Letter www.acsami.org
Bactericidal Dendritic Polycation Cloaked with Stealth Material via Lipase-Sensitive Intersegment Acquires Neutral Surface Charge without Losing Membrane-Disruptive Activity Lulu Xu,†,‡,§ Chen He,†,‡,§ Liwei Hui,‡,§ Yuntao Xie,§,⊥ Jia-Min Li,‡,§ Wei-Dong He,*,‡,§ and Lihua Yang*,‡,§ ‡
CAS Key Laboratory of Soft Matter Chemistry, §School of Chemistry and Materials Science, and ⊥CAS Key Laboratory of Materials for Energy Conversion, University of Science and Technology of China, Hefei, Anhui 230026, China S Supporting Information *
ABSTRACT: Net cationicity of membrane-disruptive antimicrobials is necessary for their activity but may elicit immune attack when administered intravenously. By cloaking a dendritic polycation (G2) with poly(caprolactone-b-ethylene glycol) (PCL-b-PEG), we obtain a nanoparticle antimicrobial, G2-g-(PCL-b-PEG), which exhibits neutral surface charge but kills >99.9% of inoculated bacterial cells at ≤8 μg/mL. The observed activity may be attributed PCL’s responsive degradation by bacterial lipase and the consequent exposure of the membrane-disruptive, bactericidal G2 core. Moreover, G2-g-(PCL-b-PEG) exhibits good colloidal stability in the presence of serum and insignificant hemolytic toxicity even at ≥2048 μg/mL. suggesting good blood compatibility required for intravenous administration. KEYWORDS: drug resistance, antimicrobial, nanoparticle, stimulus responsive, stealth coating
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Inflammation induced by microbial infection exhibits vascular permeability (i.e., enhanced permeation effect) as do cancers,26 suggesting that nanoparticles may leak from blood vessels to accumulate within infection sites. To avoid triggering immune attack, most nanoparticles developed for biomedical applications are coated with stealth materials.25,27,28 Poly(ethylene glycol) (PEG) is currently the gold standard stealth material. After PEGylation (i.e., coating a particle with PEG), even highly swollen microgels containing polyelectrolytes acquire neutral surface charge without having their interior charged groups affected.29 Therefore, we herein propose to develop a core− shell nanoparticle that has a membrane-disruptive core coated with stealth material shell via intersegment/bond that is responsive degradable by bacteria, in efforts to achieve both activity against bacterial membrane integrity and blood compatibility required for intravenous administration. A long-chain dendrimer―a highly branched polymer with systematic architecture and long subchains―exhibits a molecular conformation that can be approximated with a compact nanosphere.30 As antibacterial agents, cationic longchain dendrimers may out-perform their linear-polymer counterparts, because of enhanced abilities to adsorb on bacterial surfaces, to diffuse bacterial cell walls, to bind with bacterial cytoplasmic membranes, and to disrupt the membrane integrity.31 Note that long linear polycations preferentially disrupt membranes of bacteria over mammalian cells.32 We
he emergence of antibiotic-resistant bacteria and their accelerating spread pose a threat to global public health.1,2 It is thus imperative to develop and/or discover novel antimicrobials that are active against antibiotic-resistant bacteria and less prone to evoke resistance.2−5 Many antimicrobial peptides (AMPs) from the innate immunity of multicellular organisms act by impairing the integrity of bacterial membranes, a generic mode that appears to be more difficult for bacteria to circumvent than the metabolic targeting modes of antibiotics, and are thus widely viewed as a promising source of novel anti-infective agents.6−12 The membrane-destabilization processes are generally attributed to two structural motifs common to most AMPs: They are cationicity (i.e., an AMP has net cationic charges) and amphipathicity (i.e., an AMP has discrete cationic and hydrophobic faces).6−8 Designed to be simultaneously cationic and hydrophobic, synthetic mimics of AMPs (SMAMPs) including non-natural peptides,13−16 