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Effects of charged lipids on the physicochemical and biological properties of lipid–styrene maleic acid copolymer discoidal particles
BBA - Biomembranes 1862 (2020) 183209
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Effects of charged lipids on the physicochemical and biological properties of lipid–styrene maleic acid copolymer discoidal particles
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Masafumi Tanakaa,b, , Hisayasu Miyakea, Satoko Okaa, Shintaro Maedac, Kenji Iwasakic, Takahiro Mukaia a
Laboratory of Biophysical Chemistry, Kobe Pharmaceutical University, Kobe 658-8558, Japan Laboratory of Functional Molecular Chemistry, Kobe Pharmaceutical University, Kobe 658-8558, Japan c Laboratory of Protein Synthesis and Expression, Institute for Protein Research, Osaka University, Suita 565-0871, Japan b
A R T I C LE I N FO
A B S T R A C T
Keywords: Discoidal particles Charged lipids Cell uptake Cytotoxicity Styrene maleic acid copolymer
Styrene maleic acid copolymers (SMA) form discoidal lipid nanoparticles (lipid nanodisks) that mimic plasma high-density lipoproteins. We have previously prepared and characterized lipid nanodisks composed of SMA and the neutral phospholipid 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC). In the present study, we tested whether the surface charges can alter the physicochemical and biological properties of lipid–SMA discoidal particles. Unlike the case of DMPC alone, addition of saline to the buffer was necessary to induce the formation of lipid–SMA complexes containing either 1,2-dimyristoyl-sn-glycero-3-phosphoglycerol (DMPG) or 1,2-dimyristoyl-3-trimethylammonium-propane (DMTAP), with formation efficiency being dependent on the concentration of charged lipids. After purification, DMPG- or DMTAP-containing discoidal particles with an approximate size of 10 nm were obtained in a manner similar to DMPC alone. Although DMPG and DMTAP appeared to be similarly incorporated into the lipid nanodisks, the zeta potentials of both particles were comparable. That is, no significant differences were observed in the physicochemical properties between the lipid–SMA nanodisks. Compared to DMPC–SMA nanodisks, the uptake of DMPG or DMTAP-containing discoidal particles by RAW264 cells was increased for both particle types, whereas in MDA-MB-231 cells, only DMTAP-containing discoidal particle uptake was increased. In addition, fluorescence microscopy revealed that lipid–SMA nanodisks are localized adjacent to the plasma membrane of RAW264 cells but in MDA-MB-231 cells they accumulated in the center of the cell. Furthermore, these particles caused cytotoxicity in a cell-type dependent manner, with high toxicity in MDA-MB-231. These results raised the possibility that compositional alterations in lipid–SMA discoidal particles may modulate biological reactions in vivo.
1. Introduction High-density lipoprotein (HDL) is a natural carrier of lipophilic biomolecules such as cholesterol in plasma [1]. Nascent HDLs, lipidprotein complexes generated mainly through the efflux of cellular phospholipids and cholesterol by apolipoproteins, are discoidal in shape [2,3]. Such discoidal HDLs can be reconstituted with their major constituents, phospholipids and apolipoproteins [4]. Taking advantage of simple preparation methodologies, reconstituted HDL particles have been developed to apply as biocompatible delivery vehicles of drugs and/or imaging agents [5,6]. In general, apolipoproteins or their
fragmented peptides are utilized as materials for HDL reconstitution. However, the use of apolipoproteins usually obtained by extraction from animal plasma or a recombinant expression system is considered to be an important issue from the viewpoint of productivity, including safety and cost. Recently, it has been reported that a synthetic polymer, styrene maleic acid (SMA), can form discoidal HDL-like particles (SMA nanodisks) [7]. Heretofore, most studies on SMA nanodisks were performed to evaluate their application as a platform for structural analyses of membrane proteins [8,9]. To explore other uses of SMA nanodisks as novel delivery vehicles for drugs, radioactive compounds, or nucleic acids, we have launched an investigation to examine their
Abbreviations: DLS, dynamic light scattering; DMPC, 1,2-dimyristoyl-sn-glycero-3-phosphocholine; DMPG, 1,2-dimyristoyl-sn-glycero-3-phosphoglycerol; DMTAP, 1,2-dimyristoyl-3-trimethylammonium-propane; HDL, high-density lipoprotein; SMA, styrene maleic acid copolymer; TEM, transmission electron microscopy ⁎ Corresponding author at: Laboratory of Functional Molecular Chemistry, Kobe Pharmaceutical University, 4-19-1 Motoyamakita-machi, Higashinada-ku, Kobe 658-8558, Japan. E-mail address: [email protected] (M. Tanaka). https://doi.org/10.1016/j.bbamem.2020.183209 Received 10 November 2019; Received in revised form 8 January 2020; Accepted 27 January 2020 Available online 28 January 2020 0005-2736/ © 2020 Elsevier B.V. All rights reserved.
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2.4. Physicochemical characterization
physicochemical and biological properties. Charged lipids incorporated into lipoproteins can affect their functionality [10,11]. For example, anionic phospholipids in reconstituted lipoprotein particles have been shown to inhibit apolipoprotein E–lowdensity lipoprotein (LDL) receptor interactions [12]. Recently, it has been reported that discoidal HDL-like particles modified with GM1ganglioside, a glycosphingolipid covalently attached to an oligosaccharide with a sialic acid, play multifunctional roles in the combination therapies for Alzheimer's disease [13]. In contrast, cationic lipids, widely incorporated into liposomes for gene delivery [14], in reconstituted HDL have been shown to confer an ability to bind small interfering (si)RNA and the resulting complexes to possess target gene knockdown activity in a cultured cell model [15]. These results were obtained by lipid particles reconstituted with apolipoproteins, which are comprised of an amphipathic helical structure with positively charged residues at the polar-nonpolar interface and negatively charged residues at the center of the polar face [16]. Because SMA is a highly negatively charged polymer, influences from the incorporation of charged lipids on the physicochemical and biological properties of SMA nanodisks are likely to differ from those reconstituted with apolipoproteins. Previously, we prepared SMA nanodisks composed solely of neutral phospholipid and investigated their colloidal properties [17]. In the present study, to modify the physicochemical and biological properties of SMA nanodisks, we investigated the effects of the incorporation of anionic or cationic lipids.
