen36 biofilms is evaluated using a bio-optical imaging system for high-throughput comparison. Results confirm that our Triclosan-loaded, surface-adaptive MSPMs are more effective in killing staphylococci deep into a biofilm as compared with nonencapsulated Triclosan or Triclosan encapsulated in SSPMs.
RESULTS
Micelle Characterization and Triclosan-Release. MSPMs and SSPMs were prepared with and without Triclosan-loading. First, we measured the effects of loading the micelles with Triclosan on their diameters using dynamic light scattering (Figure 1A). Figure 1A shows relatively narrow size distributions for all micellar nanocarriers (polydispersity < 0.15) and a spherical morphology is confirmed in transmission electron micrographs. MSPMs and SSPMs in the absence of Triclosan-loading had a diameter of around 80−100 nm, regardless of pH (Figure 1B). While Triclosan-loading had no effect on the diameter of SSPMs, the diameter of MSPMs increased significantly (p < 0.05; Student’s t test), also regardless of pH,
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upon hydrophobic drug loading to 160 nm, possibly due to a higher drug loading efficiency of the MSPMs compared to SSPMs (78 ± 2% and 49 ± 1%, respectively, see Table S1). The absence of a pH-dependence of the micelle diameters indicates that both unloaded and Triclosan-loaded nanocarriers remain intact during environmental pH changes. 50 100 150 200 0 50 100 Micelle diameter (nm) R e la tiv e in te n s ity (% ) 0 20 40 0 200 400 Time (h) Cu m u la ti ve re lease ( g) pH 7.4 pH 5.0 pH 7.0 pH 7.4 0 50 100 150 200 M icel le D iam eter (n m ) 0 20 40 0 200 400 Time (h) Cu m u la ti ve re lease ( g) pH 5.0 pH 5.0 pH 7.0 pH 7.4 -10 0 10 20 M icel le D iam et er (n m ) SSPM SSPM+T MSPM MSPM+T SSPM+T+Lipase MSPM+T+Lipase
A
B
C
D
E
Figure 1. Micelle characterization and Triclosan (T)-release at different pH values. (A) Micelle
diameters of MSPMs and SSPMs at pH 5.0 with and without Triclosan-loading, measured using dynamic light scattering. (B) Micelle diameters of MSPMs and SSPMs at different pH values with and without Triclosan-loading. Error bars denote the width of the intensity peaks in dynamic light scattering. (C) Zeta potentials in 10 mM phosphate buffer of MSPMs and SSPMs at different pH values with and without Triclosan-loading. Error bars denote the standard deviations over triplicate measurements with separately prepared micelle suspensions. (D) Cumulative release of Triclosan from MSPMs and SSPMs at pH 7.4 in 10 mM phosphate buffer in absence or presence of lipase. Experiments were conducted in a total suspension volume of 2 mL at a micelle concentration of 1 mg mL−1
at 37 °C. Error bars denote the standard deviations over triplicate measurements with separately prepared micelle suspensions. (E) As in panel D, now at pH 5.0.
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Zeta potentials of SSPMs are negative at all pH values both in absence and presence of Triclosan-loading (see Figure 1C). MSPMs have less negative zeta potentials than SSPMs under all conditions, but become positively charged at pH 5.0, regardless of Triclosan-loading, illustrating their surface-adaptive nature.
Triclosan-release from the micellar nanocarriers was assessed using UV−vis spectrometry after calibration for Triclosan. Cumulative release of Triclosan was significantly higher (p < 0.05; Student’s t test) at pH 7.4 than at pH 5.0 both for SSPMs and MSPMs (compare Figure 1, panels D and E). For both SSPMs and MSPMs, release of Triclosan in absence of lipase corresponded with approximately 15% and 60% of their total loading at pH 5.0 and 7.4, respectively, as can be calculated using Table S1. In the presence of lipase, however, both types of micelles demonstrated a clear burst release that was significantly higher than in absence of lipase (p < 0.01; Student’s t test), with both types of micelles releasing similar amounts of Triclosan over a time period of at least 50 h. Release corresponded with approximately 80−90% of the total loading of either type of micelles at pH 5.0 and pH 7.4, respectively (compare Figure 1D,E with Table S1).