peptoids,17 oligomers,18−20 and polymers,20−23 have demonstrated similar in vitro antimicrobial activities as do AMPs. The net cationic charges of an AMP/SMAMP, however, may be a double-edged sword for its potential in vivo intravenous administration. On the one hand, they are necessary for its association with bacterial surface, the initial step for subsequent membrane-disruptive processes that lead to cell death.6,7,9,10,24 On the other hand, because of these cationic charges, an AMP/ SMAMP may exert nonspecific interactions with blood components and, as a result, trigger immune attack when intravenously administered in vivo.25 How to reconcile these two contradicting effects conferred by cationic charges of an AMP/SMAMP remains an ongoing challenge. © XXXX American Chemical Society
Received: October 9, 2015 Accepted: December 3, 2015
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DOI: 10.1021/acsami.5b09581 ACS Appl. Mater. Interfaces XXXX, XXX, XXX−XXX
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IR spectra, GPC traces with RI detector, and GPC traces with MALLS detector (Figure S4−8) consistently confirm the successful preparation of G2-Boc (Table S3). TFA-treatment removes Boc groups in G2-Boc and yields G2, as confirmed with the 1H NMR spectra (Figure S8). In HEPES buffer, the resulting G2 exhibits a hydrodynamic radius and a gyration radius of 12 and 13 nm, respectively, indicative of a molecular conformation that can be approximated with a compact nanosphere (Table S2 and Figure S9). Grafting PCL-b-PEG onto G2-Boc via click chemistry and subsequent TFAdeprotection successfully leads to the as-proposed G2-g(PCL-b-PEG), as confirmed consistently with 1H NMR spectra and GPC traces (Figures S11 and S12 and Table S3). Zeta-potential (ζ-potential) measurements (Figure 2b) show that whereas G2 exhibits a significantly cationic ζ-potential, G2g-(PCL-b-PEG) exhibits a neutral one, indicative of neutral surface charge, likely due to its PCL-b-PEG shell, which effectively shields the cationic charges on its G2 core.29 Dynamic light scattering (DLS) characterizations (Figure 2c and Figure S13) reveal that in contrast to G2, the G2-g-(PCL-bPEG) nanoparticle exhibits negligible increase in hydrodynamic radius over a span of 5 days in HEPES buffer supplemented with fetal bovine serum (FBS) (50%, v/v), indicative of good colloidal stability, likely due to its PCL-b-PEG shell which blocks protein adsorption onto G2 via electrostatic interactions. Closer examinations on DLS results reveal that, on day 0, G2-g(PCL-b-PEG) exhibits a large hydrodynamic radius (∼95 nm) with high scatter intensity, whereas G2 exhibits a small hydrodynamic radius (∼10 nm) with fairly low scatter intensity, suggesting that the former may form multimolecular aggregates due to its hydrophobic PCL intersegments whereas the latter is molecularly dispersed. Transmission electron microscopy images further confirm the formation of multimolecular aggregate by G2-g-(PCL-b-PEG) (Figure 2d and Figure S14). Collectively, these results suggest that, owing to PEGylation with PCL-b-PEG, G2-g-(PCL-b-PEG) exhibits neutral surface charge and good colloidal stability even in the presence of serum. PCL undergoes responsive degradation upon exposure to bacterial lipase.33,34 Will G2-g-(PCL-b-PEG) responsively
hence use long polycations as subchains for preparing the dendritic core. As a proof-of-concept, we develop G2-g-(PCL-bPEG) (Figure 1), by preparing the second generation of long-
Figure 1. Schematic illustration of G2-g-(PCL-b-PEG), a second generation of long-chain dentritic poly(ethylamino acrylate) (G2) grafted with polyethylene glycol (PEG) via bacterial lipase-sensitive poly(caprolactone) (PCL) intersegment. Prior to bacterial exposure, G2-g-(PCL-b-PEG) has neutral surface charge. Upon sensing bacteria, G2-g-(PCL-b-PEG) effectively kills both S. aureus (ball) and E. coli (cylinder) cells, likely due to the responsive degradation of PCL intersegments by lipases, enzymes produced by bacteria for lipid metabolism.
