Dipalmitoyl-phosphatidylserine-filled cationic maltodextrin nanoparticles exhibit enhanced efficacy for cell entry and intracellular protein delivery in phagocytic THP-1 cells

Synthesis, labelling and lipidation of nanoparticles

The nanoparticles were synthesised as described previously [26]. The maltodextrin (Glucidex 6, Roquette, France) was dissolved in a Eq/L sodium hydroxide solution with magnetic stirring at room temperature, then epichlorohydrin and glycidyl trimethyl ammonium chloride (Sigma-Aldrich, France) were added, leading to the formation of a cationic hydrogel. The gel was then neutralised with acetic acid and crushed by a high-pressure homogeniser (LM20-30 microfluidizer, Microfluidics, France). The nanoparticles were purified from oligosaccharides, low-molecular weight reagents, and salts by tangential flow ultra-filtration (Akta Flux6, GE Healthcare, France) using a 300 kDa cut-off hollow fiber. The resulting purified cationic particles, called NP⁺, were the common intermediate for obtaining NPPG and NPPS. NPPGs are cationic and porous nanoparticles loaded with 1,2-dipalmitoyl-sn-glycero-3-PG (DPPG, Bertin bioreagent, France) while NPPS are loaded with 1,2-dipalmitoyl-sn-glycero-3-PS (DPPS, Bertin bioreagent, France). Whenever necessary, NP⁺ were labelled by fluorescein isothiocyanate (FITC, Thermo Fisher Scientific, France) according to Dombu et al. [5]. In a 0.1 M bicarbonate buffer with pH = 8.3, NP⁺ were labelled by 1% (w:w) of FITC for 12 h. Labelled NP⁺ were further purified by gel permeation chromatography on a PD-10 Sephadex desalting column (Sigma Aldrich, France). NP⁺ (or NP⁺-FITC) was loaded with 70% (w:w) of phospholipids (DPPG or DPPS) as previously described [26]. Phospholipids were dissolved in a 17% (w:v) aqueous solution of solutol HS15 (Sigma Aldrich, France), then mixed at 80°C with nanoparticles until complete incorporation. The resulting NPPGs or NPPSs were filtered through 0.2 µm polyethersulfone membrane.

Characterization of nanoparticles

The size and zeta potential of the nanoparticles were determined as previously described [26]. The hydrodynamic diameter (number) of the nanoparticles was measured by dynamic light scattering (Zetasizer nano ZS, Malvern UK) at 25°C, using 1 mg/mL nanoparticles in an aqueous solution of 15 mM NaCl. The zeta potential of the nanoparticles was measured by electrophoretic light scattering (Zetasizer nano ZS) at 25°C, using nanoparticles at 1 mg/mL in ultrapure water.

Cell culture

THP-1 cell (ATCC #TIB-202) is a monocyte isolated from peripheral blood from an acute monocytic leukaemia patient. Mycoplasma-free cells were maintained in Roswell Park Memorial Institute-1640 (RPMI-1640; Thermo Fisher Scientific) medium supplemented with 2 mM of l-glutamine, 10% heat-inactivated fetal bovine serum, 100 U/mL penicillin, and 100 μg/mL streptomycin (Thermo Fisher Scientific). Cells were incubated at 37°C in a humidified atmosphere with 5% CO₂. For cytometry analyses, THP-1 cells were seeded at a density of 2.0 × 10⁵ cells per well (1.9 cm²) in 24-well plates. For live-cell imaging, THP-1 cells were plated at a density of 0.4 × 10⁴ cells per cm² on 35 mm glass bottom microwell dishes (MatTek Corporation, Ashland, MA, USA). The cells were then differentiated into macrophages (macrophage-like THP-1 cells) with 40 ng/mL of phorbol 12-myristate 13-acetate (PMA, Sigma-Aldrich, France) for 48 h followed by a recovery period of 24 h without PMA.

