Interaction of laponite with membrane components - consequences for

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Biological and Medical Applications of Materials and Interfaces

Interaction of laponite with membrane components consequences for bacterial aggregation and infection confinement Sara Malekkhaiat Häffner, Lina Nyström, Kathryn L Browning, Hanne Mørck Nielsen, Adam A. Strömstedt, Mariena J.A van der Plas, Artur Schmidtchen, and Martin Malmsten ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.9b03527 • Publication Date (Web): 05 Apr 2019 Downloaded from http://pubs.acs.org on April 7, 2019

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ACS Applied Materials & Interfaces

Interaction of laponite with membrane components consequences for bacterial aggregation and infection confinement Sara Malekkhaiat Häffner1,*, Lina Nyström2, Kathryn L. Browning1, Hanne Mørck Nielsen1, Adam A. Strömstedt3, Mariena J.A. van der Plas1,4, Artur Schmidtchen4,5, and Martin Malmsten1,2

1Department

of Pharmacy, University of Copenhagen, DK-2100 Copenhagen, Denmark

2Department

3Pharmacognosy,

of Pharmacy, Uppsala University, SE-75123, Uppsala, Sweden

Department of Medicinal Chemistry, Uppsala University, SE-751 23 Uppsala, Sweden

ACS Paragon Plus Environment

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ACS Applied Materials & Interfaces 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

4Division

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of Dermatology and Venereology, Department of Clinical Sciences, Lund University, SE-22184 Lund, Sweden

5Wound

Healing Center, Bispebjerg Hospital, Department of Biomedical Sciences, University of Copenhagen, DK-2100 Copenhagen, Denmark

KEY WORDS: antimicrobial, bacteria flocculation, infection confinement, laponite, membrane

ABSTRACT

The antimicrobial effects of laponite nanoparticles with or without loading of the antimicrobial peptide LL-37 was investigated along with their membrane interactions. The study combines data from ellipsometry, circular dichroism, fluorescence spectroscopy, particle size/ζ-potential measurements, and confocal microscopy. As a result of the net negative charge of laponite, loading of net positively charged LL-37 increases with increasing pH. The peptide was found to bind primarily to the outer surface of the laponite nanoparticles in a predominantly helical conformation, leading to charge reversal. Despite


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their net positive charge, peptide-loaded laponite nanoparticles did not kill Gram-negative

Escherichia coli bacteria or disrupt anionic model liposomes. They did however cause bacteria flocculation, originating from the interaction of laponite and bacterial lipopolysaccharide (LPS). Free LL-37, in contrast, is potently antimicrobial through membrane disruption, but does not induce bacterial aggregation in the concentration range investigated. Through LL-37-loading of laponite nanoparticles, the combined effects of bacterial flocculation and membrane lysis are observed. However, bacteria aggregation seems to be limited to Gram-negative bacteria, as laponite did not cause flocculation of Gram-positive Bacillus subtilis bacteria, nor did it bind to lipoteichoic acid from bacterial envelopes. Taken together, the present investigation reports on several novel phenomena by demonstrating that nanoparticle charge does not invariably control membrane destabilization, and by identifying the ability of anionic laponite nanoparticles to effectively flocculate Gram-negative bacteria through LPS binding. As demonstrated in cell experiments, such aggregation results in diminished of LPS-induced cell activation, thus outlining a promising approach for confinement of infection and inflammation caused by such pathogens.


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INTRODUCTION In the wake of increasing bacterial resistance against conventional antibiotics,1 there is a growing interest in alternative approaches for treating resistant bacterial infections. Nanoparticles are currently attracting considerable interest due to the comparatively low cost, good scalability, and broad versatility of such materials, but also due to presently undeveloped

bacterial

resistance.

Various

nanomaterials,

including

carbon

nanomaterials (graphene, nanotubes, fullerenes), metal nanoparticles (silver, gold, copper), metal oxides (e.g., titanium dioxide), quantum dots, mesoporous silica, and nanoclays, are attracting particular interest.2 Nanoparticles can kill bacteria through several mechanisms, including direct membrane lysis,3 oxidative degradation of bacterial lipids or proteins by reactive oxygen species (ROS),4 interference with bacterial proteins through release of metal ions,5 or heat-induced membrane destabilization caused by oscillating magnetic fields 6 or light.7 In addition to their antimicrobial properties, inorganic nanomaterials may be combined with both conventional antibiotics and with other antimicrobial agents.2 One class of alternative antimicrobial agents attracting current interest are antimicrobial peptides.8


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These peptides display broad-spectrum antimicrobial effects, primarily through membrane disruption. During the last few years in particular, various strategies have been reported for designing antimicrobial peptides with potency against antibiotic-resistant bacteria which selectively destabilize bacterial membranes, whilst maintaining low toxicity towards human cells.9 Much less attention, however, has been payed to delivery systems for antimicrobial peptides. This is somewhat surprising, as such peptides are relatively large, net positively charged and amphiphilic molecules. Thus, delivery systems may offer opportunities for improved efficacy and reduced toxicity.10 In the present investigation, attention is placed on nanoclays. Due to their layered structure, such materials have been suggested to be able to incorporate antimicrobial peptides between phyllosilicate layers, thus providing opportunities for sustained peptide release, protection from (infectionrelated) proteolytic degradation, and pH-dependent intracellular release,11-12 similar to those offered, e.g., by nanogels13 and mesoporous silica nanoparticles.14 Considering this, we previously investigated layered double hydroxides (LDHs) and found that membrane binding, extraction of anionic lipids, and membrane lysis, all increased with decreasing LDH particle size. While such nanoparticles were not found to provide any