chain dendritic poly(aminoethyl acrylate) (G2) as its nanoparticle core, followed by PEGylation with poly(caprolactone-bethylene glycol), (PCL-b-PEG), in which the PCL intersegment is responsive degradable by lipases,33,34 enzymes produced by bacteria for lipid metabolism.35 Such a strategy may offer novel antimicrobials active against antibiotic-resistant bacteria yet compatible with blood components when intravenously administered. Briefly, G2 is prepared (Scheme S1) by synthesizing its Bocprotected precursor dendrimer (G2-Boc) with a divergent synthetic strategy via single-electron-transfer living radical polymerization of t-butyloxycarbonyl-aminoethyl acrylate (Boc-AEA), followed by deprotection treatment with trifluoroacetic acid (TFA) to remove Boc groups. 1H NMR spectra, FT-
Figure 2. (a) Schematic illustration of the chemical structure of G2-g-(PCL-b-PEG). (b) Zeta-potentials (ζ-potentials) of G2-g-(PCL-b-PEG) in HEPES buffer, with that of G2 as a reference. Data points are reported as mean ± standard deviation. (c) Hydrodynamic radii (Rh) of G2-g-(PCL-bPEG) in HEPES buffer supplemented with fetal bovine serum (50%, v/v) over a span of 5 days. Those of G2 are included as a reference. Data points are reported as mean ± standard deviation. (d) Transmission electron microscopy images of G2-g-(PCL-b-PEG). (e) Distributions of hydrodynamic radii (Rh) for G2-g-(PCL-b-PEG) in HEPES buffer at different time points after addition of bacterial lipase, with that of G2 included as a reference. B
DOI: 10.1021/acsami.5b09581 ACS Appl. Mater. Interfaces XXXX, XXX, XXX−XXX
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Figure 3. (a, b) Plate killing assays using (a) G2-g-(PCL-b-PEG) and (b) G2 against E. coli and S. aureus. Data points are reported as mean ± standard deviation. (c-d) Inner membrane permeability assays with E. coli ML-35 cells show that both (c) G2-g-(PCL-b-PEG) and (d) G2 induce detectable ΔOD400 signal and, to do so, their minimal threshold concentrations are 16 and 8 μg/mL, respectively. (e) Fluorescence microscopy images of E. coli and S. aureus cells show that, after 3 h treatment, cells treated with G2-g-(PCL-b-PEG) (64 μg/mL) stain intensely red, as do those treated with G2, indicative of dead cells, whereas those in control remain dark in the red channel, indicative of live cells. The treatment is carried out in sterile HEPES buffer (10 mM HEPES, 150 mM NaCl, pH 7.4), followed by staining briefly (15 min) with SYTO-9 and PI. Controls are those assayed similarly but without a nanoparticle. Scale bar = 20 μm.
nm; pretreating lipase with G2 leads to G2-lipase aggregates with Rh distribution peak also around ∼100 nm and delays, rather than completely deactivating, lipase in degrading PCL-bPEG micelle (Figure S15). We hence attribute the peak around ∼100 nm at ≥300 min after lipase addition (Figure 2e) to aggregates formed by lipase with the as-liberated PEG and/or G2 core (Figure S15), although intact or partially degraded G2g-(PCL-b-PEG) may exist as well. Collectively, these observations suggest that, upon exposure to bacterial lipase, G2-g-(PCL-b-PEG) responsively expose its G2 core as expected. Now G2-g-(PCL-b-PEG) responsively exposes its G2 core likely via PCL degradation by bacterial lipase. Are G2 and G2-g(PCL-b-PEG) antibacterial? To address this, we perform
degrade to its G2 core upon exposure to bacterial lipase? To assess this, we monitor the distribution of hydrodynamic radius (Rh) for G2-g-(PCL-b-PEG) dispersion (in HEPES buffer) at different time-points after lipase addition. Our results (Figure 2e) reveal that, within 30 min after lipase addition, the peak around 100−200 nm becomes significantly broader and shifts slightly toward the smaller Rh end than that at t = 0 min and, starting at 20 min, a broad peak around ∼10 nm emerges, indicative of degradation of G2-g-(PCL-b-PEG) toward G2. Nevertheless, at ≥300 min after lipase addition, the distribution of Rh still exhibits two well-separated and narrow peaks, rather a single peak around ∼10 nm indexable to G2. Note that lipase degrades PCL-b-PEG micelles within 30 min, leading to PEGlipase aggregates that have Rh distribution peak around ∼100 C
DOI: 10.1021/acsami.5b09581 ACS Appl. Mater. Interfaces XXXX, XXX, XXX−XXX
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Figure 4. Hemolysis assays using (a) G2-g-(PCL-b-PEG) and (b) G2 against mouse red blood cells in HEPES buffer (10 mM HEPES, 150 mM NaCl, pH 7.4). # indicates 0% hemolysis. Data points are reported as mean ± standard deviation.