-20 0 20 40 Ze ta p o ten ti a l (m V ) S. aureus ATCC12600GFP S. aureus Xen36 SSPM+Nile red MSPM+Nile red 5 6 7 8 0.0 0.2 0.4 0.6 0.8 pH MSPM+Nile red SSPM+Nile red In te ra c ti o n (% )
A
B
C
D
E
50mFigure 2. pH-dependent electrostatic interactions between Nile red loaded micelles and
staphylococci. (A−C) Fluorescence micrographs of S. aureus ATCC12600GFP (A) and Nile red loaded MSPMs (B) after interaction in suspension at pH 5.0, together with a merged-channel overlayer image, showing interacting micelles and staphylococci as yellow pixels (C). (D) Zeta potentials of Nile red loaded SSPM and MSPM micelles in 10 mM phosphate buffer as a function of pH as well as of both staphylococcal strains involved in the study. Error bars denote the standard deviations over triplicate measurements with separately prepared micelle suspensions or different bacterial cultures. (E) Interaction between Nile red loaded micelles and planktonic
S. aureus ATCC12600GFP as a function of pH. Interaction percentage is defined as the percentage area of yellow appearing pixels in merged-channel images of green-fluorescent staphylococcal and red-fluorescent micelle images, setting the total area of green-fluorescent pixels in panel A at 100%. Error bars denote the standard deviations over triplicate measurements with separately prepared micelle suspensions and different bacterial cultures.
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Penetration of Nile Red Loaded Micelles into Staphylococcal Biofilms and Their pH-Dependent Bacterial Targeting. To demonstrate the pH-dependent targeting of MSPMs, we have loaded both types of micelles with a hydrophobic red-fluorescent dye, Nile red. First, we studied the pH-dependent interaction of the micelles with planktonic S. aureus ATCC12600GFP
in 10 mM phosphate buffer. Using fluorescence microscopy, images of green-fluorescent staphylococci in a suspension droplet on a glass slide (see Figure 2A for an example image) were overlaid with corresponding red fluorescent images of Nile red loaded MSPMs (Figure 2B) to yield yellow-fluorescent images of interacting staphylococci and micelles (Figure 2C). Next, the electrostatic nature of the pH dependent targeting of MSPMs toward staphylococci was demonstrated by relating the zeta potentials of Nile red loaded micellar nanocarriers and both staphylococcal strains involved in this study (Figure 2D) with the pH-dependence of their interaction. As can be seen in Figure 2D, both staphylococcal strains involved in this study remain negatively charged over the pH range of 5.0−7.4, while the zeta potentials of SSPMs are slightly negatively charged over this pH trajectory. The zeta potentials of MSPMs, however, become increasingly more positive when the pH decreases to below pH 6.5. By comparison with Figure 2E, it can be seen that the interaction between MSPMs and planktonic S. aureus ATCC12600GFP
concurrently increases with decreasing pH. Note that the staphylococcal interaction with SSPMs remains low. This demonstrates that the pH-stimulated change in zeta potential of MSPMs targets the micelles toward negatively charged bacterial cell surfaces. Importantly, also Nile red loaded micelles retained their integrity with similar hydrodynamic diameters upon lowering the pH.
Next, to demonstrate stealth penetration and accumulation in staphylococcal biofilms, biofilms of green-fluorescent S. aureus ATCC12600GFP
were exposed to suspensions of Nile red loaded micelles and imaged using confocal laser scanning microscopy (CLSM) (Figure 3A−D). SSPMs had no affinity for the staphylococcal biofilms and penetration and accumulation could not be demonstrated at pH 7.4 (Figure 3A) nor at pH 5.0 (Figure 3C), likely due to lack of interaction with the staphylococci in the biofilm. MSPMs, however, showed a completely different pattern of penetration and accumulation in staphylococcal biofilms (Figure 3B,D). Whereas demonstrable penetration and accumulation of red-fluorescent MSPMs at pH 7.4
Figure 3. Penetration and pH-dependent bacterial targeting of Nile red loaded micelles in a
staphylococcal biofilm. (A and C) CLSM micrographs demonstrating the absence of penetration and accumulation of Nile red loaded SSPMs into a biofilm of S. aureus ATCC12600GFP
at pH 7.4 (A) and pH 5.0 (C). (B and D) CLSM micrographs demonstrating penetration and accumulation of Nile red loaded MSPMs into a biofilm of S. aureus ATCC12600GFP
at pH 5.0 (D), while being absent at pH 7.4 (B).
