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The pressing need to treat multi-drug resistant bacteria in the chronically infected lungs of cystic fibrosis (CF) patients has given rise to novel nebulized antimicrobials. We have synthesized a silver–carbene complex (SCC10) active against a variety of bacterial strains associated with CF and chronic lung infections. Our studies have demonstrated that SCC10-loaded into l-tyrosine polyphosphate nanoparticles (LTP NPs) exhibits excellent antimicrobial activity in vitro and in vivo against the CF relevant bacteria Pseudomonas aeruginosa. Encapsulation of SCC10 in LTP NPs provides sustained release of the antimicrobial over the course of several days translating into efficacious results in vivo with only two administered doses over a 72 h period.
Cystic fibrosis (CF) is a life-threatening autosomal recessive disorder caused by mutations in the gene encoding the cystic fibrosis transmembrane conductance regulator . CF is a multisystem disease that affects the lungs and upper respiratory tract, the gastrointestinal tract, pancreas, liver, sweat glands and genitourinary tract. Most CF patients experience polymicrobial airway colonization with pathogens including Staphylococcus aureus, Burkholderia cepacia complex organisms, and in particular, Pseudomonas aeruginosa followed by intense inflammation in the lungs. A progressive decline in lung function usually occurs, which can lead to respiratory failure and death. The present treatment for chronically infected CF patients consists of frequent high-dose administration of intravenous antibiotics including aminoglycosides and β-lactams active against P. aeruginosa, such as tobramycin and ticarcillin plus clavulanic acid, respectively. Even with this type of aggressive therapy, complete eradication of the bacteria is difficult, due to the ability of the organisms to form biofilms . The persistence of bacteria has lead to the rise of resistant bacterial strains with enhanced tolerance to current antibiotics, leading to chronic infection in the lungs. In addition, the high and frequent dosing of the intravenous antibiotics necessary to achieve significant serum concentration of the drugs can cause serious side effects including, in the case of aminoglycosides, nephrotoxicity and ototoxicity .
In order to address these two major challenges, resistance and toxicity, a new form of therapy is necessary. Silver based antimicrobials have become an attractive alternative to address the resistance problem. Silver has been used since ancient times to treat infections and is still widely used in a variety of medical applications such as the treatment of burns, coatings on catheters and endotracheal tubes, disinfectants and wound dressings [4,5]. Of major importance are the relatively few reported accounts of silver resistance despite its wide and continuous use [6–10]. The rarity of silver resistance may be attributed to the multiple suggested mechanisms of action of silver. One theory suggests that Ag(I) ions cause structural changes to the bacterial cell wall by reacting with surface proteins [11,12]. Furthermore, silver ions have been thought to bind and inactivate bacterial DNA and RNA [11,12]. Likely due to the multiple mechanisms of cellular killing through fundamental cell processes, silver has broad spectrum activity. Thus, silver is very effective against a wide variety of Gram-positive and Gram-negative bacteria including S. aureus, P. aeruginosa and Escherichia coli . Our group has reported the antimicrobial activity of several silver N-heterocyclic carbene complexes (SCCs) against P. aeruginosa and B. cepacia complex species [13–15]. In addition, recent studies have reported the in vitro antimicrobial activity of silver, in the form of silver sulfadiazine, on mature biofilms of P. aeruginosa .
To combat the second major problem, antibiotic toxicity, direct administration of antimicrobials to the lungs via inhalation has been explored [16,17]. Aerosolized antibiotics have proved effective due to localized delivery at the site of infection. Direct delivery to the organs of interest has other favorable outcomes such as lower necessary dosages for therapeutic outcomes, decreased systemic toxicity, and diminished systemic absorption. However, the challenge of rapid clearance from the lungs as a result of the relatively small size of the antibiotics has been a major problem [17,18]. In order to maintain a drug concentration in the lungs above the minimum inhibitory concentration (MIC) effective against pulmonary pathogens, several inhaled treatments must be administered throughout the day. To avoid this inconvenience, we have explored the use of inhaled nanoparticles (NPs) to deliver, directly to the site of infection, a novel silver N-heterocyclic carbene complex, SCC10, which possesses potent antimicrobial properties. This unique formulation will allow the sustained release of SCC10, thereby creating a depot effect in chronically infected lungs. Sustained release of the active drug also has the advantage of decreased administration frequency, which should lead to improved patient compliance.