Efferocytosis assay

The ability of macrophage-like THP-1 cells to phagocytose apoptotic cells was assessed using the Efferocytosis assay kit (Bertin bioreagent, France) according to the manufacturer’s instructions. Jurkat cells, an immortalised line of human T lymphocyte, were used as bait and were cultured using the same conditions as THP-1 cells. Briefly, THP-1 cells were labelled with CytoTell Blue™ and differentiated into macrophages. Then, Jurkat cells were labelled with carboxyfluorescein succinimidyl ester (CFSE) and treated with staurosporine for 6 h to induce apoptosis. Afterwards, apoptotic Jurkat cells were co-cultured for 18 h with macrophage-like THP-1 cells (cellular ratio 1:1) and cells were collected for flow cytometry analysis (Attune NxT, Thermo Fisher Scientific, France). A minimum of 5,000 cells were first selected based on their size and internal complexity (FSC/SSC), then the fluorescence intensities related to the cellular labels were determined using the following excitation/emission wavelengths (nm): CytoTell Blue: 405/450, CFSE: 488/525.

Confocal microscopy

An inverted confocal laser scanning microscope (STELLARIS 8, Leica Microsystems, Nanterre, France) equipped with a white light laser (440–790 nm), an 86X objective (NA = 1.20, water immersion, WD = 300 μm) were used in this study. Fluorescence signal was detected (500–550 nm) in photon counting mode through new generation Power HyD detectors (Power HyD-S, Silicon Multi-Pixel Photon Counter). Confocal imaging of nanoparticle-labelled living macrophage-like THP-1 cells was performed with 488 nm WLL line (2% AOTF) for FITC excitation. A full bold line Okolab chamber (Ottaviano, Italy) installed on the inverted microscope stand was used to keep the temperature at 37°C during image acquisition. Z-stack was obtained at a step of 300 nm with a total thickness of 14.1 μm. Bright-field image was acquired through activation of transmitted light detector. Images were processed through ImageJ for maximum projection and overlay.

TIRF microscopy

For live-cell imaging, macrophage-like THP-1 cells were exposed to 5 µg/mL of NPPS or NPPG for 0.5 or 2 h in RPMI-1640 medium. Afterward, cells were washed three times with the medium for 2 min each and kept in culture medium for observation. Total internal reflection fluorescence (TIRF) imaging (Leica Microsystems) of nanoparticle-internalised living macrophage-like THP-1 cells was performed with 488 nm line (25% AOTF) for FITC excitation and sCMOS camera (Hamamatsu, Massy, France) for fluorescence collection through 100× (NA = 1.46, oil immersion, WD = 0.17 µm) objective and GFP filter (excitation/bandwidth: 475/40 nm; dichroic cut: 495 nm; emission/bandwidth: 530/50 nm). TIRF microscopy was performed with a depth penetration of 150 nm and a single focus plane was acquired. Time-lapse was obtained at a frequency of one image every 2 s during 5 min. A full bold line Okolab chamber (PeCon, Erbach, Germany) installed on the inverted microscope stand was used to keep the temperature at 37°C during image acquisition. Images were processed through ImageJ for contrast adjustment and through Imaris (version 9.0.2, Oxford Instruments, Zurich, Switzerland) for nanoparticle counting and tracking. In particular, nanoparticles were automatically detected as single spherical spots through application of segmentation algorithm and thresholding to fluorescence signal while cell surface area was determined through a cell mask. Consequently, densities of nanoparticles were obtained as a ratio of nanoparticle number to cell surface (mm²).

Determination of nanoparticles cell entry by flow cytometry

Macrophage-like THP-1 cells were exposed to 10 µg/mL of FITC-labelled nanoparticles for 0.5, 1, 2, 4, or 6 h and collected using trypsin before flow cytometry analysis (Attune NxT, Thermo Fisher Scientific, France) using the following excitation/emission wavelengths (nm): FITC: 488/525. A minimum of 5,000 cells were first selected according to their size and internal complexity (FSC/SSC), then the fluorescence intensity related to the cellular content was determined. To compete with the PS receptors, cells were pretreated for 30 min with 2 mM phospho-L-serine (L-pSer) as previously described [27].