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K KK K
K K
Figure 4. Reduction in metabolic activity and killing of staphylococci in their biofilm mode of
growth by Triclosan in solution and Triclosan loaded micellar nanocarriers using bio-optical imaging. (A) Bioluminescence images of S. aureus Xen36 biofilms after exposure for different times at pH 7.4 to different concentrations of Triclosan encapsulated in MSPMs. Note that reduction in bioluminescence is not necessarily associated with cell death, but moreover with a decreased metabolic activity.40 (B) Fluorescence images of S. aureus ATCC12600GFP biofilms after exposure for different times at pH 7.4 to different concentrations of Triclosan encapsulated in MSPMs. Note that green-fluorescent bacteria stay fluorescent even in their starvation phase until cell death occurs.41 (C, E, and G) Relative bioluminescence intensity of S. aureus Xen36 biofilms after 1 h (C), 3 h (E), and 5 h (G) as a function of the concentration of Triclosan in solution or in Triclosan-loaded micelles at pH 7.4. Bioluminescence intensity directly after harvesting in buffer was set at 100%. All data are expressed as means ± SD over triplicate experiments with separately prepared micelles and different staphylococcal cultures. (D, F, and H) Relative fluorescence intensity ofS. aureus ATCC12600GFP biofilms after 1 h (D), 3 h (F), and 5 h (H) as a function of the concentration of Triclosan in solution or in Triclosan-loaded micelles at pH 7.4. Fluorescence intensity directly after harvesting in buffer was set at 100%. All data are expressed as means ± SD over triplicate experiments with separately prepared micelles and different staphylococcal cultures.
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was completely absent (Figure 3B), Nile red loaded MSPMs penetrated well into the staphylococcal biofilms at pH 5.0, on certain locations even up to the bottom of the biofilm (Figure 3D). In line with the interaction between Nile red loaded MSPMs and planktonic staphylococci (Figure 2E), we also observe strong targeting of MSPMs at pH 5.0 toward staphylococci in their biofilm mode of growth, that is absent at pH 7.4 as well as for SSPMs at both pH values. Evidently, the targeted interaction of MSPMs with bacterial cell surfaces allows the accumulation of demonstrable concentrations of Nile red loaded MSPMs in a low pH environment, opposite to SSPMs.Staphylococcal Killing by Triclosan and Triclosan-Loaded Micelles in Planktonic and Biofilm Modes of Growth. To evaluate the killing efficacy of Triclosan in solution and Triclosan-loaded micellar nanocarriers, MIC and MBC for planktonic bioluminescent S. aureus Xen36 and fluorescent S. aureus ATCC12600GFP
were determined, as listed in Table 1. Both strains appear nearly similar with respect to their susceptibility for Triclosan in solution as well as for encapsulated Triclosan. In a planktonic state at pH 7.4, staphylococci appear slightly more susceptible to Triclosan in solution than to Triclosan when encapsulated in micellar nanocarriers, although differences were very small and only involved one or two dilution steps. Staphylococcal killing (MBC) required 2- to 8-fold higher amounts of Triclosan than growth inhibition (MIC). Next, the reduction in metabolic activity and killing efficacy of Triclosan and Triclosan encapsulated in both types of micelles was compared against staphylococci in their biofilm mode of growth. To do these experiments as a function of concentration and time, we turned to a new technique, bio-optical imaging.38,39
The use of bio-optical imaging enabled the quantitative analysis of numerous bioluminescent (Figure 4A, indicative of reduced metabolic activity) or fluorescent (Figure 4B, indicative of cell death) biofilms after exposure to Triclosan under different conditions in one single experiment. Taking the relative bioluminescence or fluorescence with respect to the one immediately after harvesting in buffer as 100%, we quantitatively determined the influence of Triclosan concentration and exposure time (Figure 4) on the metabolic activity and viability of S. aureus Xen36 and S. aureus ATCC12600GFP
biofilms, respectively. The advantage with respect to a reduction in metabolic activity of staphylococci in a biofilm mode of growth of Triclosan-loaded MSPMs becomes evident for the bioluminescent strain already at a low encapsulated Triclosan concentration of around 5 μg mL−1
at an exposure time of 3 h (compare Figure 4, panels C, E, and G). This demonstrates that targeting and enzymatic degradation of the MSPMs requires time in order to become effective as compared with effects of Triclosan in solution that shows most, but limited, effects at an exposure time of 1 h. Staphylococcal killing of Triclosan-loaded MSPMs becomes evident for the fluorescent strain (Figure 4D,F,H) at higher Triclosan concentrations than for which reduced metabolism was observed and occurred above 40 μg mL−1
at an exposure time of 3 h or longer (Figure 4D,F,H).