Dynamic light scattering (DLS) technique was used to estimate the sizes and zeta potentials of the lipid–SMA complexes with a Zetasizer Nano ZS (Malvern, Worcestershire, UK). Transmission electron microscopy (TEM) observations of freshly prepared lipid–SMA complexes were performed on JEM-2200FS transmission electron microscope (JEOL, Tokyo, Japan) as described [17]. 2.5. Cell culture Murine macrophage cell line (RAW264) and human breast cancer cell line (MDA-MB-231) were used as testing cell lines. These cells were obtained from the RIKEN BRC through the National Bio-Resource Project of the MEXT, Japan, and American Type Culture Collection (ATCC, Manassas, VA), respectively. Cells were grown in a humidified incubator (5% CO2) at 37 °C in DMEM (for RAW264) or RPMI (for MDA-MB-231) supplemented with 10% fetal bovine serum (FBS), nonessential amino acids, penicillin and streptomycin. 2.6. Cell uptake
1,2-Dimyristoyl-sn-glycero-3-phosphocholine (DMPC) and 1,2-dimyristoyl-sn-glycero-3-phosphoglycerol (DMPG) were purchased from NOF Corporation (Tokyo, Japan). 1,2-Dimyristoyl-3-trimethylammonium-propane (DMTAP) was purchased from Avanti Polar Lipids (Alabaster, AL). Pre-hydrolyzed styrene–maleic anhydride copolymer 3:1 (SMA) was purchased from Sigma-Aldrich (St. Louis, MO). Lissamine™ Rhodamine B 1,2-dihexadecanoyl-sn-glycero-3-phosphoethanolamine (Rhodamine-PE) was obtained from Molecular Probes (Eugene, OR). 10 mM HEPES and HEPES-buffered saline (HBS) were used as buffer solutions (pH 7.4).
Cells were precultured in 24-well plates (1–3 × 105 cells/well) for two days. After washing, the cells were incubated with Rhodamine-PE labeled lipid particles (40 μg/mL of PC) at 37 °C for 30 or 120 min. After incubation, the cells were washed, dissolved in 0.2 M NaOH, and the concentration of cellular protein was measured. The fluorescence intensity of Rhodamine-PE was measured after centrifugation to remove insoluble matters. Fluorescence measurements were performed at room temperature on a Hitachi F-2500 spectrophotometer (Tokyo, Japan). Rhodamine-PE fluorescence spectra were recorded in a 4 × 4 mm cuvette from 560 to 650 nm at an excitation wavelength of 560 nm, and the intensities at 585 nm were chosen to quantitate cell uptake. Fluorescence microscopy was carried out for cells incubated with Rhodamine-PE labeled lipid particles (40 μg/mL of PC) on glass bottom dishes (Matsunami Glass, Kishiwada, Japan) at 37 °C for 120 min. Images were obtained using either a ZeissLSM700 confocal laser scanning system (Carl Zeiss, Oberkochen, Germany) or a BZ-X710 All-inOne Fluorescence Microscope (KEYENCE, Osaka, Japan). For confocal imaging, cells were treated with CellBrite™ Blue, as per the manufacturer's instructions (Biotium, Inc., Fremont, CA).
2.2. Phospholipid solubilization assay
2.7. Cytotoxicity assay
Lipids in organic solvents were dried under vacuum, dispersed in buffer with vigorous vortex mixing, and subjected to 5 freeze–thaw cycles. The kinetics of vesicle solubilization after the addition of SMA were monitored with right-angle light scattering using a Hitachi F-2500 spectrophotometer (Tokyo, Japan), as previously described [17]. Excitation and emission wavelengths were set at 600 nm. Vesicles (final PC concentrations of 50 μg/mL) were incubated at a PC to SMA weight ratio of 1:2 for 10 min at 24.6 °C.
The Cell Counting Kit-8 (CCK-8) assay was employed to test the cytotoxicity of lipid particles in RAW264 and MDA-MB-231 cells. Cells were precultured in 96-well plates (5 × 103 cells/well) for one day. After washing, lipid–SMA complexes (10, 25, 50, and 100 μM of PC) diluted in serum-free media were added to the cells and incubated at 37 °C for 6 h. After incubation, 20 μL of reagent was added and incubated for another 2 h for RAW264 cells or 4 h for MDA-MB-231 cells to quantify the living cells. Finally, the 96-well plates were analyzed on a microplate reader at 450 nm. Untreated cells were used as the 100% viability control.
2. Experimental procedures 2.1. Materials
2.3. Particle preparation 2.8. Concentration determination Lipid–SMA complexes composed of either DMPC alone or DMPC mixed with 20 mol% of DMPG or DMTAP were prepared as previously described [17]. Isolation of lipid–SMA complexes was performed on a Superdex 200 prep grade XK 16/600 column (GE Healthcare, Buckinghamshire, UK) at a flow rate of 1.0 mL/min controlled by a Biologic FPLC (Bio-Rad, Hercules, CA). Ultraviolet (UV) absorbance at 280 nm, possibly attributable to light scattering, was monitored. Fluorescence of a trace amount (1.0 mol%) of Rhodamine-PE incorporated into lipids was also measured.
DMPC concentrations were determined using a colorimetric enzymatic assay kit for choline from Wako Pure Chemicals (Osaka, Japan). Measuring the concentration of DMTAP accurately after particle preparation was technically challenging; therefore, the concentrations of lipid particles containing anionic or cationic lipids were normalized to DMPC instead. Cellular protein concentrations were determined by the Lowry procedure using bovine serum albumin (Bio-Rad, Hercules, CA) as a standard. SMA concentrations were determined as described [17]. 2
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Fig. 1. Time-dependent changes in light scattering intensity of lipid vesicles at 600 nm. (A and B) DMPC/DMPG vesicles in the absence (open) and presence (closed) of SMA (blue for 9:1, red for 8:2, and black for 7:3.). (C and D) DMPC/DMTAP vesicles in the absence (open) and presence (closed) of SMA (blue for 9:1, red for 8:2, and black for 7:3.). A and C; HEPES buffer, B and D: HBS.