DPPS accessibility

To assess the accessibility of DPPS from the surrounding environment of the NPPS, annexin V, a PS binding protein, was used. FITC-labelled nanoparticles (5 µg) was mixed with 0.25, 1.25, or 2.5 µg of annexin V-AF647 (640943, Biolegend, France) in a calcium-containing buffer (10 mM HEPES, 140 mM NaCl, 2.5 mM CaCl₂, pH = 7.4) for 30 min at room temperature. The binding of annexin V to the nanoparticles was evaluated by flow cytometry (CytoFlex, Beckman Coulter, France). A minimum of 450,000 nanoparticles were first selected with FITC fluorescence intensity (excitation/emission (bandwidth) wavelengths: 488 nm/525(40) nm) in comparison with the unlabelled nanoparticles. The annexin V-AF647 (excitation/emission (bandwidth) wavelengths: 638 nm/660(10) nm) was analysed in the FITC positive nanoparticle region.

Association of proteins with nanoparticles

Bovine serum albumin (BSA, Sigma Aldrich, France) was used as a model protein. The loading of BSA in nanoparticles was performed by mixing both components in an aqueous solution at room temperature for 1 h. Different ratios (w:w) of nanoparticles/BSA were used from 1/0 to 1/5. The size and zeta potential were determined as described for the characterisation of the nanoparticles. The association of BSA with the nanoparticles was analysed by non-denaturing PAGE as previously described [10]. Gels were scanned and quantified using the ImageJ software, setting the 0% of association over the BSA alone input while 100% of association was set to the blank protein lane (NPPS or NPPG alone).

Intracellular delivery of proteins

BSA labelled with FITC (BSA-FITC, Sigma Aldrich, France) was associated with nanoparticles at a ratio (w:w) nanoparticle/protein of 3/1. Then, macrophage-like THP-1 cells were exposed to 10 µg/mL of the formulations for 0.5, 1, 2, 4 or 6 h and collected using trypsin before flow cytometry analysis (Attune NxT, Thermo Fisher Scientific) using the following excitation/emission wavelengths (nm): FITC: 488/525. A minimum of 5,000 cells were first selected according to their size and internal complexity (FSC/SSC) then the fluorescence intensity related to the BSA-FITC cellular content was determined.

Intracellular degradation of proteins

BSA-DQ (DQ™-green BSA, Sigma Aldrich, France) is a Bodipy-FL over-labelled protein used as a reporter of protein degradation. BSA-DQ was associated with nanoparticles at a nanoparticle/protein ratio (w:w) of 3/1. Then, macrophage-like THP-1 cells were exposed to 10 µg/mL of the formulations for 0.5, 1, 2, 4, or 6 h and collected using trypsin before flow cytometry analysis (Attune NxT, Thermo Fisher Scientific) using the following excitation/emission wavelengths (nm): 488/525. A minimum of 5,000 cells were first selected based on their size and internal complexity (FSC/SSC), then the fluorescence intensity related to the cellular content of Bodipy-FL-labelled peptides released by the degradation of BSA-DQ was determined. To assess the involvement of acid endosomes (i.e. lysosomes) in the process of BSA degradation delivered by nanoparticles, macrophage-like THP-1 cells were pretreated for 15 min with 50 mM of chloroquine diphosphate salt (Thermo Fisher Scientific), then exposed for 30 min to the BSA-DQ formulations before flow cytometry analysis. For longer time of exposure, a pH-sensitive fluorophore was used. BSA was labelled with pHrodo (pHrodo™ Red, SE, Thermo Fisher Scientific) according to the manufacturer’s instructions using 1% (w:w) of label. BSA-pHrodo was further purified by gel permeation chromatography on a PD-10 Sephadex desalting column (Sigma Aldrich, France) and the protein content was determined by the microBCA method (microBCA assay kit, Thermo Fisher Scientific). BSA-pHrodo was associated with nanoparticles at a nanoparticle/protein ratio (w:w) of 3/1. Then, macrophage-like THP-1 cells were exposed to 10 µg/mL of the formulations for 0.5, 1, 2, 4, or 6 h and collected using trypsin before flow cytometry analysis (Attune NxT, Thermo Fisher Scientific) using the following excitation/emission wavelengths (nm): 560/585. A minimum of 5,000 cells were first selected based on their size and internal complexity (FSC/SSC), then the fluorescence intensity related to the cellular content of low pH-activated BSA-pHrodo was determined.