DISCUSSION
The lack of susceptibility of bacteria in their biofilm mode of growth has plagued infection control for many ages. In this paper, we present a surface-adaptive, antimicrobially loaded, micellar nanocarrier system that penetrates and accumulates in a staphylococcal biofilm to kill bacteria over the depth of the biofilm. Whereas antimicrobials in solution only penetrate and kill bacteria on the outside of a biofilm (see Figure 5A), we here show that micellar nanocarriers equipped with a PEG shell and surface-adaptive charge properties fully penetrate and accumulate in a biofilm (Figure 5C). Essential for penetration are both the stealth PEG-shell and the negative micellar charge upon initial penetration, as cationic antimicrobials7
and pH-activated, cationic micellar nanocarriers27
can interact with negatively charged surfaces, including bacterial cell surfaces28,42
at the periphery of a biofilm. Due to the lack of surface-adaptive charge properties, however, SSPMs that remain negatively charged also after initial penetration into a biofilm are not able to demonstrably accumulate in a biofilm in antibacterially effective concentrations, neither do they target to bacterial cell surfaces and effectively degrade while diffusing through biofilm channels to release their antimicrobial content (Figure 5B).
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Stealth MSPMs on the other hand (Figure 5C) are retained in a biofilm since once inside, they encounter low pH conditions that cause protonation of the hydrophobic PAE block on the micelle core, therewith making them hydrophilic and positively charged (Figure 5D). This has a dual effect: first, they adsorb to the negatively charged, bacterial cell surfaces,28,42
while second, in the close vicinity of bacteria they are more amenable to enzymatic degradation of the micelle core and release their antimicrobial content.43
Therefore, we have developed an antimicrobial nanocarrier system that, owing to their stealth properties and surface-adaptive features, bypasses biofilm recalcitrance to antimicrobial penetration, therewith constituting a major progress in combatting EPS producing bacteria in their biofilm mode of growth. Although proteins in EPS as produced by our S. aureus strain used can also become positively charged during a pH drop in a biofilm due to fermentation,44
this did not impede penetration of our MSPMs into the biofilm nor hampered their killing efficacy.
This study makes use of Triclosan as an antimicrobial and S. aureus as a target strain. Triclosan is sufficiently hydrophobic for encapsulation in micelles, but might have been replaced by any other hydrophobic
Figure 5. Schematic presentation (not drawn to scale) of our hypotheses and findings that
antimicrobially loaded MSPMs composed of PEG and pH-responsive PAE kill bacteria better through the depth of a biofilm than nonencapsulated antimicrobials or antimicrobials encapsulated by SSPMs. (A) Nonencapsulated antimicrobials penetrate to a limited degree into a biofilm and kill only bacteria on the outside of the biofilm. Penetration is limited by adsorption to bacterial cell surfaces and matrix components. (B) Antimicrobials encapsulated in a SSPM nanocarrier with stealth properties will show better penetration into a biofilm than nonencapsulated ones and thus kill bacteria in deeper layers of the biofilm, provided sufficient antimicrobial release. Due to the stealth properties of the SSPM nanocarriers, there will be no targeting to bacterial cell surfaces and, as a consequence, little enzymatic degradation of micelles and antimicrobial release. (C) Antimicrobials encapsulated in a MSPM nanocarrier with stealth properties will show full penetration in a biofilm due to their stealth properties and become positively charged in the low pH vicinity of bacteria to target themselves to the bacterial cell surface and expose their micelle core (see panel D). The micelle core subsequently becomes hydrolyzed by bacterial lipases to release its antimicrobial content. (D) Summary of the surface-adaptability of MSPMs under the influence of pH changes and lipase degradation.