profiles of DMPC/DMPG–SMA and DMPC/DMTAP–SMA complexes, respectively. Corresponding data of DMPC-SMA complexes are shown in [17]. Absorbance at 280 nm attributable to light scattering, largely overlapped with Rhodamine-PE fluorescence as a lipid marker. Lipid–SMA complexes were highly homogeneous only by mixing, which could be advantageous in practical applications. Interestingly, the peak was relatively broader when HBS was used instead of HEPES buffer as an elution buffer (data not shown). Therefore, the addition of salt is essential for the interaction of SMA with lipids to form complexes, however, once complexes are formed, they are rather stable in a buffer without saline. Fig. 3A and B show results of DLS analyses for DMPC/DMPG–SMA and DMPC/DMTAP–SMA particles, respectively. Corresponding data of DMPC-SMA complexes are shown in [17]. These particles remained stable for up to 5 days when kept at either 4 °C or 37 °C. Condensation process by ultrafiltration did not affect the particles size. The zeta potentials of DMPC/DMPG–SMA (−19.2 ± 2.4 mV) and DMPC/ DMTAP–SMA (−19.3 ± 2.4 mV) particles were similar to that of DMPC–SMA (−18.2 ± 2.1 mV) particles, despite the addition of charged lipids. It seems strange because zeta potentials of DMPC/DMPG and DMPC/DMTAP liposomes with the same molar ratio were approximately −24 mV and 44 mV, respectively. This is possibly due to the negative charge of SMA prevailing over the charge brought by the addition of the charged lipids. As expected, the particle morphologies of DMPC/DMPG–SMA and DMPC/DMTAP–SMA, as examined by TEM, were discoidal in shape (Fig. 3C and D), suggesting that particle integrity was not greatly influenced by the addition of charged lipids. Taken together, no significant differences were observed in terms of the physicochemical properties between lipid-SMA particles regardless of addition of charged lipids.
2.9. Statistical analysis For assessing the difference from the control group (DMPC–SMA complexes), Mann-Whitney test was carried out using GraphPad Prism 7 (GraphPad Software, San Diego, CA, USA). Significance level was set at p < 0.05. 3. Results 3.1. Solubilization assay SMA has been shown to induce the turbidity clearance of DMPC vesicles at 24.6 °C, the gel-to-liquid crystalline phase transition temperature of DMPC, due to solubilization by formation of discoidal particles [17]. Here, we examined whether the addition of charged lipids influences the solubilization behavior. Because DMPG possesses a transition temperature equivalent to DMPC [18], it was expected that solubilization of DMPC/DMPG vesicles after addition of SMA would also occur at 24.6 °C. However, no profound decrease in the turbidity of DMPC/DMPG vesicles by SMA was observed when HEPES buffer was used (Fig. 1A). In contrast, when HBS was used as a buffer, the addition of SMA caused turbidity clearance of DMPC/DMPG vesicles. In this case, the degree of the turbidity was not affected by the DMPG concentration (Fig. 1B). Similar results were obtained using phosphate buffer and phosphate buffered saline (Fig. S1), suggesting that the addition of salt is an important factor in the solubilization of DMPC/ DMPG vesicles, and distinct from vesicles composed solely of DMPC. Similarly, the turbidity derived from DMPC/DMTAP vesicles was almost unchanged by the addition of SMA in HEPES buffer (Fig. 1C). When HBS was used as a buffer, the turbidity clearance occurred in DMPC/DMTAP vesicles, but the degree of decrease in the turbidity was inversely correlated to the DMTAP concentration (Fig. 1D). These results suggest that electrostatic interactions between SMA and vesicles govern the efficiency of formation of these complexes.
3.3. Cell uptake of lipid–SMA complexes To evaluate the effect of charged lipids on cellular uptake, SMA nanodisks were incubated with RAW264 or MDA-MB-231 cells for 30 or 120 min (Fig. 4). Addition of either DMPG or DMTAP to lipid–SMA complexes enhanced their uptake into RAW264 cells (Fig. 4A). At 120 min, the uptake of DMPC/DMPG–SMA and DMPC/DMTAP–SMA particles was increased by approximately 2-fold compared to uptake of
3.2. Physicochemical properties of lipid–SMA complexes Lipid–SMA complexes formed in the presence of 20 mol% DMPG or DMTAP were further analyzed by gel filtration chromatography with HEPES buffer as the eluent. Fig. 2A and B show the gel filtration elution 3
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Fig. 2. Representative gel filtration profiles of lipid–SMA complexes monitored by UV absorbance at 280 nm (open) and Rhodamine-PE fluorescence (closed). (A) DMPC/DMPG (8:2) and (B) DMPC/DMTAP (8:2) using HEPES buffer as eluent.
periphery in RAW264 cells (Fig. 5A and B), whereas, in MDA-MB-231 cells, particles localized to the center of the cells (Fig. 5C and D). Confocal images clearly showed the internalization of SMA nanodisks by MDA-MB-231 cells (Fig. 5E). Collectively, the subcellular distribution pattern was cell-type dependent.