Unless otherwise specified, all manipulations were carried out under aerobic conditions. 4,5-Dichloroimidazole (Alfa Aesar), iodomethane (Alfa Aesar), sodium tetraphenylborate (Acros Organics), silver acetate (Acros Organics) and bromohexane (TCI) were all used as received. The l-tyrosine polyphosphate (LTP) polymer, with a molecular weight of 8–10 kDa, was prepared using established methods [19,20]. The molecular structure of LTP used for this study has been previously published . Linear polyethylenimine (LPEI) with a molecular weight of 25 kDa was purchased from PolyScience Inc. Polyethylene glycol grafted to chitosan (PEG-g-CHN) (80% acetylation) was purchased from CarboMer Inc. The surfactant, 10% polyvinylpyrrolidone (PVP) was purchased from Sigma–Aldrich. Dry solvents were used, which were obtained from a PureSolv™ solvent purification system. Water was distilled and deionized with Barnstead NanoPure II System. 1H and 13C NMR data were collected on a Varian Gemini 300 MHz instrument. The 1H and 13C NMR spectra obtained were referenced to the residual protons of the deuterated solvents and the solvent resonances, respectively. Mass spectrometry data were collected on a Bruker Daltons (Billerica, MA) Esquire-LC mass spectrometer equipped with ESI. Elemental analyses were performed at the Microanalyses Laboratory at The University of Illinois and Galbraith Laboratories.
4,5-Dichloroimidazole (1.23 g, 9.00 mmol) and potassium hydroxide (2.24 g, 40.0 mmol) were stirred in acetonitrile (50 mL) for 2 h at room temperature (Scheme 1). The unreacted KOH was filtered from the solution and bromohexane (1.26 mL, 9.00 mmol) was added (Eq. 1). The reaction mixture was refluxed for 24 h. Excess iodomethane (2.5 mL, 40.2 mmol) was added. The mixture was refluxed at 85 °C for 24 h. The volatile components were removed and the crude product was redissolved in dichloromethane. The solid, presumably KBr, was filtered and discarded and the volatile components were removed in vacuo to yield a yellow oily product. The product was stirred in diethyl ether and filtered to yield a yellow powder. Characterization results including the 1H and 13C NMR spectra of 1 can be found in the Supplementary data.
For recrystallization purposes, 1 (0.5 mmol, 0.182 g) was dissolved in water (20 ml) and sodium tetraphenylborate (1 mmol, 0.342 g) was added. A white solid of the tetraphenylborate (BPh¯4) adduct, 1a, precipitated from the solution and was filtered and washed with 10 mL of water. Crystals suitable for single X-ray diffraction studies were grown from acetone. The X-ray crystal structure determination detail including the crystal structure analysis of 1a can be found in the Supplementary data.
Compound 1 (2.20 g, 6.07 mmol) was dissolved in dichloromethane (50 mL) and silver acetate (2.03 g, 12.1 mmol) was added. The mixture was stirred at room temperature for 3 h. The yellow precipitate, presumably AgI, was filtered and discarded. The volume of the reaction mixture was reduced under pressure to 5 mL. Hexane (300 mL) was added and the mixture was cooled in an ice bath. A fine off-white solid precipitated from the solution. It was filtered and washed with 20 mL of hexane. The powder was placed under a vacuum overnight to ensure all traces of volatiles had been removed. Characterization results including the 1H and 13C NMR spectra of SCC10 can be found in the Supplementary data.