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antimicrobial. On the other hand, Triclosan is a very suitable chemical for biofilm control, as it is used in personal care and hygiene products, such as deodorants and soaps and in oral health products to decrease oral biofilm formation and treat inflammatory lesions.45
Also, Triclosan coated polyglactin 910 sutures have been introduced and demonstrated to be effective in preventing suture-colonization by wild-type and methicillin-resistant S. aureus and Staphylococcus epidermidis.46
Moreover, there is a lack of clinical evidence to suggest that the use of Triclosan has led to the propagation of antibiotic resistant staphylococci, and toxicity to mammals is relatively low.47
Planktonically, the MIC and MBC of both our staphylococcal strains used are very similar (Table 1), while also in their charge properties there are no important differences (Figure 2D). This enables a direct comparison of the beneficial effects of our Triclosan-loaded MSPMs on staphylococcal bioluminescence and fluorescence when in a biofilm mode of growth for the two strains (Figure 4). We see clear reductions in bioluminescence indicative of reduced metabolic activity40
at low encapsulated Triclosan concentrations of around 5 μg mL−1
as compared with Triclosan in solution or encapsulated in SSPMs. Fluorescence reductions indicative of staphylococcal killing37
occur at higher encapsulated Triclosan concentrations of around 40 μg mL−1
. These differences are in line with the higher MBC values of Triclosan in solution or encapsulated against planktonic staphylococci as compared with their MIC values, presented in Table 1.
Some papers48
report cytotoxic limits of Triclosan in the range of several tens of micrograms per milliliter (μg mL−1
), while other papers47,49
negate a cytotoxic limit. Triclosan concentrations as high as 40 μg mL−1
required for staphylococcal killing can easily be reached in in vitro release studies (see, e.g., Figure 1D,E) carried out in fluid volumes that are small with respect to the volume of blood in the human circulation in which micelles would have to be injected in a clinical application and seek their way to a blood-accessible infection site. This means that clinically, even when the total Triclosan content of our micelles would be released within 1 h, its concentration in blood would still be below any cytotoxic limit of Triclosan reported in the literature. An important feature of our surface-adaptive micellar nanocarriers, however, is that they target themselves to bacterial cell surfaces to become degraded by bacterial lipases, stimulating Triclosan release from the nanocarriers inside the biofilm. This turns our Triclosan-loaded MSPMs into a highly local drugdelivery device that can yield high concentrations far above MBC in a biofilm while leaving systemic Triclosan concentrations well below the toxic limit. Thus, for our self-targeting local Triclosan delivering micellar nanocarriers, “local” refers to “inside the biofilm”, whereas for all other clinically applied local drugdelivery devices,50
including antibiotic-loaded beads and bone cements, “local” release refers to “in the neighborhood” of an infection, but not to the inside of an infectious biofilm.
S. aureus Xen36 S. aureus ATCC12600GFP
MIC (μg mL−1) MBC (μg mL−1) MIC (μg mL−1) MBC (μg mL−1) Triclosan 1.25 10 1.25 10 SSPM+T 5 10 2.5 20 MSPM+T 2.5 10 2.5 10
Table 1. Minimal inhibitory and minimal bactericidal concentrations (MIC and MBC,
respectively) of boluminescent S. aureus Xen36 and green-fluorescent S. aureus ATCC12600GFP
against Triclosan in solution (10 mM phosphate buffer at pH 7.4) and encapsulated in SSPMs or MSPMsa
a
Note that in case of MIC and MBC values against Triclosan-loaded micelles (+T), encapsulated Triclosan concentrations are derived from the micelle concentration in suspension and their drug loading content (see Table S1).
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CONCLUSIONS
Due to the rise in antimicrobial-resistant bacterial strains, modern medicine is in dire need of improved strategies to control bacterial infections associated with biofilms. We have developed surface-adaptive, pH-responsive mixed-shell polymeric micelles as nanocarriers for hydrophobic antimicrobials, such as Triclosan. When loaded with Triclosan, these nanocarriers penetrate S. aureus biofilms owing to their stealth properties at physiological pH, adapt a positive charge under low, pathological pH conditions in the close vicinity of bacterial cell surfaces to accumulate in a biofilm and target themselves to negatively charged bacterial cell surfaces where they become hydrolyzed by bacterial lipases to cause release of an encapsulated drug. In our study, we used a Gram-positive bacterial strain, but the mechanisms of action of our MSPMs are nonspecific, based on electrostatic repulsion and attraction, and can be expected to also work in biofilms of other Gram-positive and Gram-negative bacteria with a similar pH-dependence of their zeta potentials. Therewith, this approach constitutes a major progress toward killing bacteria in their biofilm mode of growth, bypassing biofilm recalcitrance to antimicrobial penetration.