DMPC–SMA particles. Since lipid concentrations were normalized to DMPC, the differences in the absolute uptake may be larger when considering total lipids. In contrast, only DMTAP but not DMPG significantly enhanced the uptake by MDA-MB-231 cells (Fig. 4B). These results were similar to those observed for the uptake of charged liposomes by macrophages (J774) or cervical cancer cells (HeLa) [19]. However, considering that the physicochemical properties, particularly the zeta potential of these particles, were not affected by the addition of charged lipids, the observed results suggest that the partial (local) charges in the planar bilayer surface of the discoidal particles confers the cell selectivity. Fluorescence microscopy images were obtained to visualize the subcellular distribution of SMA nanodisks. Fig. 5 shows a representative microscopy image of RAW264 and MDA-MB-231 cells after incubation with SMA nanodisks. Although the differences in cell localization caused by compositional alterations in the lipid–SMA complexes were not specifically defined, all three SMA nanodisks seemed to locate to the
3.4. Cytotoxicity of lipid–SMA complexes To examine the effect of charged lipids in lipid–SMA complexes on cytotoxicity, SMA nanodisks were incubated with RAW264 and MDAMB-231 cells for 6 h (Fig. 6). Although SMA with lower molecular weight has been shown to induce no appreciable effect on cell viability [20,21], SMA nanodisks even composed solely of neutral DMPC caused cytotoxicity, especially in MDA-MB-231 cells. In RAW264 cells, DMPC/ DMPG–SMA particles showed slightly higher cytotoxicity when compared to DMPC/DMTAP–SMA particles. In MDA-MB-231 cells, no significant differences in cytotoxicity were observed between all three
Fig. 3. DLS measurements and TEM analyses of lipid–SMA complexes. (A and B) Size distribution of lipid–SMA complexes. (C and D) TEM images of lipid–SMA complexes stained with ammonium molybdate. A and C; DMPC/DMPG–SMA complexes, B and D; DMPC/DMTAP–SMA complexes. 4
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Fig. 4. Effects of the addition of anionic or cationic lipids on cellular uptake of SMA nanodisks. (A) RAW264 and (B) MDA-MB-231 cells. Open bars; DMPC–SMA complexes, closed bars; DMPC/DMPG–SMA complexes, hatched bars; DMPC/DMTAP–SMA complexes. *p < 0.05 vs DMPC–SMA complexes.
nanoparticles due to electrostatic interaction with cells. However, surface charges of nanoparticles optimum for the cellular uptake virtually depend on the type of the nanoparticles [31]. Especially, molecular modeling study suggests that cellular uptake mechanism of discoidal lipid nanoparticles is quite different from traditional nanoparticles [32]. The addition of positively charged DMTAP into SMA nanodisks enhanced its uptake by both RAW264 and MDA-MB-231 cells, despite DMPC/DMTAP–SMA particles showing an equivalent zeta potential to either DMPC–SMA or DMPC/DMPG–SMA. Since these three nanodisks are discoidal in shape and SMA molecules are assumed to wrap around the circumference of bilayer disks, the cells seemed to recognize the positive charge enriched in the planar lipid membranes of the nanodisks. In contrast, the uptake of SMA nanodisks with negatively charged DMPG was enhanced in RAW264 cells, but not in MDA-MB-231. Scavenger receptors are usually involved in the uptake of negatively charged liposomes by macrophages [33], thus, it is conceivable that the cellular uptake mechanism for DMPC/DMPG–SMA and DMPC/ DMTAP–SMA particles by RAW264 cells is distinct. Indeed, the amount of uptake for DMPC/DMPG–SMA particles was increased from 30 min to 2 h whereas that for DMPC/DMTAP–SMA particles remained constant, as shown in Fig. 4, although we did not fully quantify these changes. Cellular uptake of nanoparticles can also be modulated by the presence of serum [34,35]. When administered intravenously, the surface of nanoparticles would be covered with serum proteins, which may alter the cellular uptake process. Thus, the effect of the protein corona formed around the nanoparticles will need further consideration in future experiments. It is well known that positively charged nanoparticles can exhibit greater cytotoxicity [36]. Our preliminary experiments suggested that DMPC/DMTAP liposomes with a diameter of approximately 100 nm formed at a ratio of 8:2 caused more evident cytotoxic effects in RAW264 cells compared with DMPG-containing liposomes (data not shown). However, as shown above, the effect of DMTAP addition into SMA nanodisks on the cytotoxicity was negligible possibly because the presence of SMA maintained the net negative charge and might have compensated for the toxic effect. Such reduced cytotoxic activity even when in the presence of positively charged lipids can be an advantage of SMA nanodisks. The cytotoxicity results in the present study did not vary between particles, but rather varied in a cell dependent manner, similarly to the subcellular distribution pattern visualized by fluorescence microscopy. These results suggest that the potency of the cytotoxic effects may be correlated with the degree of internalization by the cells. The mechanistic basis for such cell specificity is still unclear. Because SMA is an artificial polymer with no postulated physiological functions, SMA nanodisks are assumed to be biologically inert. Thus, surface modifications may be required in future pharmacologic applications. In the present study, we proposed that the cellular uptake of SMA nanodisks is modulated simply by changes in the lipid composition. Ideally SMA nanodisks would specifically accumulate and exert toxic effects in cancer cells but not in normal cells, and this could be achieved by combining SMA with specifically designed lipid
SMA nanodisks, but notably, they all showed significantly greater cytotoxicity in MDA-MB-231 cells than in RAW264 cells. These results may be due to SMA nanodisks' internalization by MDA-MB-231 cells as observed by the fluorescence microscopy (Fig. 5E). In the cellular uptake assay, no apparent cell damage was observed even at the PC concentration of 40 μg/mL (corresponding to approximately 60 μM). This was likely due to the differences in the incubation times between the experiments, 120 min in the cell uptake assay versus 6 h for toxicity assay.
4. Discussion SMA required the presence of NaCl to solubilize vesicles with charged lipids, unlike in the case of vesicles composed of DMPC alone. The addition of NaCl facilitated the vesicle solubilization both in the presence of positively and negatively charged lipids. This result is consistent with the previous observation that there is no significant preference of SMA to solubilize homogeneous lipid mixtures [22]. We have previously reported that SMA was not able to solubilize 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC) vesicles in phosphate buffer [17]. We have, however, confirmed the turbidity clearance of POPC vesicles in phosphate buffer containing NaCl (Fig. S2). Indeed, Scheidelaar et al. demonstrated that increased salt concentrations could promote vesicle solubilization [23]. Even in the presence of NaCl, the degree of the clearance decreased with increasing amount of DMTAP, which may have been due to the shift of transition temperature upon DMTAP enrichment [24]. Alternatively, the capacity of nanodisks to accommodate positively charged lipids may be limited. However, 30% of DMTAP (by weight) could be incorporated into nanodisks composed of apoA-I and DMPC as the major lipids, in phosphate buffered saline [15]. Liposomes composed of phospholipids are rapidly trapped in the reticuloendothelial system (RES) when administered systemically. To evade RES uptake and prolong the blood circulation time, the inclusion of polyethylene glycol (PEG) is most commonly used [25]. However, PEGylation often hampers the functionality of the affinity ligand attached to liposomes [26] and also affects the uptake of liposomes containing phosphatidylserine by macrophages [27]. In contrast, SMA nanodisks mimicking nascent HDL are thought to be biocompatible and consequently it is anticipated that PEGylation may not be required. We have previously reported that POPC–SMA complexes are not recognized as foreign substances [28]. Furthermore, immunogenicity induced by PEGylation is one of the major issues in liposomal formulations. For example, acceleration of blood clearance (ABC phenomenon) upon repeated injection of PEGylated liposomes is typically observed [29]. Although there is a possibility that SMA, being an artificial polymer similar to PEG, could exert an immunogenic response, drug-conjugated SMA has so far demonstrated no immunogenicity, in a previous study [30]. Several reports have shown that positively charged nanoparticles can have a higher cellular uptake when compared to negatively charged 5
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Fig. 5. Fluorescence microscopy images of (A, B) RAW264 and (C, D) MDA-MB-231 cells treated with DMPC/DMPG–SMA complexes. A and C; differential interference contrast image, B and D; Rhodamine-PE fluorescence image. (E) Confocal fluorescence image of MDA-MB-231 cells showing internalization of DMPC–SMA complexes.