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protection of LDH-bound LL-37 against proteolytic degradation, they were demonstrated to offer other functional advantages, notably through their ability to flocculate Escherichia

coli (E. coli), an effect again more pronounced for smaller particles.15 In the present study, we expand on this previous investigation by studying membrane interactions of laponite, a nanoclay consisting of negatively charged silicate sheets and cationic counterions. Previously, laponite has been used as delivery system for a range of different types of drugs, e.g., donepezil, dexamethasone, and ciprofloxacin.16-18 The relatively high counterion exchange capacity of laponite (53-63 meq/100 g) indicated the potential as a delivery system for cationic antimicrobial peptides.16,

18

In this study,

particular focus is placed on the role of laponite charge density on the binding/release of the antimicrobial peptide LL-37 (LLGDFFRKSKEKIGKEFKRIVQRIKDFLRNLVPRTES), interactions with cell membrane components, and antimicrobial effects of LL-37-loaded nanoparticles. A combination of ellipsometry, circular dichroism, fluorescence spectroscopy, particle size/ζ-potential measurements, and confocal microscopy were used to study the effect of laponite, LL-37, and peptide-loaded nanoparticles both regarding membrane lysis and bacterial aggregation. Furthermore, nanoparticle-induced


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aggregation is compared between Gram-negative and Gram-positive bacteria. These effects are correlated to laponite binding to bacterial lipopolysaccharide (LPS) and lipoteichoic acid (LTA), key anionic surface molecules for Gram-negative and Grampositive bacteria, respectively. As a result of being net negatively charged, laponite is demonstrated not to bind to bacterial-mimicking lipid membranes. It does, however, bind readily to LPS but not to similarly negatively charged LTA. Laponite also and selectively aggregates Gram-negative bacteria, despite Gram-negative and Gram-positive bacteria being characterized by similar negative electrostatic potentials. Together, these results show that particle interactions with bacterial membranes are more complex than dictated by simple electrostatics, and involve lipopolysaccharide and lipid components in different ways. We also demonstrate that laponite-induced LPS aggregation results in diminished LPS-induced cell activation. Such entrapment has previously been found to represent a key host defense mechanism, e.g., by thrombin-derived peptides in wounds.19-20 The present study extends on these previous findings by demonstrating that laponite may provide similar entrapment of bacteria and inflammatory bacterial LPS, thereby opening up possibilities for use as therapeutics in wound care for the spread of infection leading


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to uncontrolled systemic responses. Taken together, the present study therefore reports on several novel phenomena of potential interest for nanoparticle-induced confinement of infection and inflammation.

EXPERIMENTAL

Materials. LL-37 (LLGDFFRKSKEKIGKEFKRIVQRIKDFLRNLVPRTES, >95% purity, determined by mass spectrometry and HPLC) was obtained from Biopeptide Co. (San Diego, CA, USA). Laponite RD®, powder lot 0001731219, RD grade, was provided by BJØRN THORSEN A/S (Copenhagen, Denmark), calcined at 200 °C for 2 days, followed by extensive dialysis against Milli-Q water for one week. LPS from E. coli (0111:B4) and LTA from Staphylococcus aureus (S. aureus) was from Sigma (St. Louis, MO, USA). All other chemicals used were of analytical grade.

Microorganisms. E. coli ATCC 25299, a clinical strain, was obtained from the Department of Clinical Bacteriology, Lund University Hospital, while Bacillus subtilis (B.

subtilis) CCUG 163BT, a type strain, was obtained from the Department of Molecular Sciences, Swedish University of Agricultural Science.


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Liposome preparation and leakage assay. Negatively charged DOPE/DOPG (75/25 mol/mol) liposomes, frequently used as a bacterial membrane model,21 were employed to monitor membrane interactions of laponite nanoparticles, in the absence and presence of LL-37, under different ambient conditions. For this, dioleoylphosphatidylglycerol (DOPG) and dioleoylphosphatidylethanolamine (DOPE) were obtained from Avanti Polar Lipids (Alabaster, AL, USA). Lipids were dissolved in chloroform, followed by evaporation under nitrogen flow in order to form multilayers on a glass vial wall. Residual solvent was removed by vacuum for 24 h at room temperature. After this, 10 mM Tris, pH 7.4 (Sigma, St. Louis, MO, USA), was added together with 0.1 M carboxyfluorescein (CF) (Sigma, St. Louis, MO, USA). After hydration, the lipid mixture was exposed to eight freeze-thaw cycles, followed by 30 extrusions through polycarbonate filters (100 nm pore size) (Avestin, Ottawa, Canada) and removal of the un-encapsulated CF. CF release from the liposomes was followed by fluorescence at 515 nm, as described previously,22-23 using 0.8 mM Triton X-100 (Sigma-Aldrich, St. Louis, MO, USA) as positive control. Measurements were performed in triplicate at 25 °C.