respective minimum threshold dosages, ΔOD400 caused by G2g-(PCL-b-PEG) at a certain time-point is lower than that by G2, likely due to delay caused by lipase-responsive degradation of PCL in the former, suggesting necessity of PCL degradation in its activity. To assess whether G2-g-(PCL-b-PEG) and G2 permeabilize membranes of both Gram-negative and -positive bacteria, we perform bacterial Dead/Live viability assays under fluorescence microscopy (Figure 3e and Figure S16), using E. coil and P. aeruginosa as representative strains for Gram-negative bacteria, whereas S. aureus and B. subtilis as representative strains for Gram-positive bacteria. SYTO-9 and propidium iodide (PI), two nucleic acid stains with drastically different capabilities in permeating healthy membranes and spectral characteristics, are used to label all and dead bacteria, respectively. After 3-h treatment, cells treated with G2-g-(PCL-b-PEG) stain intensely red, as do those treated with G2, indicative of dead cells with compromised membranes; this is the case for all four bacterial strains tested irrespective of their Gram property. In contrast, cells treated similarly but without a nanoparticle (i.e., control) remain dark in the red channel, indicative of live cells with intact membranes. Collectively, these results suggest that both G2-g-(PCL-b-PEG) and G2 may kill Gram-negative and -positive bacteria by permeabilizing their cytoplasmic membranes (albeit other targets may exist). The G2-g-(PCL-b-PEG) is developed with intension for intravenous administration. Hence we examine its blood compatibility, as blood is the first barrier for its in vivo applicaitons. In contrast to G2, the G2-g-(PCL-b-PEG) nanoparticle exhibits good colloidal stability even in the presence of fetal bovine serum, suggesting potential of good colloidal stability in blood. Moreover, hemolysis assays using mouse red blood cells reveal that despite its antibacterial potency, G2-g-(PCL-b-PEG) induces ≤30% hemolysis up to 2048 μg/mL (Figure 4a), indicative of insignificant hemolytic toxicity and a selectivity, HC50/MBC99.9, of >256 (where HC50 is the minimal concentration to lyse 50% inoculated red blood cells). It is noteworthy that G2 lacks hemolytic toxicity as well (Figure 4b), suggesting that even after bacterial exposure that degrades G2-g-(PCL-b-PEG) to G2, the product may still lack hemolytic toxicity. Taken together, these observations suggest that G2-g-(PCL-b-PEG) may have good blood compatibility required for intravenous administration. Still, comprehensive examination on the in vivo toxicity of G2-g-(PCL-b-PEG) is necessary for its future therapeutic development, yet, it is out of the focus of this work.