NPs loaded with SCC10 were prepared using water-in-oil-in-water emulsion method  and solvent evaporation. For the blank LTP nanoparticle formulation, the initial water-in-oil emulsion consisted of 300 mg of LTP dissolved in 3.0 mL of chloroform, 3.0 mg of PEG-g-CHN dissolved in 0.9 mL of 0.1 N acetic acid (for 48 h at 37 °C under rotation), 3 mg of LPEI in 1 mL of distilled and deionized water (DH2O), and 1 mL of DH2O in a double-neck round-bottom flask. The initial water-in-oil emulsion was vortexed for 2 min at 3000 rpm by an impeller. The second water-in-oil-in water emulsion had 100 mL of 10% PVP containing 10 mg/mL NaNO3 added to the initial emulsion and vortexed for 3 min at 1600 rpm. Chloroform was allowed to evaporate for 5 h while being stirred and vented. NPs were collected and washed with DH2O by centrifugation at 6000 × g for 10 min. Finally, the NPs were shell frozen in 10 mL of DH2O, and placed in a lyophilizer for 72 h. The lyophilized NPs were stored in a desiccator. To prepare SCC10–LTP NPs, the only deviation from the previously described method is dissolving both SCC10 and LTP into 3 mL of chloroform. The resulting NPs were aliquoted and stored in desiccated containers until the day of the experiment.
Scanning electron microscopy (SEM, Hitachi S2150) was used to measure the morphology of the NPs. The SEM samples were prepared by suspending 1 mg of NPs in 1 mL of DH2O. Then, 200 μL of the suspended NPs was transferred onto a stub, dehydrated, sputter coated with silver/palladium, and examined.
Dynamic laser light scattering was used to further quantify the size of the LTP NPs. The nanoparticle sample was prepared by suspending 1 mg of NPs in 10 mL of 1× phosphate buffered saline (PBS) that was previously passed through a 0.2-μm filter. The suspended NPs were centrifuged at 1000 × g for 10 s to remove any large aggregates. Then, samples were decanted into a glass scintillation vial. A dynamic laser light scattering system (Brookhaven Instruments BI-200SM) calculated the nanoparticle diameter by the Regularized Non-negatively Constrained Least Squares (CONTIN) method. The range of nanoparticle size was reported as differential distribution values.
The loading of SCC10 was determined by measuring the amount of SCC10 extracted from the NPs using an absorbance plate reader (Molecular Devices). Blank LTP (1 mg) or SCC10–LTP NPs (1 mg) were dissolved in 600 μL of chloroform for 15 min in 1.5 mL centrifuge tubes. HCl (400 μL, 0.1 N) was added to the dissolved NPs, emulsified for 2 min by shaking, and allowed to phase separate for 10 min. A sample of the HCl phase (200 μL) was diluted to 1 mL, and the absorbance of the precipitated silver was measured from 450 to 510 nm using an absorbance plate reader (Molecular Devices). The concentration of the SCC10 was calculated according to a standard curve.
Release profiles were determined by measuring the absorbance of the supernatant acquired from SCC10–LTP NPs incubated in PBS at physiological temperature. Release samples were prepared by suspending 1 mg of NPs in 1 mL of PBS and incubated at 37 °C under rotation. At pre-determined intervals, the samples were centrifuged at 6000 × g and 900 μL of buffer was removed and exchanged with fresh buffer. The samples were lyophilized and suspended in 1 mL of chloroform. The absorbance was measured at a pre-determined wavelength using an absorbance plate reader and the SCC concentration was calculated according to a standard curve.
Both SCC10–LTP NPs and the blank LTP NPs were reconstituted in 5 mL of a phosphate buffer (18 mg/mL Na2HPO4 and 3 mg/mL KH2PO4) to a concentration of 7.2 and 6.8 mg of NPs/mL, respectively. The solution contained no Cl− to avoid precipitation of AgCl. The reconstituted loaded and blank LTP NPs were delivered via an Aeroneb Lab apparatus (Aerogen Inc., Galway, Ireland) connected to a multi-dosing animal chamber. The Aerogen nebulizer is based on micropump technology that produces fine particles (1–5 μm) in a low velocity aerosol . The multi-dosing chamber is a square plexiglass box with inner dimensions of 8 × 8 × 4.5 inches height. The nebulizer is mounted in the center of the lid. To test whether the LTP NPs could be readily nebulized, approximately 5 mg of SCC10–LTP NPs was suspended in 1 mL of phosphate buffer. The NPs were placed into the nebulizer, aerosolized, and the vapor was collected directly into a 50 mL conical tube. The condensed mist was placed on a glass slide and examined under a Zeiss microscope. Microscopy images were acquired using Axiovision 3.1 software (Zeiss).