Statistics

All experiments were done at least three times. The distribution of the quantitative variables was assessed by the Shapiro–Wilk test completed by the calculation of the residuals and the coefficients of Skewness and Kurtosis. Student’s tests and ANOVA with Tukey-Kramer correction were used to compare quantitative values. For kinetic analyses, multivariate regression with repeated measurements was used. All tests were performed with a risk alpha of 5% using the SAS Software (version 9.04.01M6P11072018). Only statistically significant differences are indicated in the figures.

Characterisation of the cationic maltodextrin nanoparticles and macrophage-like THP-1 cells

NPPSs and NPPGs are NP⁺ containing DPPS and DPPG, respectively. To strictly compare the effect of lipids on cell entry and protein delivery, a common batch of NP⁺ was used, which was filled with either DPPS or DPPG. The characterisation of nanoparticles using light scattering approaches revealed that NPPS and NPPG had similar (i) median hydrodynamic diameter (number size), (ii) interquartile range (from 0.178 to 0.269) of the polydispersity index (PDI), and (iii) zeta potential values (Table 1).

Since the polysaccharide matrix, the size, and the zeta potential of NPPS and NPPG were identical, the differential cellular effects could only be attributed to the associated lipid composition. The colloidal stability of nanoparticles was then determined weekly by measuring the size of the NPPS and NPPG stored in water at room temperature. No significant variations in the nanoparticle size could be detected during 28 weeks (Figure 1) demonstrating a robust colloidal stability of NPPS and NPPG over time in aqueous medium. Then, the ability of macrophage-like THP-1 cells to phagocyte apoptotic Jurkat cells via a PS-dependent efferocytosis mechanism was assessed through flow cytometry. Derived from monocytes, THP-1 cells were differentiated into macrophages after stimulation with PMA and labelled with CytoTell Blue. In parallel, immortalised human T lymphocyte Jurkat cells were labelled with CFSE and treated with staurosporine to induce apoptosis and exposure of the PS eat me signal on the outer plasma membrane. Both cell types were then co-cultured for 18 h and flow cytometry analyses revealed that more than 20% of Jurkat cells were engulfed by macrophage-like THP-1 cells (Figure 2). These data show that phagocytic macrophage-like THP-1 cells are competent for efferocytosis and expressed the relevant PS-receptors that can potentially be targeted by PS-containing nanoparticles.

Figure 1

Colloidal stability of NPPS and NPPG. The nanoparticles are stored for 28 weeks at room temperature in water. The hydrodynamic diameter of NPPS and NPPG are monitored by dynamic light scattering after sterile sampling of the nanoparticles. The graph represents the median and the interquartile of at least 6 samples. No significant difference is observed between NPPS and NPPG.

Figure 2

Macrophage-like THP-1 cells carry out efferocytosis. THP-1 are labelled with CytoTell Blue^® (left) while apoptotic Jurkat cells are labelled with CFSE (middle) as revealed by cytometry. THP-1 are co-cultured with apoptotic Jurkat (ratio 1:1) for 18 h (right), then cytometry analyses show that more than 20% of THP-1 cells have done efferocytosis. Grey: unlabelled cells, blue: CytoTell Blue^®-positive THP-1 cells, green: CFSE-positive Jurkat cells, red: CytoTell Blue^®-CFSE double positive THP-1 cells having captured apoptotic Jurkat cells.