Fig. 6. Viability of (A) RAW264 and (B) MDA-MB231 cells incubated with SMA nanodisks for 6 h. DMPC–SMA complexes (closed circles), DMPC/ DMPG–SMA complexes (closed triangles), DMPC/ DMTAP–SMA complexes (closed square). No significant differences existed between all three SMA nanodisks, but significant differences were observed between RAW264 and MDA-MB-231 cells at 6 h.
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compositions. More studies will be required to further our understanding of the cellular uptake mechanisms and the immunogenic properties of SMA nanodisks for the development of novel cell specific and biocompatible nano carriers for use in pharmacological settings.
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Declarations of competing interest The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper. CRediT authorship contribution statement Masafumi Tanaka: Conceptualization, Writing - original draft. Hisayasu Miyake: Investigation. Satoko Oka: Investigation. Shintaro Maeda: Investigation. Kenji Iwasaki: Resources, Investigation. Takahiro Mukai: Supervision, Writing - review & editing. Acknowledgments TEM observation was supported by the Platform Project for Supporting in Drug Discovery and Life Science Research (Platform for Drug Discovery, Informatics, and Structural Life Science) from the Japan Agency for Medical Research and Development (AMED), Japan. We would like to thank Dr. Kohei Sano for the help with fluorescence microscopy imaging, Ms. Ayano Yuge for technical assistance, and Enago for the English language review. Appendix A. Supplementary data Supplementary data to this article can be found online at https:// doi.org/10.1016/j.bbamem.2020.183209. References [1] C.J. Fielding, P.E. Fielding, Molecular physiology of reverse cholesterol transport, J. Lipid Res. 36 (2) (1995) 211–228. [2] P.G. Frank, Y.L. Marcel, Apolipoprotein A-I: structure-function relationships, J. Lipid Res. 41 (6) (2000) 853–872. [3] K.A. Rye, P.J. Barter, Formation and metabolism of prebeta-migrating, lipid-poor apolipoprotein A-I, Arterioscler. Thromb. Vasc. Biol. 24 (3) (2004) 421–428. [4] C.G. Brouillette, G.M. Anantharamaiah, Structural models of human apolipoprotein A-I, Biochim. Biophys. Acta 1256 (2) (1995) 103–129. [5] R.O. Ryan, Nanodisks: hydrophobic drug delivery vehicles, Expert Opin. Drug Deliv. 5 (3) (2008) 343–351. [6] K.K. Ng, J.F. Lovell, G. Zheng, Lipoprotein-inspired nanoparticles for cancer theranostics, Acc. Chem. Res. 44 (10) (2011) 1105–1113. [7] M.C. Orwick, P.J. Judge, J. Procek, L. Lindholm, A. Graziadei, A. Engel, G. Grobner, A. Watts, Detergent-free formation and physicochemical characterization of nanosized lipid-polymer complexes: Lipodisq, Angew. Chem. Int. Ed. Engl. 51 (19) (2012) 4653–4657. [8] R. Zhang, I.D. Sahu, L. Liu, A. Osatuke, R.G. Comer, C. Dabney-Smith, G.A. Lorigan, Characterizing the structure of lipodisq nanoparticles for membrane protein spectroscopic studies, Biochim. Biophys. Acta 1848 (1 Pt B) (2015) 329–333. [9] J.M. Dorr, S. Scheidelaar, M.C. Koorengevel, J.J. Dominguez, M. Schafer, C.A. van Walree, J.A. Killian, The styrene-maleic acid copolymer: a versatile tool in membrane research, Eur. Biophys. J. 45 (1) (2016) 3–21. [10] S. Lund-Katz, P.M. Laplaud, M.C. Phillips, M.J. Chapman, Apolipoprotein B-100 conformation and particle surface charge in human LDL subspecies: implication for LDL receptor interaction, Biochemistry 37 (37) (1998) 12867–12874. [11] D.L. Sparks, C. Chatterjee, E. Young, J. Renwick, N.R. Pandey, Lipoprotein charge and vascular lipid metabolism, Chem. Phys. Lipids 154 (1) (2008) 1–6. [12] T. Yamamoto, R.O. Ryan, Anionic phospholipids inhibit apolipoprotein E–lowdensity lipoprotein receptor interactions, Biochem. Biophys. Res. Commun. 354 (3)
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Effects of charged lipids on the physicochemical and biological properties of lipid–styrene maleic acid copolymer discoidal particles
T
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Masafumi Tanakaa,b, , Hisayasu Miyakea, Satoko Okaa, Shintaro Maedac, Kenji Iwasakic, Takahiro Mukaia a
Laboratory of Biophysical Chemistry, Kobe Pharmaceutical University, Kobe 658-8558, Japan Laboratory of Functional Molecular Chemistry, Kobe Pharmaceutical University, Kobe 658-8558, Japan c Laboratory of Protein Synthesis and Expression, Institute for Protein Research, Osaka University, Suita 565-0871, Japan b
A R T I C LE I N FO
A B S T R A C T
Keywords: Discoidal particles Charged lipids Cell uptake Cytotoxicity Styrene maleic acid copolymer
Styrene maleic acid copolymers (SMA) form discoidal lipid nanoparticles (lipid nanodisks) that mimic plasma high-density lipoproteins. We have previously prepared and characterized lipid nanodisks composed of SMA and the neutral phospholipid 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC). In the present study, we tested whether the surface charges can alter the physicochemical and biological properties of lipid–SMA discoidal particles. Unlike the case of DMPC alone, addition of saline to the buffer was necessary to induce the formation of lipid–SMA complexes containing either 1,2-dimyristoyl-sn-glycero-3-phosphoglycerol (DMPG) or 1,2-dimyristoyl-3-trimethylammonium-propane (DMTAP), with formation efficiency being dependent on the concentration of charged lipids. After purification, DMPG- or DMTAP-containing discoidal particles with an approximate size of 10 nm were obtained in a manner similar to DMPC alone. Although DMPG and DMTAP appeared to be similarly incorporated into the lipid nanodisks, the zeta potentials of both particles were comparable. That is, no significant differences were observed in the physicochemical properties between the lipid–SMA nanodisks. Compared to DMPC–SMA nanodisks, the uptake of DMPG or DMTAP-containing discoidal particles by RAW264 cells was increased for both particle types, whereas in MDA-MB-231 cells, only DMTAP-containing discoidal particle uptake was increased. In addition, fluorescence microscopy revealed that lipid–SMA nanodisks are localized adjacent to the plasma membrane of RAW264 cells but in MDA-MB-231 cells they accumulated in the center of the cell. Furthermore, these particles caused cytotoxicity in a cell-type dependent manner, with high toxicity in MDA-MB-231. These results raised the possibility that compositional alterations in lipid–SMA discoidal particles may modulate biological reactions in vivo.