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Ellipsometry. Peptide adsorption to nanoparticles was studied in situ by null ellipsometry at 532 nm, using an Optrel Multiskop (Optrel, Kleinmachnow, Germany), as described previously.22-23 The amount adsorbed was calculated using a refractive index increment of 0.154 cm3/g. Data were corrected for bulk refractive index variations induced by changes in temperature and excess electrolyte concentration. For studies of peptide binding to laponite nanoparticles, the latter were adsorbed onto poly-L-lysine-modified silica surfaces to a surface coverage of 1.54±0.23 mg/m2 prior to peptide addition. Poly-L-lysine-modification reduces peptide adsorption directly to the silica substrate by neutralizing its negative surface potential (-40 mV), allowing peptide binding to the adsorbed nanoparticles to be monitored without much background adsorption. For such studies, laponite particles were added to the poly-L-lysine coated wafer in Milli-Q water at a concentration of 50 ppm, followed by rinsing with 10 mM Tris, pH 7.4 (5 ml/min for 30 min). After stabilization of the signal for 15 min, LL-37 was added to a concentration of 1.0 µM, in turn followed by another addition to 2.0 µM, in all cases monitoring the adsorption for 30 min until stabilization.


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For studies of laponite interactions with LPS, methylated silica surfaces (contact angle 90°, surface potential -40 mV ) were coated with E. coli LPS (0.4 mg/ml) in 10 mM Tris, pH 7.4 over 2 h, followed by rinsing with 10 mM Tris, pH 7.4 for 30 min with flow and 15 min of stabilization, resulting in a LPS coverage of 1.48±0.38 mg/m2. After stabilization, 50 ppm laponite was adsorbed for 2 h and rinsed with 10 mM Tris, pH 7.4 for 30 min followed by rinsing with 10 mM Tris, pH 7.4, containing also 150 mM NaCl. For LTAcoated surfaces, LTA (0.2 mg/mL in 10 mM Tris, pH 7.4) was adsorbed at poly-L-lysinemodified silica surfaces. Subsequently, laponite was adsorbed from a 50 ppm solution for 1 h and rinsed for 15 min with 10 mM Tris, 150 mM NaCl, pH 7.4, in order to remove loosely attached nanoparticles. All ellipsometry measurements were performed at 25 °C in at least duplicate.

Particle size and ζ-potential measurements. Measurements of particle size (hydrodynamic diameter) and ζ-potential were performed by dynamic light scattering at 173°, using a Zetasizer Nano ZS (Malvern Instruments, Malvern, UK). Experiments were performed as a function of peptide concentration at a fixed laponite concentration of 50 ppm in 10 mM Tris, pH 7.4 or acetate pH 5.4 and 3.4 buffer. Laponite/peptide mixtures


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were incubated for 45 min in RT before measurement. Measurements were performed in triplicate at 25 °C.

CD spectroscopy. Circular dichroism spectroscopy (CD) experiments were performed using a Jasco J-810 Spectropolarimeter (Jasco, Easton, PA, USA). Measurements were performed under stirring with 50 ppm laponite dispersions in 10 mM Tris, pH 7.4 or acetate pH 5.4 and 3.4 buffer, in the absence and presence of LL-37. The background was subtracted to eliminate instrumental fluctuations between measurements. Measurements were performed in duplicate, and secondary structure quantification performed as described previously.22, 24

X-ray diffraction (XRD). Powder XRD measurements were performed with a PANalytical X’Pert Pro diffractometer (Eindhoven, Netherlands), using a Cu Kα radiation source (λ = 1.541 Å). Measurements were taken in reflection mode between 2 and 40° (2θ) at 45 kV and 40 mA. The interlayer spacing was determined using the d(001) value of the 001 peak. Samples were prepared by equilibrating 1000 ppm laponite with 400 µM LL-37 for either 24 or 72 hours. The samples were freeze-dried for 24 hours using an Epsilon 2-4


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LSC (Martin Christ Gefriertrocknungsanlagen GmbH, Germany) before duplicate XRD measurement.

Cryogenic transmission electron microscopy (cryoTEM). CryoTEM analyses were performed using a Tecnai G2 20 TWIN Transmission Electron Microscope (FEI, Hillsboro, OR, USA), operating at 80 kV in zero loss bright-field mode. To enhance and improve visualization, an underfocus of 1-2 m was used. Samples were equilibrated at 25C for 1 h, in the absence and presence of LL-37 and prepared by using a FEI Vitrobottm Mark IV. A small drop (~5 l) of sample was deposited on a copper grid covered with a perforated polymer film and thin evaporated carbon layers. Excess liquid was removed by blotting with filter paper. Immediately after blotting, samples were vitrified in liquid ethane, held just above its freezing point. Samples were kept below -165°C in a protected atmosphere during transfer and examination.

Confocal microscopy. Bacteria were plated onto a cover slide at 108 cfu/ml. Detection of live and dead bacteria was achieved by SYTO 9 and propidium iodide dye (LIVE/DEAD® BacLight™ Bacterial Viability Kits L7012, ThermoFischer Scientific, Waltham, MA, USA), respectively. For this, 1.5 µL of a 1:1 mixture of SYTO 9 and


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propidium iodide was added to 500 µL of sample. After incubation for 15 min at room temperature in 10 mM Tris, pH 7.4, samples were imaged with a 100x/1.25 oil objective using Zeiss 510 Confocal Microscope (Jena, Germany). Quantification of the fraction of live/dead bacteria, as well as of fraction of bacteria present in aggregates, was performed using ImageJ Software by comparing the fluorescence intensity and counting of bacteria respectively.