The P. aeruginosa isolate designed PAO1 is a standard laboratory strain. A clinical isolate of P. aeruginosa designed PA M57-15 was provided by Dr. Thomas Ferkol (Washington University, St. Louis, MO). This isolate is a mucoid strain obtained from a patient with cystic fibrosis and has been extensively studied in animal models . The E. coli strains were provided by Dr. Simon Silver (University Of Chicago, IL). The J53 strain is known to be sensitive to killing by silver cations and serves as a positive control. In contrast, the J53 + pGM101 is a J53 derivative that harbors the pGM101 plasmid originally conferring silver resistance to a burn ward isolate of Salmonella and serves as a negative control. B. cepacia complex species either were obtained from the Clinical Microbiology Laboratory at St. Louis Children's Hospital (CF clinical isolates of Burkholderia multivorans) or were provided by Dr. John LiPuma of Ann Arbor, MI, Dr. Johannes Huebner of Boston, MA (CF clinical isolates of Burkholderia dolosa), or the American Type Culture Collection.
Male C57BL/6J mice (Jackson Laboratories, Bar Harbor, Maine) at 7–10 weeks of age were used for these studies that were approved by the Washington University School of Medicine Animal Studies Committee. Animals were housed in a barrier facility under pathogen-free conditions until they were inoculated with bacteria.
Minimal inhibitory concentrations (MICs) and minimum bactericidal concentrations (MBCs) were determined by a standard Clinical and Laboratory Standards Institute (CLSI) micro-dilution method. Bacteria were streaked from glycerol-frozen stocks onto blood agar plates and incubated overnight at 37 °C. Cells from the fresh plates were suspended in the CLSI standard Mueller–Hinton broth (M–H) to an optical density at 650 nm (OD650) of 0.25 and grown at 37 °C in a shaking incubator at 200 rpm to an OD650 of 0.4, which corresponds to ~2 × 108 colony forming units (CFU)/mL, confirmed by plating serial dilutions. The bacteria were diluted in broth to a concentration of 105 CFU in 100 μL, which was added to triplicate wells of a 96-well plate containing 100 μL of SCC10 stock diluted in sterile water. The SCC10 stock solution was made by dissolving 10 mg of SCC10 in 1 mL of DMSO. The final SCC10 concentrations tested were, 1, 2, 4, 6, 8, and 10 μg/mL. The plate was incubated for 18–20 h at 37 °C and the MIC determined as the lowest concentration with clear wells. The MBC of SCC10 was determined by plating the wells with growth inhibition (clear) on blood agar plates and noting the lowest concentration that resulted in no growth.
A clinical isolate of P. aeruginosa, PA M57-15, was streaked from glycerol-frozen stocks onto tryptic soy agar (TSA) plates and incubated overnight at 37 °C. Cells from the fresh plates were suspended in M–H (5 mL) to an OD650 of 0.2 and grown at 37 °C in a shaking incubator at 200 rpm to an OD650 of 0.4 (~2 × 108 CFU/mL). Serial dilutions of SCC10 and SCC10–LTP NPs were made from stock solutions of 10 mg/mL and 64 μg (SCC10 component)/mL, respectively. Aliquots of 200 μL of the bacterial suspension were loaded in triplicate into wells of a 96-well plate containing 50 μL of SCC10 or SCC10–LTP NPs diluted in sterile water to give final concentrations of SCC10 or SCC10–LTP NPs of 4, 8, 16 and 32 μg/mL. The 96-well plate was placed in a shaking microplate reader (Synergy 2 Plate reader, BioTek) overnight at 37 °C with OD650 readings every 15 min. Data were graphed using Prism 4 (GraphPad Software, Inc., San Diego, CA).