1. Introduction High-density lipoprotein (HDL) is a natural carrier of lipophilic biomolecules such as cholesterol in plasma [1]. Nascent HDLs, lipidprotein complexes generated mainly through the efflux of cellular phospholipids and cholesterol by apolipoproteins, are discoidal in shape [2,3]. Such discoidal HDLs can be reconstituted with their major constituents, phospholipids and apolipoproteins [4]. Taking advantage of simple preparation methodologies, reconstituted HDL particles have been developed to apply as biocompatible delivery vehicles of drugs and/or imaging agents [5,6]. In general, apolipoproteins or their
fragmented peptides are utilized as materials for HDL reconstitution. However, the use of apolipoproteins usually obtained by extraction from animal plasma or a recombinant expression system is considered to be an important issue from the viewpoint of productivity, including safety and cost. Recently, it has been reported that a synthetic polymer, styrene maleic acid (SMA), can form discoidal HDL-like particles (SMA nanodisks) [7]. Heretofore, most studies on SMA nanodisks were performed to evaluate their application as a platform for structural analyses of membrane proteins [8,9]. To explore other uses of SMA nanodisks as novel delivery vehicles for drugs, radioactive compounds, or nucleic acids, we have launched an investigation to examine their
Abbreviations: DLS, dynamic light scattering; DMPC, 1,2-dimyristoyl-sn-glycero-3-phosphocholine; DMPG, 1,2-dimyristoyl-sn-glycero-3-phosphoglycerol; DMTAP, 1,2-dimyristoyl-3-trimethylammonium-propane; HDL, high-density lipoprotein; SMA, styrene maleic acid copolymer; TEM, transmission electron microscopy ⁎ Corresponding author at: Laboratory of Functional Molecular Chemistry, Kobe Pharmaceutical University, 4-19-1 Motoyamakita-machi, Higashinada-ku, Kobe 658-8558, Japan. E-mail address: [email protected] (M. Tanaka). https://doi.org/10.1016/j.bbamem.2020.183209 Received 10 November 2019; Received in revised form 8 January 2020; Accepted 27 January 2020 Available online 28 January 2020 0005-2736/ © 2020 Elsevier B.V. All rights reserved.
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2.4. Physicochemical characterization
physicochemical and biological properties. Charged lipids incorporated into lipoproteins can affect their functionality [10,11]. For example, anionic phospholipids in reconstituted lipoprotein particles have been shown to inhibit apolipoprotein E–lowdensity lipoprotein (LDL) receptor interactions [12]. Recently, it has been reported that discoidal HDL-like particles modified with GM1ganglioside, a glycosphingolipid covalently attached to an oligosaccharide with a sialic acid, play multifunctional roles in the combination therapies for Alzheimer's disease [13]. In contrast, cationic lipids, widely incorporated into liposomes for gene delivery [14], in reconstituted HDL have been shown to confer an ability to bind small interfering (si)RNA and the resulting complexes to possess target gene knockdown activity in a cultured cell model [15]. These results were obtained by lipid particles reconstituted with apolipoproteins, which are comprised of an amphipathic helical structure with positively charged residues at the polar-nonpolar interface and negatively charged residues at the center of the polar face [16]. Because SMA is a highly negatively charged polymer, influences from the incorporation of charged lipids on the physicochemical and biological properties of SMA nanodisks are likely to differ from those reconstituted with apolipoproteins. Previously, we prepared SMA nanodisks composed solely of neutral phospholipid and investigated their colloidal properties [17]. In the present study, to modify the physicochemical and biological properties of SMA nanodisks, we investigated the effects of the incorporation of anionic or cationic lipids.