NF-B and AP-1 activation of human monocytes. THP1-XBlue-CD14 reporter monocytes (InvivoGen, Toulouse, France), were cultured and stimulated as described previously.25 In short, laponite (at the indicated final concentrations in 10 mM Tris, pH 7.4) with or without LPS (20 ng/mL final concentration) were incubated for 45 min at RT before being centrifuged at 14000g for 10 min. 10 mM Tris, pH 7.4, with or without LPS were used as positive and negative controls. Next, cells (1x106 cells/mL) were stimulated with the centrifuged supernatants. After incubation (18-20 h) at 37°C and 5% CO2, NF-κB and AP-1 activation was determined by incubating cell culture 14000 g supernatants with detection substrate (Quanti-BlueTM, InvivoGen, Toulouse, France) for 1-2 h, followed by quantification of cleaved substrate at an absorbance of 600 nm (n=3). In order to


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investigate cell toxicity effects, a lactic acid dehydrogenase assay was employed. For this, THP1 reporter cells were incubated as above for 18-20 h. Lactic acid dehydrogenase release was monitored by a lactic acid dehydrogenase-based in vitro toxicology assay kit (Sigma-Aldrich, St. Louis, MO, USA) according to manufacturer’s instructions.

RESULT LL-37 binding to laponite depends on charge contrast and occurs primarily at the outer surface As a first step, the size and charge of laponite nanoparticles was investigated. Due to the relatively poor electron contrast in this system, only particles oriented side-on are clearly seen in the cryoTEM images.26 Dynamic light scattering shows an average particle size of 45±5.4 nm in Milli-Q water, while some particle aggregation is observed after incubation for 45 min (RT) at an ionic strength of 10 mM at pH 3.4, 5.4, and 7.4, respectively (Fig. 1b), despite the weakly increasing negative surface charge with increasing pH. (Fig. 1c). On addition of net positively charged LL-37, the ζ-potential of the composite nanoparticles becomes less negatively charged, subsequently displaying charge reversal and a positive ζ-potential at saturation peptide binding. As a result of the


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pH-dependent laponite negative charge (Fig. 1c), more peptide is needed to neutralize the laponite charges with increasing pH, as demonstrated by the PZC being reached at a peptide concentration of 3.0, 7.9, and 11.5 µM at pH 3.4, 5.4, and 7.4, respectively. Furthermore, the limiting ζ-potential of the loaded particles after saturation peptide binding depends on pH, reaching 61±2, 36±3, and 22±1 mV at pH 3.4, 5.4, and 7.4, respectively (Fig. 1d), mirroring the pH-dependent net charge of LL-37.

Figure 1. (A) Representative cryoTEM images of 1000 ppm laponite dispersions in the absence (left) and presence (right) of 400 µM LL-37 in 10 mM Tris, pH 7.4. (B) Size


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distributions of laponite particles as a function of pH. Shown also in (C) and (D) are results on pH-dependent ζ-potential of laponite nanoparticles in the absence and presence of LL-37 at the indicated concentrations. Adsorption of LL-37 to laponite nanoparticles was then quantified by ellipsometry (Fig. 2). In agreement with the ζ-potential results, ellipsometry showed LL-37 binding to laponite nanoparticles to increase with increasing pH. Thus, at pH 3.4, where LL-37 carries a net charge of +11 and the bare laponite particles display a ζ-potential of -10±2 mV, LL-37 binding saturates at a density of 0.44±0.01 mg/m2, corresponding to ≈740 peptide molecules per laponite nanoparticle. In comparison, saturation is reached at ≈3300 peptide molecules/nanoparticle at pH 5.4, at which the net charge of LL-37 is +6.5 and the ζ-potential of laponite -28±2 mV. At pH 7.4, where the peptide net charge is +6 and the laponite ζ-potential is -26±4 mV, saturation binding is reached at ≈3800 peptide molecule per laponite nanoparticle (Fig. 2a). Together, this shows that LL-37 binding is largely dictated by the laponite negative charge. We also note that LL-37 bound to laponite does not desorb readily on rinsing with buffer (Fig. 2b).


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Figure 2. (A) LL-37 adsorption to laponite nanoparticles pre-adsorbed to poly-L-lysinecoated surfaces at pH 3.4, 5.4, and 7.4, keeping the ionic strength constant at 10 mM in all cases. Shown also is the LL-37 adsorption to the underlying poly-L-lysine-modified surface. (B) Time-resolved LL-37 adsorption to laponite in 10 mM Tris, pH 7.4. Rinsing with 10 mM Tris, pH 7.4 is indicated with an arrow.

We next performed XRD experiments to elucidate whether LL-37 is intercalated between the basal sheets of laponite, bound to the outside of the nanoparticles, or both. As shown in Supp. Info. Fig. SI1a, incubating laponite at 1000 ppm with 400 µM LL-37 over 24 hours does not result in any appreciable changes in the lattice spacing, indicating


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that binding of LL-37 occurs primarily on the outer surface of the laponite nanoparticles. Increasing the incubation time to 3 days resulted in a minor change in D-space from 13.11±0.17 to 13.77±0.42 Å (Supp. Info. Fig. SI1b). These results demonstrate that LL-37 intercalation into the laponite particle is a very slow process, occurring over a time scale of several days, if occurring at all. In contrast, X-ray

diffraction

studies

of

laponite

loading

with

the

cationic

surfactant

cetyltrimethylammonium bromide (CTAB) showed a change in layer spacing of 3.44 nm after incubation at 18.7 mM.27 The slow intercalation of LL-37 also means that the results above, as well as those below on bacterial aggregation and killing, correspond to a situation in which the overwhelming majority of the LL-37 molecules are bound to the outer surface of the laponite nanoparticles. Despite being restricted primarily to the outer surface of the laponite nanoparticles, LL-37 undergoes a pronounced conformational change on binding to the laponite nanoparticles. Thus, while displaying a largely disordered conformation in aqueous solution at pH 3.4-7.4, CD results show pronounced helix formation after laponite binding (Fig. 3a). The extent of helix conformation increased


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with increasing charge contrast between nanoparticle and peptide, thus being higher at pH 7.4 than at pH 3.4 (Fig. 3b).