MATERIALS AND METHODS
Polymer Synthesis. PEG-b-PCL and PCL-b-PAE were synthesized as previously reported (Scheme S1).37 Briefly, PEG-b-PCL was synthesized through ring-opening polymerization of monomeric ε-caprolactone with PEG−OH as an initiator and Sn(Oct)2 as a catalyst in refluxed toluene (Scheme S1A). Next, the solvent was
removed and the crude product was dissolved in dichloromethane, followed by precipitation into an excess amount of diethyl ether and the precipitate was dried in vacuum. For the synthesis of PCL-b-PAE, PCL− OH was first synthesized by ring-opening polymerization of ε-caprolactone using BOC-NH-C2H4OH as an
initiator, similar to the synthesis of PEG-b- PCL, described above. Subsequently, PCL−OH was turned into PCL monoacrylate by reacting with acryloyl chloride at low temperature in the presence of triethylamine as the acid-binding agent. Finally, PCL monoacrylate was added to hexane-1,6-dioldiacrylate (HDD), 4,4′- trimethylene dipiperidine (TDP) in chloroform to yield PCL-b-PAE (Scheme S1B).37 All polymers were subjected to characterization by nuclear magnetic resonance and gel permeation chromatography to confirm their molecular structure and degree of polymerization degree. Stock solutions of PEG5k-b-PCL10k and PCL10k
-b-PAE11k in tetrahydrofuran with a concentration of 5 mg mL −1
were used for the preparation of micelles. Preparation of Triclosan-Loaded, Mixed-Shell and Single-Shell Polymeric Micelles. For the preparation of MSPM suspensions, equal volumes (1 mL) of both polymer solutions in tetrahydrofuran were mixed. The resulting solution was added dropwise into 7 mL of an acetate buffer (pH 4.5, 100 mM) at a rate of 1 droplet (20 μL) per 20 s under magnetic stirring for 4 h to form a micelle suspension. The micelle suspension was dialyzed in a dialysis bag with molecular weight cut off of 6−8 kDa against ultrapure water for 24 h to remove tetrahydrofuran. The dialyzed micelle suspension was diluted to a volume of 10 mL with a final concentration of 1mg mL−1
and stored in a refrigerator at 4 °C. For the preparation of SSPM suspensions, only PEG5k-b-PCL10k in tetrahydrofuran was used. Triclosan-loaded micelles were essentially prepared in the
same way as described above, the only difference being that the polymer solutions in tetrahydrofuran were first mixed with a Triclosan solution in the same solvent (1 mg mL−1
). In the resulting polymer/Triclosan solution, the mass ratio of Triclosan to the polymers was kept at 30 wt %. The mixed polymer/Triclosan solution was then added to 7 mL of acetate buffer (pH 4.5, 100 mM), followed by dialysis to remove tetrahydrofuran.
Triclosan-Release by Loaded Mixed-Shell and Single-Shell Polymeric Micelles. To determine the release of Triclosan from both types of micelles, 2 mL of freshly prepared Triclosan-loaded micelle suspensions (1.0 mg mL−1) was transferred into a dialysis bag (molecular weight cutoff: 12−14 kDa), and subsequently immersed in 20 mL of a 10 mM phosphate buffer (pH 5.0 or pH 7.4) at 37 °C. Aliquots (1 mL) of the dialysis solution were collected every 30 min up to 50 h, and the absorbance of the solutions at 281 nm
2
was recorded on an UV−vis spectrophotometer (Shimadzu, Japan). The volume of the stock dialysis solution was kept constant by adding 1 mL of fresh buffer after each aliquot was taken. With the use of a calibration curve obtained over a Triclosan-concentration range from 2 to 100 μg mL−1, the UV absorbance was correlated to the amounts of Triclosan that released from the micelles. A key assumption of the current work is that bacterial enzymes such as lipase that exist in the biofilm can hydrolyze the micelle PCL core to speed up the release of the loaded antimicrobials. Therefore, we also studied lipase-triggered release of Triclosan from both types of micelles. Lipase (Sigma, from Pseudomonas cepacia, ≥30 U mg−1
) was added to the micelle suspension to a final concentration of 0.5 mg mL−1
. Triclosan release was measured as described above.