bacterial killing assays, using E. coli and S. aureus as representative Gram-negative and−positive bacteria, respectively. Our results (Figure 3a) show that G2-g-(PCL-b- PEG) exhibits MBC99.9 (minimal concentration to kill 99.9% of inoculated bacterial cells) values of 4 and 8 μg/mL against E. coli and S. aureus, respectively; it is noteworthy that these MBC99.9 correspond to nM scale when expressed in molar concentration (Figure S17). In similar assays, G2 exhibits an MBC99.9 value of 4 μg/mL against S. aureus and kills 99.8% of inoculated E. coli cells at 3−8 μg/mL. The average molecular weight of G2-g-(PCL-b-PEG) is approximately 2-fold of that of G2 (Table S3). Hence, G2-g-(PCL-b-PEG) is similarly potent against bacteria as does G2 (in molar dosage), suggesting that the observed activity of G2-g-(PCL-b-PEG) may be attributed to its G2 core that is responsively exposed to bacteria via PCL degradation by bacterial-secreted lipase. G2-g-(PCL-b-PEG) and G2 exhibit similar antibacterial potency (in molar dosage). Do they act by disrupting bacterial membrane integrity? To assess this, we perform the following two membrane permeability assays. We first quantitatively examine their abilities to permealize cytoplasmic membranes of Gram-negative bacteria, by assessing the inner membrane permeability of E. coli ML-35 indicated by β-galactosidase activity using o-nitrophenyl-β-D-galactoside (ONPG) as substrate (Figure 3c and Figure 3d). The mutant E. coli ML-35 (i-, y-, z+) has constitutive cytoplasmic β-galactosidase activity but no lactose permease. Intact E. coli ML-35 cells therefore cannot hydrolyze ONPG until their cytoplasmic membranes are permeabilized to allow efflux of β-galactosidase and/or influx of ONPG across cytoplasmic membranes. o-Nitrophenol (ONP), the product of ONPG hydrolysis by β-galactosidase, shows strong absorption at 400 nm, whereas ONPG does not. Hence, the hydrolysis of ONPG catalyzed by β-galactosidase can be monitored by measuring optical density at 400 nm (OD400). As a result, the kinectics in OD400 increase reflects that in ONP production, which indicates β-galactosidase efflux and/or ONPG influx across cytoplasmic membranes. Within 120 min after addition into E. coli ML-35 suspension, G2-g(PCL-b-PEG) causes appreciable ΔOD400 once ≥16 μg/mL (Figure 3c), indicative of inner membrane disruption caused by G2-g-(PCL-b-PEG) above a minimal threshold concentration of 16 μg/mL. Similarly, G2 causes detectable ΔOD400 once ≥8 μg/mL. That the minimum threshold mass concentration of G2-g-(PCL-b-PEG) is 2-fold of that for G2 (16 versus 8 μg/ mL) may be attributed to that the former’s larger average molecular weight than the latter’s (Table S3). Closer examinations on the kinetics of ΔOD400 indicate that, at their D
DOI: 10.1021/acsami.5b09581 ACS Appl. Mater. Interfaces XXXX, XXX, XXX−XXX
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In conclusion, we have developed a core−shell polycation nanoparticle, G2-g-(PCL-b-PEG), which has neutral surface charge bestowed by its stealth material shell but, upon sensing bacteria, exhibits membrane-disruptive activity likely attributable to its bactericidal dendritic core, which is responsively exposed via PCL degradation by bacterial lipase. Moreover, G2g-(PCL-b-PEG) nanoparticle may have good blood compatibility, as indicated by its good colloidal stability in the presence of serum and lack of significant hemolytic toxicity. When administered intravenously in vivo, such an antimicrobial nanoparticle may avoid eliciting nonspecific interactions with blood components that may otherwise trigger immune attack but accumulate preferentially at infection sites via vascular permeability and eradiate pathogen bacteria therein. This work demonstrates an alternative, feasible strategy toward novel antimicrobials that may have both potency against antibioticresistant bacteria and good blood compatibility required for intravenous administration.
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ASSOCIATED CONTENT
S Supporting Information *
The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsami.5b09581. Schemes S1 and S2, Figures S1−S17, Tables S1−S3, additional Results and Discussion, as well as Materials and Methods (PDF)
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AUTHOR INFORMATION
Corresponding Authors
*E-mail:
[email protected]. *E-mail:
[email protected]. Author Contributions †
L.X and C.H. contributed equally.
Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS The E. coli ML-35 strain was a kind gift from Professor Andrè J. Ouellette at the University of Southern California. This work was supported in part by NSFC (21174138, L.Y.; 20934005 and 21274136, W.-D.H.), CAS Youth Innovation Promotion Association (L.Y.), and Ministry of Education of China (NCET-13-0547, L.Y.; FRF for CU WK2060200012, L.Y. and W.-D.H.).
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