Figure 3. (A) Representative CD spectra of LL-37 in the presence and absence of laponite at pH 3.4 and 7.4. (B) Helix content in of LL-37 in the absence and presence of laponite in pH 3.4 and 7.4. The measurements were performed at a constant ionic strength of 10 mM.

Membrane destabilization by laponite is limited As a result of the negative ζ-potential of bare laponite particles, they do not bind to strongly negatively charged (z≈-35 mV15) DOPE/DOPG membranes (results not shown). As a consequence of this, laponite nanoparticles do not induce leakage in DOPE/DOPG


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liposomes in the range 0.5 – 200 ppm for 10 mM Tris, pH 7.4 (Fig. 4a). Similar results were obtained at pH 5.4 (Supp. Info. Fig. SI2). Despite LL-37 being located at the outer surface of the composite laponite nanoparticles and despite the latter carrying a positive ζ-potential (Fig. 1d), membrane disruption induced by free LL-37 was strongly suppressed after laponite binding, indicating that the conformational restrictions of the particle-bound peptide molecules affected their membrane-destabilizing capacity detrimentally (Fig. 4a, b).

Figure 4. (A) Laponite-induced leakage of DOPE/DOPG liposomes in 10 mM Tris, pH 7.4. (B) Leakage of DOPE/DOPG liposomes in mixtures of laponite (50 ppm) and LL-37 at the indicated concentrations. Laponite and peptide was pre-mixed in 10 mM Tris, pH 7.4, 45


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min prior to addition to the liposome solution. For comparison, leakage induced by LL-37 alone is shown as well.

Laponite causes aggregation of Gram-negative bacteria, but not of Gram-positive To study the effects of laponite nanoparticles on bacteria in the absence and presence of LL-37, confocal microscopy experiments were performed for E. coli bacteria. As shown in Fig. 5a and 5b, laponite nanoparticles do not induce membrane defects in these bacteria, at least up to a particle concentration of 50 ppm. They do, however, induce bacterial aggregation, starting at 2.5 ppm and becoming prominent at 5 ppm, with large clusters of bacteria being formed. In contrast, well separated individual bacteria were observed prior to addition of laponite nanoparticles (Fig. 5a, c).


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Figure 5. (A) Representative confocal microscopy images of E. coli bacteria, using livedead assay (red, green, and DIC images overlaid), for 108 cfu/ml bacteria after treatment for 45 min with 0.5-5 ppm laponite in 10 mM Tris, pH 7.4. For all images, the size bar indicates 10 µm. Shown also in (B) and (C) are the fraction of live bacteria and the fraction of bacteria in aggregates, respectively.


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Due to these effects of the bare nanoparticles, we investigated whether it was possible to obtain combined membrane lysis and bacterial aggregation by combining laponite nanoparticles with LL-37. In order to do this, the LL-37 concentration was increased to 20 M, whilst maintaining a laponite concentration of 2.5 ppm. Based on the results shown in Fig. 6a and 6b, saturation binding is inferred to be reached at approximately 10 µM LL37, above which additional unbound peptide will be free to interact with the bacteria.


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Figure 6. (A) Representative confocal microscopy images of E. coli bacteria, using livedead assay (red, green and DIC images overlaid), in the presence of indicated concentrations of LL-37 with 2.5 ppm laponite in 10 mM Tris, pH 7.4 (incubation time 45 min). Shown also in (B) is the fraction of live bacteria. For all images, the size bar indicates 10 µm.


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From Fig. 6a and 6b, showing bacterial killing and aggregation versus peptide concentration, no significant difference in viability was observed in the absence and presence of the laponite nanoparticles, indicating that bacterial membrane rupturing is caused primarily by free LL-37, and that the presence of laponite nanoparticles does not provide any additional effect on membrane destruction. These findings are in agreement with the non-activity of the bare laponite nanoparticles on both bacteria and DOPE/DOPG liposome lysis (Fig. 4a, b and Fig. 5b). Also with regards to bacterial aggregation, there was no observable synergistic effect between laponite and LL-37, demonstrated by similar laponite-induced bacterial aggregation, both in the absence and presence of LL37 (Fig. 7a, b).


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Figure 7. (A) Representative confocal microscopy images of E. coli bacteria, using livedead assay (red, green, and DIC images overlaid), for 108 cfu/ml bacteria after treatment for 45 min with 5 ppm laponite in the presence and absence of 50 µM LL-37 in 10 mM Tris, pH 7.4. For all images, the size bar indicates 10 µm. Shown also in (B) and (C) is the fraction of live bacteria and the fraction of bacteria in aggregates, respectively. Included also are results for positive (live) and negative (dead) controls.