Mice were anesthetized using brief exposure to isoflurane. PA M57-15 in Luria broth (LB) was delivered to each mouse intranasally. The dose ranged from 1.1 × 107 to 4.3 × 107 CFU per mouse in each of 4 separate experiments with 8 mice per group. Mice were weighed daily and observed for a total of 72 h after inoculation for survival. To allow nose-only delivery of drug, animals were placed in CH-247 tubes (CH Technologies, Westwood, NJ). Four animals at a time, each housed in individual tubes, were placed into a multi-dosing chamber. One hour after inoculation with P. aeruginosa, the animals were exposed for 15 min to either nebulized blank LTP NPs (34 mg) or SCC10–LTP NPs (36 mg) suspended in 5 mL each of phosphate buffer. The animals received a second nebulized dose of NPs 24 h after the first. Once the concentration throughout the dosing chamber equilibrated as indicated by diffusion of the mist cloud, the tail movements of the animals were used as an indicator of their status. Mice were then euthanized 72 h after inoculation by carbon dioxide inhalation. Lungs and spleens were harvested for quantitative bacteriology within 1 h after death for animals that did not survive to 72 h and immediately after euthanasia for the remaining animals as previously described . The organs were homogenized in 1 mL of LB broth (PowerGen 125 homogenizer, Fischer Scientific) and after performing a 10-fold serial dilution, 10 μL of each of the dilutions was placed in duplicate on TSA plates overnight at 37 °C. Presence or absence of bacterial colonies was noted to determine bacteremia. The results are pooled from 4 separate experiments for a total of 16 animals in each treatment group.
All analyses were performed using Prism 4. The in vivo survival curves in the infection model were compared using a log-rank test. Changes in animal weights were compared by ANOVA. The lung and spleen bacteriology of surviving animals were compared by t-test. Data are given as mean ± standard deviation. The risk of bacteremia between the two groups, animals receiving blank LTP NPs versus SCC10–LTP NPs, was compared by Fisher's exact test of a contingency table (number of animals with bacteria in the spleen versus number of animals cleared).
The 1H NMR spectrum of the imidazolium salt precursor 1 shows a resonance at 9.51 ppm which is indicative of the formation of an imidazolium salt and is consistent with the general C2–H acidic proton shift of other imidazolium salts (δ 8–10 ppm) . The 13C NMR shift of the N–C–N sp2 carbon, which later becomes the carbene center was observed at 136.3 ppm.
The molecular structure of the cationic portion of 1 obtained by X-ray crystallography is depicted in Fig. 1. Following metallation of 1 with silver acetate, the 1H NMR spectrum of SCC10 showed a loss of the imidazolium proton resonance at 9.51 ppm indicating carbene formation. The most definitive evidence for the formation of the silver–carbene was observed in the 13C NMR spectrum of SCC10. A shift of the resonance corresponding to the N–C–N sp2 carbon from 136.3 ppm to 179.6 ppm was observed confirming formation of a carbene center. This shift is consistent with those typically observed for N-heterocyclic carbene complexes of Ag(I) [25,26].
SCC10 was encapsulated into NPs formulated with LTP using a water-in-oil-in-water emulsion technique. The resulting NPs are spherical with smooth surface morphology for both SCC10–LTP NPs and blank LTP NPs (Fig. 2A and B, respectively) as determined by scanning electron microscopy (SEM). Using emulsion techniques, the encapsulation of low molecular weight drugs can be difficult since the encapsulated drugs can diffuse into the external water phase during the solvent evaporation [27,28]. Indeed, previous attempts to encapsulate silver–carbene complexes without a hydrophobic group resulted in undetectable amounts of the drug as determined by loading studies. Thus, the conjugation of the hexyl side chain and the ionic charge gradient created by the addition of the NaNO3 in the external aqueous phase are carefully designed to confine SCC10 in the polymeric phase. Furthermore, incorporation of PEG-g-CHN and LPEI during nanoparticle formulation stabilizes the emulsion and prevents aggregations . Due to the amphiphilic nature of these polymers, they enriched at the water–oil interface and prevented coalescence of the LTP droplets by steric repulsion [29,30].