Dynamic light scattering (DLS) technique was used to estimate the sizes and zeta potentials of the lipid–SMA complexes with a Zetasizer Nano ZS (Malvern, Worcestershire, UK). Transmission electron microscopy (TEM) observations of freshly prepared lipid–SMA complexes were performed on JEM-2200FS transmission electron microscope (JEOL, Tokyo, Japan) as described [17]. 2.5. Cell culture Murine macrophage cell line (RAW264) and human breast cancer cell line (MDA-MB-231) were used as testing cell lines. These cells were obtained from the RIKEN BRC through the National Bio-Resource Project of the MEXT, Japan, and American Type Culture Collection (ATCC, Manassas, VA), respectively. Cells were grown in a humidified incubator (5% CO2) at 37 °C in DMEM (for RAW264) or RPMI (for MDA-MB-231) supplemented with 10% fetal bovine serum (FBS), nonessential amino acids, penicillin and streptomycin. 2.6. Cell uptake
1,2-Dimyristoyl-sn-glycero-3-phosphocholine (DMPC) and 1,2-dimyristoyl-sn-glycero-3-phosphoglycerol (DMPG) were purchased from NOF Corporation (Tokyo, Japan). 1,2-Dimyristoyl-3-trimethylammonium-propane (DMTAP) was purchased from Avanti Polar Lipids (Alabaster, AL). Pre-hydrolyzed styrene–maleic anhydride copolymer 3:1 (SMA) was purchased from Sigma-Aldrich (St. Louis, MO). Lissamine™ Rhodamine B 1,2-dihexadecanoyl-sn-glycero-3-phosphoethanolamine (Rhodamine-PE) was obtained from Molecular Probes (Eugene, OR). 10 mM HEPES and HEPES-buffered saline (HBS) were used as buffer solutions (pH 7.4).
Cells were precultured in 24-well plates (1–3 × 105 cells/well) for two days. After washing, the cells were incubated with Rhodamine-PE labeled lipid particles (40 μg/mL of PC) at 37 °C for 30 or 120 min. After incubation, the cells were washed, dissolved in 0.2 M NaOH, and the concentration of cellular protein was measured. The fluorescence intensity of Rhodamine-PE was measured after centrifugation to remove insoluble matters. Fluorescence measurements were performed at room temperature on a Hitachi F-2500 spectrophotometer (Tokyo, Japan). Rhodamine-PE fluorescence spectra were recorded in a 4 × 4 mm cuvette from 560 to 650 nm at an excitation wavelength of 560 nm, and the intensities at 585 nm were chosen to quantitate cell uptake. Fluorescence microscopy was carried out for cells incubated with Rhodamine-PE labeled lipid particles (40 μg/mL of PC) on glass bottom dishes (Matsunami Glass, Kishiwada, Japan) at 37 °C for 120 min. Images were obtained using either a ZeissLSM700 confocal laser scanning system (Carl Zeiss, Oberkochen, Germany) or a BZ-X710 All-inOne Fluorescence Microscope (KEYENCE, Osaka, Japan). For confocal imaging, cells were treated with CellBrite™ Blue, as per the manufacturer's instructions (Biotium, Inc., Fremont, CA).
2.2. Phospholipid solubilization assay
2.7. Cytotoxicity assay
Lipids in organic solvents were dried under vacuum, dispersed in buffer with vigorous vortex mixing, and subjected to 5 freeze–thaw cycles. The kinetics of vesicle solubilization after the addition of SMA were monitored with right-angle light scattering using a Hitachi F-2500 spectrophotometer (Tokyo, Japan), as previously described [17]. Excitation and emission wavelengths were set at 600 nm. Vesicles (final PC concentrations of 50 μg/mL) were incubated at a PC to SMA weight ratio of 1:2 for 10 min at 24.6 °C.
The Cell Counting Kit-8 (CCK-8) assay was employed to test the cytotoxicity of lipid particles in RAW264 and MDA-MB-231 cells. Cells were precultured in 96-well plates (5 × 103 cells/well) for one day. After washing, lipid–SMA complexes (10, 25, 50, and 100 μM of PC) diluted in serum-free media were added to the cells and incubated at 37 °C for 6 h. After incubation, 20 μL of reagent was added and incubated for another 2 h for RAW264 cells or 4 h for MDA-MB-231 cells to quantify the living cells. Finally, the 96-well plates were analyzed on a microplate reader at 450 nm. Untreated cells were used as the 100% viability control.
2. Experimental procedures 2.1. Materials
2.3. Particle preparation 2.8. Concentration determination Lipid–SMA complexes composed of either DMPC alone or DMPC mixed with 20 mol% of DMPG or DMTAP were prepared as previously described [17]. Isolation of lipid–SMA complexes was performed on a Superdex 200 prep grade XK 16/600 column (GE Healthcare, Buckinghamshire, UK) at a flow rate of 1.0 mL/min controlled by a Biologic FPLC (Bio-Rad, Hercules, CA). Ultraviolet (UV) absorbance at 280 nm, possibly attributable to light scattering, was monitored. Fluorescence of a trace amount (1.0 mol%) of Rhodamine-PE incorporated into lipids was also measured.
DMPC concentrations were determined using a colorimetric enzymatic assay kit for choline from Wako Pure Chemicals (Osaka, Japan). Measuring the concentration of DMTAP accurately after particle preparation was technically challenging; therefore, the concentrations of lipid particles containing anionic or cationic lipids were normalized to DMPC instead. Cellular protein concentrations were determined by the Lowry procedure using bovine serum albumin (Bio-Rad, Hercules, CA) as a standard. SMA concentrations were determined as described [17]. 2
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Fig. 1. Time-dependent changes in light scattering intensity of lipid vesicles at 600 nm. (A and B) DMPC/DMPG vesicles in the absence (open) and presence (closed) of SMA (blue for 9:1, red for 8:2, and black for 7:3.). (C and D) DMPC/DMTAP vesicles in the absence (open) and presence (closed) of SMA (blue for 9:1, red for 8:2, and black for 7:3.). A and C; HEPES buffer, B and D: HBS.
profiles of DMPC/DMPG–SMA and DMPC/DMTAP–SMA complexes, respectively. Corresponding data of DMPC-SMA complexes are shown in [17]. Absorbance at 280 nm attributable to light scattering, largely overlapped with Rhodamine-PE fluorescence as a lipid marker. Lipid–SMA complexes were highly homogeneous only by mixing, which could be advantageous in practical applications. Interestingly, the peak was relatively broader when HBS was used instead of HEPES buffer as an elution buffer (data not shown). Therefore, the addition of salt is essential for the interaction of SMA with lipids to form complexes, however, once complexes are formed, they are rather stable in a buffer without saline. Fig. 3A and B show results of DLS analyses for DMPC/DMPG–SMA and DMPC/DMTAP–SMA particles, respectively. Corresponding data of DMPC-SMA complexes are shown in [17]. These particles remained stable for up to 5 days when kept at either 4 °C or 37 °C. Condensation process by ultrafiltration did not affect the particles size. The zeta potentials of DMPC/DMPG–SMA (−19.2 ± 2.4 mV) and DMPC/ DMTAP–SMA (−19.3 ± 2.4 mV) particles were similar to that of DMPC–SMA (−18.2 ± 2.1 mV) particles, despite the addition of charged lipids. It seems strange because zeta potentials of DMPC/DMPG and DMPC/DMTAP liposomes with the same molar ratio were approximately −24 mV and 44 mV, respectively. This is possibly due to the negative charge of SMA prevailing over the charge brought by the addition of the charged lipids. As expected, the particle morphologies of DMPC/DMPG–SMA and DMPC/DMTAP–SMA, as examined by TEM, were discoidal in shape (Fig. 3C and D), suggesting that particle integrity was not greatly influenced by the addition of charged lipids. Taken together, no significant differences were observed in terms of the physicochemical properties between lipid-SMA particles regardless of addition of charged lipids.