Effect of associated lipid composition on cell entry efficacy

To study the impact of the lipid composition on the nanoparticle ability to penetrate macrophage-like THP-1 cells, two complementary strategies were used. Nanoparticle cell entry was first investigated in isolated living macrophage-like THP-1 cells with FITC-labelled NPPS and NPPG (5 µg/mL) through confocal and TIRF microscopies. On the cover glass, adherent macrophage-like THP-1 cells mainly showed rounded “fried egg”-like morphology with variable-sized basal lamellipodia (Figure 3a, white arrows). Confocal imaging revealed the presence of NPPS and NPPG in both lamellipodia and cytoplasm surrounding the nucleus (Figure 3a) suggesting that there is no specific site for nanoparticle entry in macrophage-like THP-1 cells. However, TIRF experiments indicated that NPPS and NPPG were mainly motionless in the lamellipodia in contrary to the upper cytoplasm. Consequently, for time resolution and standardisation purposes, images were only acquired at the cell body mid-plane and considered as a cell index for NPPS and NPPG TIRF evaluation. In this configuration, NPPS and NPPG were mainly distributed in the periphery of the cells after 0.5 h exposure and more widely found within the entire cytoplasm after 2 h of cell contact (Figure 3b and c). In particular, the density of NPPS and NPPG within the focus plan significantly increased over time of exposure (Figure 3d). In addition, NPPS density was slightly higher than NPPG (565 ± 47 vs 459 ± 45 particles/mm²) after 0.5 h-exposure, suggesting that NPPS may have entered the macrophage-like THP-1 cells faster than NPPG. Then, nanoparticle cell entry was studied in living cell population through flow cytometry. In microplates, macrophage-like THP-1 cells were exposed to 10 µg/mL of FITC-labelled nanoparticles up to 6 h and the overall cellular content of fluorescent nanoparticles was detected. Macrophage-like THP-1 cells loaded with NPPG showed a constant increase (linear slope) in intracellular NPPG content from 0.5 to 6 h (Figure 4). In contrast, NPPS cell entry displayed a strong initial start within the first 2 h followed by a progressive increase during the next 4 h (Figure 4). Although NPPG and NPPS loading exhibited parallel increase in slope from 2 to 6 h, the NPPS intensity signal is about two times higher than NPPG one. These data demonstrate that cell entry was faster and higher for NPPS than for NPPG as shown by the early boosted penetration of NPPS in macrophage-like THP-1 cells.

Figure 3

Internalised nanoparticle imaging in living macrophage-like THP-1 cells. (a) Macrophage-like THP-1 cells were exposed to 5 µg/mL of NPPS for 30 min and imaged by confocal microscopy (Green, FITC-labelled nanoparticles, maximum projection; Grey, bright field, white arrows indicated lamellipodia edges). Scale bar: 10 µm. (b and c) Macrophage-like THP-1 cells were exposed to 5 µg/mL of NPPS for 30 min (b) or 2 h (c) and imaged in a single focal plan at the mid-cell body level by TIRF microscopy. Scale bar: 10 µm. (d) Density of nanoparticles within a single focus plane of macrophage-like THP-1 cells through TIRF microscopy are represented as histograms expressing the mean value (±SEM) of a minimum of 7 cells. *p < 0.05; **p < 0.01.

Figure 4

Endocytosis of NPPS and NPPG by macrophage-like THP-1 cells. THP-1 cells are exposed to FITC-labelled-NPPS or -NPPG for 30 min to 6 h then analysed by cytometry to determine the cellular contents of the nanoparticles. The graph represents the mean values (±SEM) of at least three experiments. **p < 0.01.

Mechanism of nanoparticle cell entry

Since PS-receptors expressing phagocytic THP-1 macrophages can preferentially target DPPS-filled nanoparticles, the mechanism of nanoparticle cell entry was studied in more detail. First, L-pSer was used to compete the interactions between PS (associated with nanoparticles) and PS receptors (expressed by macrophages). Cells were pretreated for 30 min with 2 mM L-pSer followed by NPPS or NPPG exposure for 0.5 or 2 h. L-pSer had no effect on NPPG or NPPS cell entry whatever the duration of exposure was (Figure 5). These data suggest that the early boost of NPPS cell loading may not be related to PS saturable receptors. Second, annexin V that displays high affinity for PS was selected to get more insight into NPPS/PS receptor interactions. Therefore, a flow cytometry-based analysis was developed to monitor nanoparticle–protein interactions in various ratio of NPPS and annexin V, and NPPG was used as a control. Consequently, NP⁺ were first labelled with FITC and then filled with either DPPS or DPPG to produce FITC-labelled NPPG or NPPS, respectively. Nanoparticles were incubated for 30 min with Alexa Fluor 647-labelled annexin V before flow cytometry analysis. The nanoparticles were successively gated according to (i) size and structure (more than 450,000 events), (ii) FITC fluorescence, and (iii) Alexa Fluor 647 fluorescence. Different ratios of NPPS/annexin V and NPPG/annexin V (mass ratio: 1/0, 20/1, 4/1, and 2/1) were examined. With a 20/1 ratio, about 60% of NPPS or NPPG were associated with annexin V. With a 4/1 ratio, 95% of NPPS or NPPG were associated with annexin V, while all nanoparticles interacted with annexin V with a 2/1 ratio (Figure 6). These data indicate a clear interaction between nanoparticle and annexin V which do not depend on lipid composition.