This method for encapsulating SCC10 has yielded NPs with properties that may alleviate rapid clearance of SCC10 from the lungs and allow delivery to biofilms in the airways. Encapsulating low molecular weight drugs into NPs has been shown to retard drug clearance from tissues [31,32]. Furthermore, studies by Owusu-Ababio et al. have shown that NPs are effective for delivering antimicrobial drugs into biofilm infections . To effectively treat biofilms in small airways, NPs must be small enough to navigate through the respiratory tract without impaction in the nasopharynx, while large enough to deposit onto the matrix of the biofilms [34–36]. The sizes of the blank LTP and SCC10–LTP NPs quantified using a dynamic laser light scattering system, ranged between 471 and 2891 nm with median diameter of 922 nm and between 555 and 1519 nm with median diameter of 1172 nm, respectively (Fig. 3). Hence, these NPs are the proper size to navigate through the respiratory tract and deposit onto biofilms. Furthermore, both SCC10–LTP NPs and LTP NPs were delivered in a suspension form by a nebulizer that produces a narrow range of droplet size, between 4 and 6 μm, optimized for delivery to human airways.
The encapsulation techniques for preparing SCC10–LTP NPs resulted in satisfactory loading efficiency. Our minimum target for SCC10 loading was 2.5% of the total polymer weight to ensure optimal dosing while minimizing the delivery of polymer. The loading has been determined by quantifying the amount of silver extracted from LTP NPs via absorbance spectrophotography. The LTP NPs were formulated to be 10% (w/w) of SCC10; however, the actual loading was determined to be 7.1 ±1%. The encapsulation efficiency translates to 71 ± 10%.
Release of the encapsulated SCC10 from the LTP NPs was characterized to determine whether this formulation would provide the desired depot delivery of the drug. The LTP NPs release the entire encapsulated drug in 7 days under in vitro incubation in PBS (Fig. 4). The cumulative release after 7 days from 1 mg of LTP NPs was 74 ± 10 μg (Fig. 4), which is comparable to the loading data of 71 ±1 μg. Approximately 80% of the SCC10 is released within the first 2 days from the LTP NPs, which corresponds to 40 μg (54%) and 20 μg (27%) SCC10 per 1 mg of NPs on days 1 and 2, respectively. For the next 5 days, the release amounts of SCC10 are minimal. This release rate is consistent with previous studies that attribute the rapid degradation rates to the presence of hydrolytically unstable phosphoester linkages in LTP's polymeric backbone [19,21].
The rapid degradation rate makes LTP NPs ideal candidates for drug delivery into the lungs, because they do not accumulate in the airways. Furthermore, the degradation products of the LTP polymer have been shown to result in an insignificant decrease in the local pH, unlike other biomaterials such as poly[lactic-co-glycolic acid] [19,20]. The probable nontoxic degradation products of LTP include l-tyrosine based derivatives, phosphate ions, and alcohols . These degradation products are easily metabolized or excreted.
The in vitro antimicrobial properties of the SCC10–LTP NPs were characterized through comparison with the well-studied SCC10 parent compound, SCC5 . Using a standard micro-dilution assay, the MIC of SCC5, when assayed in the presence of phosphate buffer (18 mg/mL Na2HPO4 and 3 mg/mL KH2PO4 without NaCl), was uniformly 1 μg/mL for all organisms tested excluding the E. coli J53 strain with the silver resistant plasmid pMG101 (Table 1). Both these MICs and the corresponding MBCs were lower than those reported previously for SCC5 against these organisms . The antimicrobial activity of SCC10 is likely linked to the release of the Ag(I) from the carbene, and thus, the free-Ag(I) concentration in the media determines the efficacy. The phosphate buffer in these assays may have driven a more rapid dissociation of the active Ag(I) from the carbene providing a higher free-Ag(I) concentration. The effects of DMSO on the MIC of SCC5 were evaluated in order to insure that the apparent MICs and MBCs of SCC10, which must be solubilized with DMSO due to the hydrophobic nature of the molecule, would not be falsely lowered due to DMSO toxicity. The DMSO changed the MIC and MBC of SCC5 against only one organism, P. aeruginosa strain PA M57-15, and rather than lowering the MIC, DMSO raised it. While no explanation for this observation is available yet, it is likely that the efficacy of SCC10 was not falsely overestimated due to the presence of DMSO in the assay.