Culturing and Harvesting of Bacterial Strains. Two bacterial strains were employed in this study: S.
aureus ATCC12600GFP and bioluminescent S. aureus Xen36 (PerkinElmer, Inc., Waltham, MA). Both strains were cultured from cryopreservative beads (Protect Technical Surface Consultants Ltd., U.K.) onto tryptone soy agar plates (TSB, OXOID, Basingstoke, U.K.) at 37 °C in ambient air. S. aureus ATCC12600GFP
was cultured on agar plates with 10 μg mL−1
tetracycline, while S. aureus Xen36 was cultured on agar plates with 200
μg mL−1 kanamycin. For experiments, one colony was transferred to inoculate 10 mL of tryptone soy broth (TSB, OXOID, Basingstoke, U.K.) at 37 °C for 24 h in ambient air. For culturing S. aureus ATCC12600GFP,
10 μg mL−1
tetracycline was added to the TSB, while S. aureus Xen36 was grown without further addition of antibiotics. This preculture was diluted 1:20 in 200 mL of TSB and grown statically for 16 h at 37 °C. Cultures were harvested by centrifugation for 5 min at 5000g, washed twice in phosphate buffered saline (PBS, 5 mM K2HPO4, 5mM KH2PO4, and 150 mM NaCl, pH 7.0), sonicated 3× for 10 s (Vibra cell model 375, Sonics
and Material, Inc., Danbury, CT) while cooling in an ice/water bath to break possible aggregates, and finally suspended in 10 mL of PBS to a concentration of 3 × 108
bacteria mL−1
, as determined in a Bü rker-Tü rk counting chamber.
Zeta Potentials of Micelles and Bacterial and Hydrodynamic Diameters of Micelles. Zeta potentials of the micelles and both staphylococcal strains were measured at 25 °C using a Zetasizer Nano- ZS (Malvern Instruments, Worcestershire, U.K.) in 10 mM phosphate buffer over the pH range from 5.0 to 7.4. The micelle concentration was 0.5 mg mL−1
, while bacteria were suspended at a concentration 1 × 108
bacteria mL−1
. In addition, hydrodynamic radii of the micelles were measured using the Zetasizer.
Interaction of Nile Red Loaded Micelles with Planktonic Staphylococci and Staphylococcal Biofilms. Loading the micelles with Nile red was essentially done as described above for Triclosan. Briefly, a Nile red stock solution in dimethylformamide (1 mg mL−1
, 30 wt % with respect to the amount of polymer) was mixed with polymer solution in dimethylformamide (1 mg mL−1
) to a total volume of 2 mL. Under rigorous magnetic stirring, acetate buffer (pH 4.5, 100 mM) was dropwise added to initialize the micellization, and then extra volume of acetate buffer was added to make a final polymer concentration of 0.5 mg mL−1
. After micelle formation, the resulting suspension was stirred for 2 h and dialyzed in a dialysis bag with molecular weight cutoff of 6−8 kDa against ultrapure water for 24 h. The micelle suspension was sterilized by filtering through a 0.45 μm Millipore filter into a sterilized vial in order to remove aggregates of non-encapsulated, hydrophobic Nile red molecules.
To study the interaction of the micelles with planktonic staphylococci, 300 μL of Nile red loaded SSPMs or MSPMs micelle suspensions (0.5 mg mL−1
) was mixed with 300 μL of a S. aureus ATCC12600GFP
(3 × 108
bacteria mL−1
) suspension and 500 μL of PBS with pH values adjusted to 5, 6, 7, 7.4, in sterile 2 mL Eppendorf tubes. After 1 h incubation at 37 °C, the suspension was centrifuged twice at 6500g for 5 min (centrifuge 5417R, Eppendorf, Germany) and resuspended in 1 mL of PBS with the appropriate pH value. Finally, the staphylococci were resuspended in 500 μL of PBS with the appropriate pH and 20 μL was placed on a microscope glass slide and covered with a coverslip. Micrographs were taken with a Leica DM4000B fluorescence microscope (40× plan apochromatic objective) equipped with a highly light-sensitive digital
2
CCD camera and an automatic light shutter. A laser with a wavelength of 485−495 nm was used to excite GFP and Nile red fluorescence for 3.2 and 4.0 s, respectively. The relative fluorescence intensities over the areas of the wells were processed with Fiji ImageJ.51
To study the interaction of Nile red loaded micelles with staphylococcal biofilms, a 100 μL droplet of a staphylococcal suspension (108
bacteria mL−1
) was placed on a sterile (5 × 5 × 2 mm3
) (L × W × H) poly(methyl methacrylate) slide to allow bacterial adhesion for 1 h at 37 °C. Next, the slides were washed carefully with PBS three times to remove the nonadhering bacteria and immersed into TSB medium for 24 h at 37 °C. The TSB medium was replaced with fresh TSB and the biofilm was grown for another 24 h. Finally, the slides with biofilm attached were washed three times with PBS. The biofilm was then exposed to PBS buffer containing Nile red loaded micelles. The penetration of the micelles into the biofilms was subsequently studied with CLSM (LEICA TCS SP2 Leica, Wetzlar, Germany) with a HCX APO L40×/0.80WU-V-1 objective. An argon ion laser at 488 nm and a green HeNe laser were used to excite the GFP and Nile red in the samples, respectively, and fluorescence was collected at 500−535 nm (GFP) and 583−688 nm (Nile red). All data were acquired and analyzed using Leica software, version 2.0.