Figure 5

Inhibition of nanoparticle endocytosis by competition of PS receptors using L-pSer in macrophage-like THP-1 cells. THP-1 cells are treated for 30 min with 2 mM of L-pSer and exposed to FITC-labelled-NPPS or -NPPG for 30 min (top) or 2 h (bottom), then analysed by cytometry to determine the cellular contents of the nanoparticles. The graph represents the mean value (±SEM) of at least three experiments. No statistically significant inhibition is observed using the L-pSer.

Figure 6

Accessibility of lipids associated with nanoparticles. Different mass ratios of FITC-labelled NP/annexin V-AF647 (Alexa Fluor 647) are formulated for 30 min and analysed by cytometry. Nanoparticles are first selected on their FITC fluorescence and then on the annexin V-AF647 content. No statistically significant difference is observed between NPPS and NPPG.

Study of protein delivery by nanoparticles

Since NPPS had better properties for macrophage-like THP-1 entry compared to NPPG, which are surprisingly not directly related to PS receptors, functional aspects of intracellular protein delivery by nanoparticles have been evaluated. For this, FITC-labelled BSA was incorporated into NPPS or NPPG, and protein association, size, and zeta potential were determined. Different mass ratios for nanoparticles/BSA (1/0, 5/1, 3/1, 1/1, 1/2, 1/3, 1/4, 1/5, and 0/1) were produced and formulations were run on non-denaturing PAGE. Under these conditions, nanoparticle-associated BSA could not migrate and gel analyses revealed only unassociated proteins. No unassociated BSA was observed for 5/1 or 3/1 ratios with NPPG or NPPS. At the ratio of 1/1, 98% of BSA was associated with NPPS and 83% with NPPG (Figure 7a and b). For further experiments, the 3/1 nanoparticles/BSA mass ratio, leading to a complete BSA association, was selected to compare the NPPS and NPPG formulations. For this ratio, BSA-NPPS and BSA-NPPG did not exhibit any significant change in size and zeta potentials compare to NPPS and NPPG (Figure 7c and d). The results demonstrate a similar association of BSA to NPPS and NPPG and suggest that BSA is integrated within the nanoparticles rather than on the surface. The capacity of NPPS and NPPG to deliver BSA into macrophage-like THP-1 cells was then investigated. Macrophage-like THP-1 cells were exposed to FITC-labelled BSA-NPPS or FITC-labelled BSA-NPPG from 0.5 to 6 h and cellular delivery of FITC-labelled BSA was analysed by cytometry. At the early time points (0.5 and 1 h), NPPS and NPPG delivered BSA in a similar manner. In contrast, from 2 to 6 h of exposure, NPPS delivered more BSA compared to NPPG (Figure 8). These data indicate that, in addition to cell entry properties, NPPS also presents enhanced capacity to deliver proteins in phagocytic cells compared to NPPG.

Figure 7

Association of protein by NPPS and NPPG. BSAs are used as a model protein. Different mass ratio of NPPS/BSA (a) or NPPG/BSA (b) are formulated for 30 min and run on non-denaturing polyacrylamide gel electrophoresis. Gels were imaged and quantified setting the 0% of association over the BSA input. Numbers on the gels represents the percentage of association of each formulation. The mass ratio nanoparticle/BSA: 3/1 is chosen since no free protein are detectable following silver nitrate staining of the gels. The relative size (c) and the relative zeta potential (d) of at least three formulations NPPS/BSA and NPPG/BSA (3/1) are represented on the graphs with the NPPS values set to 100%. No statistically significant difference is observed between NPPS and NPPG, with or without BSA.