SCC10 appeared equally efficacious in vitro compared with SCC5. The MICs of SCC10 were again, uniformly 1 μg/mL for non-silver resistant organisms. The MBCs of SCC10 were lower than those of SCC5. Encapsulating SCC10 in the LTPNPs resulted in an increase in the apparent MIC of the SCC10 component as one would expect due to the slow release and hence, lower free-Ag(I) concentration at any given SCC10 concentration. As mentioned, SCC5 cannot be readily encapsulated into LTPNPs using current techniques. Consequently, no SCC5-LTP NPs were tested. The LTP NPs themselves had no antimicrobial activity in these assays. Additional MIC and MBC studies of SCC10 on other CF relevant bacteria can be found in the Supplementary data.
To further explore the impact of SCC10 and SCC10–LTP NPs on the growth properties of the P. aeruginosa strain PA M57-15, the bacterial growth was measured at OD650 in a microplate spectrophotometer in the presence of free SCC10 and SCC10–LTP NPs (Fig. 5). The control PA M57-15 growth curve exhibited log growth between 1 and 5 h. SCC10 at 4 μg/mL shifted the onset of log growth to between 3 and 8 h, while SCC10 concentrations of 8 μg/mL and above inhibited bacterial growth. In this assay, the apparent MIC of SCC10 against PA M57-15 is between 4 and 8 μg/mL, which correlates well with the MIC of 4 μg/mL reported previously for SCC5 against PA M57-15 assayed using standard microtiter dilution methods, also in the absence of the phosphate buffer .
In contrast, incubation of PA M57-15 with SCC10–LTP NPs to give final SCC10 concentrations in the microtiter plate well of up to 8 μg/mL resulted in no obvious growth inhibition, as the growth curves appeared similar to the control. At concentrations of SCC10 in LTP NPs of 16 and 32 μg/mL, the lag before onset of log phase growth increased in a dose-dependent manner from 2 to 5 h indicating growth inhibition (Fig. 5B). Once again, the free-Ag(I) concentration in the media containing SCC10–LTP NPs is lower than that of media containing the equivalent concentration of SCC10 due to the slow release of the intact SCC10 prior to subsequent free-Ag(I) release. Thus, we have illustrated the depot delivery properties of the SCC10–LTP NPs in a biologically relevant assay.
Different solutions are nebulized with different efficiencies via the micropump technology . Similar to the findings of others , the efficiency appears in our hands to correlate with the ionic strength of the liquid, in that solutions with higher ionic strength nebulize more readily. Because the outer shell of LTP NPs is predicted to be non-polar, they were not predicted to nebulize efficiently when suspended in water without the addition of salts. This hypothesis was tested directly by suspending LTP NPs in water, placing an aliquot into the nebulizer and attempting to collect the aerosolized suspension. As expected, the nebulization of phosphate buffer alone was free of NPs. In contrast, condensate from nebulized SCC10–LTP NPs had a concentration equal to that of the original suspension demonstrating that NPs readily passed through the nebulizer (Fig. 6). These results indicate that the nebulization of the LTP NPs is a viable method for localized delivery of SCC10 into the lungs.
Encapsulating SCC10 into degradable NPs provides an avenue for targeting bacteria in biofilms. Targeting moieties such as peptides specific to the P. aeruginosa bacteria can be easily incorporated into the nanoparticle formulations. For example, toll-like receptors that recognize the lipopolysaccharide on the membranes of Gram-negative bacteria including P. aeruginosa could be bound to the surfaces of NPs [39,40]. Currently, polysaccharides and antibodies have been used in nanoparticle formulations for targeting to specific cells and bacteria [41,42].