Killing of Planktonic Staphylococci by Triclosan and Triclosan-Loaded Micelles. To determine the MIC and MBC, 100 μL of Triclosan solutions or Triclosan-loaded micelle suspensions (both with an equivalent amount of Triclosan between 0 and 80 μg mL−1
) was applied to 100 μL of a S. aureus ATCC12600GFP
suspension in PBS (3 × 108
bacteria mL–1
) with pH adjusted to 5, 6, 7, or 7.4. The MIC values were taken as the lowest Triclosan concentration at which bacterial growth was absent. Subsequently, the MBC values were determined by plating aliquots of suspensions with concentrations yielding no visible growth of bacteria on TSB agar plates after being incubated for 24 h at 37 °C, and the lowest concentration at which colony formation remained absent was taken as the MBC.
Killing of Staphylococci in Their Biofilm Mode of Growth by Triclosan and Triclosan-Loaded Micelles. To determine the killing efficiency of staphylococci in their biofilm mode of growth by Triclosan in solution or Triclosan-loaded micelles, biofilms were grown by adding 100 μL of S. aureus Xen36 or S. aureus ATCC12600GFP
suspension (108
bacteria mL−1
) in PBS to 96-wells plates at 37 °C for 1 h to allow bacteria to adhere. Next, bacterial suspensions were removed and the wells were washed with 100 μL of PBS. Subsequently, 200 μL of TSB was added and bacteria were allowed to grow for 48 h at 37 °C. To study Triclosan penetration and retention, 100 μL of solutions of Triclosan or suspensions with Triclosan-loaded micelles (both with an equivalent amount of Triclosan between 0 and 80 μg mL−1
) at pH 7.4 was added to the biofilms. A bio-optical Imaging System (IVIS Lumina II, Imaging System, PerkinElmer) was used to detect the bioluminescence of S. aureus Xen36 and fluorescence of S. aureus ATCC12600GFP
biofilms exposed either to Triclosan or Triclosan-loaded micelles. Bioluminescence or fluorescence images were taken every hour up to 5 h (image acquisition factors: 20 s exposure time, medium binning, 1 F/Stop, Open Emission Filter). Images were automatically corrected for background noise. Regions of interest (ROIs) were manually created for each well and average radiances over the ROIs were converted to photon fluxes (p/s) using Living Image software (PerkinElmer).
Statistical Analysis. All data are expressed as means ± SD. Differences between groups were examined for statistical significance with two-tailed Student’s t test, accepting significance at p < 0.05.
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SUPPORTING INFORMATION
O O O O OH n Sn(Oct)2, toluene reflux, 16 h O O O O O H n m PEG-OH (CL) PEG-b-PCL O HN OH O O O Sn(Oct)2, toluene reflux, 16 h (CL) O HN O O O H m PCL-OH O Cl O Et3N O HN O O O m PCL-allyl O O HN NH(TDP) O O O O (HDD) O HN O O O m PCL-b-PAE O O N N O O O O N NH nScheme S1. Synthesis of (A) poly(ethylene glycol)-block-poly(ε-caprolactone) (PEG-b-PCL) and
(B) poly(ε-caprolactone)-block-poly(β-amino ester) (PCL-b-PAE). HDD refers to
Hexane-1,6-dioldiacrylate, TDP to 4,4’-trimethylene dipiperidine.
Table S1. Drug loading content (DLC) and Drug loading efficiency (DLE) of Triclosan-loaded micelles. DLC and DLE were calculated according to
DLC (wt %) = Wloaded drug/W(loaded drug and polymer) ×100%
DLE (%)= Wloaded drug/Wdrug in feed × 100%
Micelle type DLC (wt%) DLE (wt%)
Triclosan-loaded SSPMs 12.8 ± 0.6 49.0 ± 1.1 Triclosan-loaded MSPMs 19.0 ± 0.3 78.0 ± 2.1
Note: the weight of the loaded drug and polymer, as well as of the drug in the feed were measured gravimetrically, while the weight of the loaded drug was measured using UV-VIS.