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In summary, Fig. 5a and 7a show that laponite nanoparticles do not kill E. coli bacteria but act as efficient bacterial flocculants. LL-37, in contrast, is effective in killing E. coli bacteria through membrane disruption, also in the presence of laponite (Fig. 7b), but does not induce any bacterial aggregation (Fig. 7c). By combining laponite and LL-37, both membrane lysis and aggregation can be obtained. The observed flocculation of E. coli bacteria by laponite is interesting due to the expected electrostatic repulsion arising from their similar net negative surface charge. Therefore, we next investigated if this phenomena is directly linked to the bacteria class by comparing these results to a Gram-positive bacteria, having similar effective ζ-potential as E. coli bacteria (Supp. Info. Fig. SI3). In order to address this, we performed similar confocal microscopy experiments as above for Gram-positive B. subtilis. From this, it was found that bare laponite does not induce membrane disruption, up to a laponite concentration of at least 5 ppm (Fig. 8a), analogous to the findings for Gram-negative E.

coli bacteria. In contrast to E. coli, however, laponite nanoparticles do not induce flocculation in B. subtilis (Supp. Info. Fig. SI4 and Fig. 8b).


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Figure 8. (A) Fraction of live B. subtilis bacteria (left) and bacteria in aggregates (right) as a function of laponite concentration. (B) Fraction of live bacteria (left) and bacteria in aggregates (right) after 45 min treatment with 5 ppm laponite in the presence and absence of LL-37.

Bacteria aggregation mirrors laponite binding to LPS, but not LTA


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Furthermore, laponite-induced flocculation was not observed for bacteria-mimicking DOPE/DOPG liposomes (Supp. Info. Fig. SI5), indicating that laponite-induced flocculation of E. coli bacteria is not driven by laponite-lipid interactions. Instead, laponiteinduced aggregation of E. coli bacteria could rather be induced by interaction of the laponite with bacterial lipopolysaccharides.


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Figure 9. Schematic illustration of the chemical structure of LPS (left) and LTAs (right). In both compounds, phosphate groups and lipid components have been highlighted in blue and purple, respectively.


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Therefore, laponite binding to LPS (dominating the outer membrane in Gram-negative bacteria, including E. coli)28 and LTA (a key lipopolysaccharide in Gram-positive bacteria, including B. subtilis)29 (Fig. 9) was investigated by ellipsometry. Results showed that laponite indeed binds readily to E. coli LPS, displaying saturation binding of 1.71±0.06 mg/m2 after rinsing with 10 mM Tris, pH 7.4, with additional 150 mM NaCl. In contrast, laponite binding to LTA was not observed (-0.036±0.008 mg/m2), in agreement to the limited nanoparticle-induced aggregation observed for B. subtilis (Fig. 10a, b, c).


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Figure 10. (A) Time-resolved laponite adsorption to LPS (left) and LTA (right) in 10 mM Tris, pH 7.4. Rinsing with 10 mM Tris, 150 mM NaCl, pH 7.4, is indicated with an arrow. (B) Limiting amount of laponite adsorbed to LSP and LTA. (C) Schematic illustration of laponite interactions with LPS and LTA in Gram-negative (left) and Gram-positive (right) bacteria, respectively.

Laponite-induced aggregation suppresses LPS-induced cell activation In order to investigate consequences of laponite nanoparticle co-aggregation with LPS, NF-κB/AP-1 activation experiments were conducted. NF-κB plays a key role in LPSinduced cell activation, subsequently resulting in the generation of pro-inflammatory cytokines, in turn playing a key role in sepsis. As seen in Fig. 11a, laponite, on its own, did not result in any appreciable NF-κB/AP-1 activation up to 100 ppm. In contrast, samples for which LPS/laponite aggregates had been removed by centrifugation showed a concentration-dependent decrease of NFκB/AP-1 activation, thus demonstrating a LPSscavenging role of laponite, which can lead to diminished LPS-induced cell activation. In agreement to this, lactate dehydrogenase experiments indicated that laponite attenuated


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LPS-induced lactate dehydrogenase-release, whereas laponite itself did not induce lactate dehydrogenase release (Fig. 11b).

Figure 11. (A) NF-κB/AP-1 activation and (B) LDH release by laponite at the indicated concentrations using THP1-XBlue-CD14 reporter monocytes in the absence and presence of LPS (20 ng/mL), as well for supernatants obtained after LPS-nanoparticle aggregation.

DISCUSSION In analogy to the ability of cationic nanoclays to bind, and intercalate, anionic molecules such as siRNA and anionic surfactants,30 anionic nanoclays have the capacity to bind cationic molecules both at their external surface and by intercalation within the basal


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layers. For laponite, examples of this include studies by Wang et al.,31 and Xiao et al.,32 who both reported that the surface potential of laponite becomes less negative on addition of doxorubicin. While binding to the external surface is straightforwardly determined by electrostatic and other interactions as for solid nanoparticles, the process of intercalation is more complex and depends on the system and experimental conditions. For example, Hamilton et al., investigated the mechanism of ciprofloxacin binding to laponite by monitoring structural changes using XRD. These authors did not observe any increase in the interlayer spacing with increasing concentration of ciprofloxacin, however, they suggest that as laponite is hydroscopic, hydration of the particle may mask such effects.18 On the other hand, Ghadiri et al., reported that tetracycline binding for 1 h resulted in an increase in the interlayer spacing in laponite in a pH-dependent manner.33 Similarly, Wang et al., loaded doxorubicin into laponite for 24 h and found this to be governed a two-step process of ion exchange and intercalation, based on XRD and ζ-potential measurements.31 Whilst laponite has so far not been investigated in relation to antimicrobial peptide incorporation, it is interesting to note that nanoclays have been previously reported to