As an initial evaluation of the in vivo antimicrobial effects of LTP NPs, two doses of aerosolized blank LTP and SCC10–LTP NPs were delivered 24 h apart in a nose-only fashion to P. aeruginosa-infected mice in a multi-dosing chamber (Fig. 7A). Treatment with SCC10–LTP NPs resulted in an almost 20% survival advantage (Fig. 7B: 12/16, 75% SCC10–LTP NPs versus 9/16, 56% blank LTP NPs; p = 0.23) and the surviving animals demonstrated less weight loss compared with surviving animals treated with blank NPs (Fig. 7C: p = 0.009 by ANOVA). In other studies, mice were pretreated with a 5 mg nebulized dose of free SCC1, a silver complex with a methylated caffeine carrier, prior to infection with P. aeruginosa (unpublished results). The mice then received 8 more doses of 5 mg each over a 72-hour study period to achieve a survival advantage of approximately 26%. Thus, treatment with SCC10–LTP NPs using less than one tenth of the silver–carbene dose and only 2 doses in 72 h achieved a similar survival advantage.
To better quantify the antimicrobial effects of the NPs, the lungs and spleens of each animal were harvested for gross examination and quantitative bacteriology. The lungs from the majority of mice treated with SCC10–LTP NPs appeared grossly normal (bright pink with a smooth surface). In contrast, the lungs from most of the mice that received blank LTP NPs appeared inflamed (red or dark pink with an enlarged volume). The spleens from both groups showed no significant morphological disparity. Treatment with SCC10–LTP NPs resulted in a significant decrease in the bacterial burden in the lungs of surviving animals (Fig. 8A: 3.6 × 104 ± 2.8 × 104 SCC10–LTP NPs versus 5.3 × 105 ± 2.6 × 105 blank LTP NPs; p = 0.026, data from one animal is missing due to contamination of the plates). The bacterial counts in the lungs of animals that died were significantly higher than these in both groups and may represent post-mortem replication of bacteria (data not shown). The bacterial burden in the spleens of the surviving animals treated with SCC10–LTP NPs was less than that of the sham-treated animals (Fig. 8B: 1.3 × 102 ± 8.9 × 101 for SCC10–LTP NPs versus 5.7 × 102 ± 3.2 × 102 for blank LTP NPs), although the difference was not statistically significant (p = 0.074). Survival is likely related to bacteremia as demonstrated by dissemination of bacteria to the spleen. Indeed, all of the animals that died had bacteria recovered from their spleens. The animals treated with SCC10–LTP NPs had a significantly lower probability of dissemination of bacteria to the spleen (Fig. 8C; p = 0.029, contingency table), perhaps explaining the survival advantage. These findings indicate that treatment with SCC10–LTP NPs can effectively decrease the burden of bacteria in the lung, bacteremia, and likelihood of death in a P. aeruginosa pneumonia model.
We have illustrated through in vitro and in vivo studies that SCC10–LTP NPs are effective against the CF relevant bacteria P. aeruginosa. SCC10–LTP NPs provide sustained release of the active drug species over the course of several days providing a significant survival advantage in mouse infection models with only two doses. These results are promising for translating clinically into less frequent dosing of the antibiotic and the added advantage of improved patient compliance.
This work was supported by the University of Akron (Firestone Fellowship), Washington University School of Medicine, the National Institute of Allergies and Infectious Diseases (1 R01 A106785601), and the National Institute of General Medical Sciences (5 R01 GM086895-02). We thank the National Science Foundation (CHE-8808587 and CHE-9977144) for funds used to purchase the Varian Gemini 300 MHz and the Varian INOVA 400 MHz NMZ instruments used in this work. We would like to acknowledge the National Science Foundation (CHE-0116041) and the Ohio Board of Regents for funds used to purchase the Bruker-Nonius Apex CCD X-ray diffractometer used in this research.