Figure 8

Protein delivery by NPPS and NPPG in macrophage-like THP-1 cells. FITC-labelled BSA is formulated (mass ratio 3/1) with NPPS or NPPG and THP-1 cells are then exposed to the formulations before cytometry analyses of the cellular BSA-FITC content. The graph represents the median fluorescence intensity (±interquartile) of three experiments. *p < 0.05.

Characterisation of nanoparticle intracellular routes and protein processing

After nanoparticles are internalized by macrophage-like THP-1 cells, next is the intracellular routes that NPPS and NPPG follow which determine their destination within cellular compartments. As previously described, speed movements of nanoparticles were investigated in a single TIRF focal plan at the mid-cell body level. 5 min time-lapses through TIRF microscopy revealed that half of NPPS and NPPG were mobile irrespective of the duration of exposure i.e. 0.5 h or 2 h (Figure 9a). The average distance around 3 µm covered by mobile NPPS and NPPG in 5 min was not different between particles irrespective of the duration of exposure (Figure 9b and c). Interestingly, half of NPPS and NPPG were motionless regardless of the duration of exposure i.e. 0.5 h or 2 h. Therefore, the results indicate that there is no difference between NPPS and NPPG, as well on the static as on the dynamic profiles in a 5 min time-lapse experiment. Along the intracellular routes, proteins (i.e. antigens) are processed to induce or stimulate an immune response, and the first step generally involves protein degradation by two complementary means: enzymatic and pH-decrease-dependent degradation in acid endosomes and/or lysosomes. Whether NPPS better directs proteins to vesicular degradation compared to NPPG is a main question and two approaches have been developed to test this hypothesis. First, BSA-DQ, which fluoresces when degraded by vesicular enzymes, was associated with NPPS and NPPG. Degradation of BSA-DQ in macrophage-like THP-1 cells was quantified by flow cytometry after 0.5–6 h of exposure. No difference in BSA-DQ enzymatic degradation was observed between NPPS and NPPG at any time of the experiment (Figure 10a). Second, the potential role of lipids in pH-induced vesicular protein degradation was studied (i) through a 30 min pretreatment with chloroquine, which is described as an inhibitor of endosome acidification, or (ii) by using BSA-pHrodo, which is a pH-sensitive labelled protein that fluoresce at lower pH. No difference in protein degradation within chloroquine-pretreated macrophage-like THP-1 cells was observed when BSA was delivered by NPPS or NPPG (data not shown). BSA-pHrodo was formulated with NPPS and NPPG, exposed to macrophage-like THP-1 cells for 0.5–6 h and the acidification of BSA was then analysed by flow cytometry. Similarly, there was no difference in the kinetics of acidification when BSA was carried by NPPS or NPPG (Figure 10b). This suggests that, contrary to protein uptake and delivery, protein degradation through enzymatic or acidification processes was not dependent on the lipid composition of nanoparticles.

Figure 9

Characteristics of nanoparticle movement in living macrophage-like THP-1 cells through TIRF microscopy. (a) Histograms indicating the proportion of mobile nanoparticles in a single focus plane after 30 min and 2 h of exposition. No statistically significant difference is observed. (b) Illustration of NPPS tracking in a single focus plane after 30 min of exposition and using a 5 min time-lapse acquisition. Scale bar = 10 µm. Rainbow scale revealed time window of NPPS movements during time-lapse (blue, beginning; red, end). (c) Histograms indicating the average distance travelled by mobile nanoparticles within the cell. Each value represents the mean value (±SEM) of minimum five cells. No statistically significant difference is observed.

Figure 10

Characterization of nanoparticle-assisted protein degradation in macrophage-like THP-1 cells. NPPS or NPPG are formulated (mass ratio 3/1) with BSA-DQ (a) or with BSA-pH-rodo (b) to monitor the enzymatic or the pH-dependent degradation of proteins, respectively. THP-1 cells are exposed for 30 min to 6 h to the formulations and the cellular content of degraded BSA is measured by cytometry. The graphs represent the median fluorescence intensity (±interquartile) of three experiments. No statistically significant difference is observed between NPPG and NPPS.