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incorporate larger molecules. For example, Chen et al., investigated the effect of LDH size on siRNA delivery and found dsDNA and siRNA to intercalate more efficiently into small LDH nanoparticles, resulting in more efficient siRNA delivery outcome.30 For anionic nanoclays,

incorporation

of

both

hydrophobic

and

hydrophilic

polymers

into

montmorillonite has been studied. Thus, poly(oxyethylene) (POE) of different sizes (2000 – 6000 g/mol) changed the d-spacing from 12.4 Å (for the clay on its own) to 19.8 Å for the largest POE. The same was seen for poly(oxypropylene)diamine (POP-D) where molecular weights of 2000 and 4000 g/mol resulted in d-spacings of 58.0 and 92.0 Å, respectively.34-35 Considering this, it is reasonable that similar anionic nanoclays should be able to intercalate larger cationic molecules under certain circumstances. Despite this, our findings indicate that LL-37 does not intercalate into the laponite gallery, but instead is confined to the outer surface of the laponite nanoparticles. The reason for this is unclear, but possible contributing factors include the larger size of LL-37 compared to siRNA, and the presence of both electrostatic and non-electrostatic (notably hydrophobic) driving forces for laponite binding, which may result in kinetically arrested states at the first point of attachment (i.e., the outer surface).


36

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Depending on whether an antimicrobial agent is intercalated or bound to the external particle surface one would expect the membrane interactions and antimicrobial effects to be affected, together with the release rate of the incorporated antimicrobial. For example, Wang et al., found the antimicrobial effect of amoxicillin against S. aureus to be reduced after incorporation into laponite nanoparticles.36 Analogous effects have been previously reported also for mesoporous silica14 and polymer microgels13. For solid nanoparticles, which cannot incorporate the antimicrobial agent into the interior of the particle, release is frequently fast for small antimicrobials, contributing to the high antimicrobial effect. In addition, nanoparticles surface-coated by polycationic antimicrobial agents can bind to bacterial membranes and destabilize them through more general electrostatic interactions, effects displayed also by cationic nanoparticles not carrying any antimicrobials.37 However, as demonstrated previously for cationic LDH, membrane destabilization of antimicrobial peptide-coated nanoclays may be more complex than seen for cationic nanoparticles without internal structure.15 Demonstrating this, small LDH nanoparticles were found to display more efficient binding to bacteria-mimicking DOPE/DOPG membranes and extraction of anionic lipids than larger LDH nanoparticles.


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In the case of laponite, one would expect the LL-37-loaded particles to behave as other cationic and antimicrobial peptide-coated nanoparticles at first sight. In this context, we previously investigated membrane interactions of mesoporous and non-porous silica nanoparticles.14 As a result of pore incorporation, membrane interactions of LL-37-loaded mesoporous silica nanoparticles relies entirely on peptide release, effects which have also been observed for antimicrobial peptide-loaded anionic microgels.13 For non-porous silica nanoparticles, on the other hand, LL-37 was demonstrated to be arrested due to strong electrostatic interactions, hence membrane interactions of such peptide-loaded particles was entirely due to membrane binding and destabilization by the peptide-loaded particles. In the present case of laponite, LL-37 binding results in a limiting composite particle ζ-potential of ≈+20 mV at binding saturation at pH 7.4, which is slightly larger than that of the LL-37-coated solid silica nanoparticles previously investigated (z≈+10 mV).14 Hence, from an electrostatic perspective, the present observation of strongly reduced membrane interactions of laponite-bound LL-37, without interparticle intercalation, is unexpected.


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In the present investigation, the ability of laponite nanoparticles to flocculate bacteria is particularly interesting. This may offer an attractive approach for confinement of infection and resulting inflammation. A dispersed inflammatory response to infection can result in sepsis and it has been argued that some host defense peptides may be able to confine infections by such a mechanism.19-20 Localization of the inflammatory response by nanoparticles may similarly offer opportunities as therapeutics for avoidance of sepsis in severe infection. In this context, we also reported recently that cationic LDH nanoparticles are able to aggregate E. coli bacteria.15 As shown by the results in the present investigations for laponite, however, global electrostatics alone does not determine bacterial aggregation. Thus, despite carrying a net charge of ≈-25 mV on their own compared to an effective ζ-potential of ≈-35 mV for bacteria (Supp. Info. Fig. SI3), laponite nanoparticles are quite effective at inducing bacterial aggregation even at low particle concentrations. In contrast, no such aggregation was observed for anionic DOPE/DOPG liposomes with a very similar ζ-potential, demonstrating that the origin of this effect is not related to anionic phospholipid interactions. Instead, these results point to laponite interactions

with

non-lipid

bacterial

membrane

components,

notably

bacterial


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lipopolysaccharides. Indeed, despite the net negative charge of LPS, laponite binds readily to LPS, in agreement with the laponite-induced flocculation of E. coli bacteria observed. Interestingly, however, laponite does not bind to the similarly net negatively charged LTA, nor does it induce flocculation of Gram-positive B. subtilis bacteria. These results indicate that laponite binding to bacterial lipopolysaccharides depends rather sensitively on the nature of the latter. Thus, both LPS and LTA are net negatively charged polysaccharides containing phosphate and carboxyl residues, as well as a hydrophobic domain. However, the latter contains only two chains in LTA,29, 38 whereas LPS contains up to six lipid chains (Figure 9).39-41 As demonstrated previously by ANS fluorescence spectroscopy, the lipid chains in LPS are partly exposed to the aqueous surrounding42, and may, therefore, contribute to the hydrophobic driving force of laponite binding. In addition, LTA contains a substantial proportion of negatively charged phosphate groups in the repeating carbohydrate region29,

38

whereas LPS is dominated by uncharged

carbohydrate residues.39-41 This contributes to a repulsive electrostatic barrier to laponite adsorption, expected to be substantially larger for LTA than for LPS. For the bacterial flocculation experiments, yet another factor influencing laponite interaction with bacterial


40

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membranes is that LTA is incorporated within the peptidoglycan layer of Gram-positive bacteria29,

38

hence interaction with laponite nanoparticles is likely to be sterically

hindered. Taken together, these contributions result in laponite binding to LPS but not to LTA, as well as in laponite-induced flocculation of Gram-negative bacteria, but not of Gram-positive ones (Fig. 10c). Here, additional studies are clearly needed to determine the effect of charge group distributions within LPS and the laponite nanoparticles. This may affect the global electrostatics determining laponite-lipopolysaccharide interactions, particularly at intermediate ionic strength, where the Debye length is smaller than the particle size. Similarly, laponite being electrostatically anisotropic with a negative face and a partially positively charged rim, the latter may become relatively more important at intermediate to high ionic strength, as seen in other systems with ‘patchy’ charge distributions.43 Irrespective of the detailed mechanism, which requires further studies involving structural analysis of charge group distributions within LPS, LTA and laponite, it seems clear that nanoparticle binding results in bridging flocculation of Gram-negative bacteria. As clearly demonstrated in the present study, the resulting entrapment of LPS presents a powerful approach for suppressing LPS-induced cell activation. Considering


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the seemingly low cytotoxicity of laponite, such confinement potentially represents an approach for localizing inflammation and avoiding scattered and uncontrolled inflammatory responses, a hallmark of sepsis.

CONCLUSION Laponite nanoparticles are layered nanoclays, displaying increasingly negative ζpotential with increasing pH due to the dissociation of silanol groups. As a result of this, binding of the net positively charged antimicrobial peptide LL-37 to laponite increases with increasing pH. Despite the loaded nanoparticles displaying a distinctly positive ζpotential at saturation peptide binding, they do not disrupt E. coli bacteria or DOPE/DOPG liposomes. They do however, cause concentration-dependent aggregation of E. coli bacteria. As no effect was observed for model liposomes, the flocculation is not due to direct membrane interactions. Instead, bacterial flocculation correlates with laponite binding to LPS. Strikingly, laponite-induced flocculation was not observed for Grampositive B. subtilis bacteria, and laponite binding to LTA was limited. From confocal microscopy experiments, it was furthermore demonstrated that laponite nanoparticles do


42

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not kill E. coli bacteria, even after flocculation. LL-37, in contrast, is efficient at killing E.

coli bacteria through membrane disruption, but does not induce any bacterial aggregation within the concentrations required for antibacterial activity. Neither peptide nor nanoparticles dramatically influence the activity of the other. For the combined LL37/laponite system both bacterial killing, through membrane disruption, and aggregation was observed. In relation to previous literature on the effects of nanoparticle loading with antimicrobial peptides on resulting membrane interactions and antimicrobial effects,14-15 the present investigation provides new insight by: i) demonstrating that net positive charge does not automatically result in efficient membrane destabilization, and ii) identifying the capacity of anionic laponite nanoparticles to effectively flocculate Gramnegative bacteria (but not Gram-positive ones) through laponite binding to LPS. Furthermore, the finding that NF-κB/AP-1 activation in monocytes induced by LPS exposure is strongly suppressed after aggregation by laponite nanoparticles points to such nanoparticle-induced LPS and bacteria aggregation as a promising strategy for confinement of infection and inflammation. For the further elucidation of this, investigations of nanoclays of different size, providing a way to investigate effects of size-


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dependent van der Waals interactions, as well as the relative importance of side and edge charges in these electrostatically anisotropic systems, represent a logical next step.

ASSOCIATED CONTENT

Supporting Information. Results of XRD spectra of laponite in the absence and presence of LL-37, after equilibrating 1000 ppm laponite with 400 µM LL-37 for (A) 24 and (B) 72 hours (Fig. SI1b), laponite-induced leakage of DOPE/DOPG liposomes in pH 5.4 (Fig. SI2), ζ-potential of E. coli and B. subtilis in 10 mM Tris, pH 7.4 (Fig. SI3), confocal microscopy images of B. subtilis bacteria using live-dead assay (Fig. SI4), and as well as on laponite-induced flocculation of DOPE/DOPG liposomes (Fig. SI5), are available free of charge in Supporting Information.

AUTHOR INFORMATION

Corresponding Author [email protected]


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Author Contributions The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript.

Funding Sources

The research was funded by the Swedish Research Council (project 2016-05157, 201702341) and the LEO Foundation Center for Cutaneous Drug Delivery (project 2016-1101), Edvard Welanders Stiftelse (Hudfonden).

Notes

The authors declare no competing financial interest.

ACKNOWLEDGMENT

Lise-Britt Wahlberg and Dan Lundgren Nørgaard are gratefully acknowledged for technical support. We acknowledge the Core Facility for Integrated Microscopy, Faculty of Health and Medical Sciences, University of Copenhagen, for support with the cryoTEM experiments.


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Interaction of laponite with membrane components - consequences for

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