2.9. Statistical analysis For assessing the difference from the control group (DMPC–SMA complexes), Mann-Whitney test was carried out using GraphPad Prism 7 (GraphPad Software, San Diego, CA, USA). Significance level was set at p < 0.05. 3. Results 3.1. Solubilization assay SMA has been shown to induce the turbidity clearance of DMPC vesicles at 24.6 °C, the gel-to-liquid crystalline phase transition temperature of DMPC, due to solubilization by formation of discoidal particles [17]. Here, we examined whether the addition of charged lipids influences the solubilization behavior. Because DMPG possesses a transition temperature equivalent to DMPC [18], it was expected that solubilization of DMPC/DMPG vesicles after addition of SMA would also occur at 24.6 °C. However, no profound decrease in the turbidity of DMPC/DMPG vesicles by SMA was observed when HEPES buffer was used (Fig. 1A). In contrast, when HBS was used as a buffer, the addition of SMA caused turbidity clearance of DMPC/DMPG vesicles. In this case, the degree of the turbidity was not affected by the DMPG concentration (Fig. 1B). Similar results were obtained using phosphate buffer and phosphate buffered saline (Fig. S1), suggesting that the addition of salt is an important factor in the solubilization of DMPC/ DMPG vesicles, and distinct from vesicles composed solely of DMPC. Similarly, the turbidity derived from DMPC/DMTAP vesicles was almost unchanged by the addition of SMA in HEPES buffer (Fig. 1C). When HBS was used as a buffer, the turbidity clearance occurred in DMPC/DMTAP vesicles, but the degree of decrease in the turbidity was inversely correlated to the DMTAP concentration (Fig. 1D). These results suggest that electrostatic interactions between SMA and vesicles govern the efficiency of formation of these complexes.
3.3. Cell uptake of lipid–SMA complexes To evaluate the effect of charged lipids on cellular uptake, SMA nanodisks were incubated with RAW264 or MDA-MB-231 cells for 30 or 120 min (Fig. 4). Addition of either DMPG or DMTAP to lipid–SMA complexes enhanced their uptake into RAW264 cells (Fig. 4A). At 120 min, the uptake of DMPC/DMPG–SMA and DMPC/DMTAP–SMA particles was increased by approximately 2-fold compared to uptake of
3.2. Physicochemical properties of lipid–SMA complexes Lipid–SMA complexes formed in the presence of 20 mol% DMPG or DMTAP were further analyzed by gel filtration chromatography with HEPES buffer as the eluent. Fig. 2A and B show the gel filtration elution 3
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Fig. 2. Representative gel filtration profiles of lipid–SMA complexes monitored by UV absorbance at 280 nm (open) and Rhodamine-PE fluorescence (closed). (A) DMPC/DMPG (8:2) and (B) DMPC/DMTAP (8:2) using HEPES buffer as eluent.
periphery in RAW264 cells (Fig. 5A and B), whereas, in MDA-MB-231 cells, particles localized to the center of the cells (Fig. 5C and D). Confocal images clearly showed the internalization of SMA nanodisks by MDA-MB-231 cells (Fig. 5E). Collectively, the subcellular distribution pattern was cell-type dependent.
DMPC–SMA particles. Since lipid concentrations were normalized to DMPC, the differences in the absolute uptake may be larger when considering total lipids. In contrast, only DMTAP but not DMPG significantly enhanced the uptake by MDA-MB-231 cells (Fig. 4B). These results were similar to those observed for the uptake of charged liposomes by macrophages (J774) or cervical cancer cells (HeLa) [19]. However, considering that the physicochemical properties, particularly the zeta potential of these particles, were not affected by the addition of charged lipids, the observed results suggest that the partial (local) charges in the planar bilayer surface of the discoidal particles confers the cell selectivity. Fluorescence microscopy images were obtained to visualize the subcellular distribution of SMA nanodisks. Fig. 5 shows a representative microscopy image of RAW264 and MDA-MB-231 cells after incubation with SMA nanodisks. Although the differences in cell localization caused by compositional alterations in the lipid–SMA complexes were not specifically defined, all three SMA nanodisks seemed to locate to the
3.4. Cytotoxicity of lipid–SMA complexes To examine the effect of charged lipids in lipid–SMA complexes on cytotoxicity, SMA nanodisks were incubated with RAW264 and MDAMB-231 cells for 6 h (Fig. 6). Although SMA with lower molecular weight has been shown to induce no appreciable effect on cell viability [20,21], SMA nanodisks even composed solely of neutral DMPC caused cytotoxicity, especially in MDA-MB-231 cells. In RAW264 cells, DMPC/ DMPG–SMA particles showed slightly higher cytotoxicity when compared to DMPC/DMTAP–SMA particles. In MDA-MB-231 cells, no significant differences in cytotoxicity were observed between all three
Fig. 3. DLS measurements and TEM analyses of lipid–SMA complexes. (A and B) Size distribution of lipid–SMA complexes. (C and D) TEM images of lipid–SMA complexes stained with ammonium molybdate. A and C; DMPC/DMPG–SMA complexes, B and D; DMPC/DMTAP–SMA complexes. 4
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Fig. 4. Effects of the addition of anionic or cationic lipids on cellular uptake of SMA nanodisks. (A) RAW264 and (B) MDA-MB-231 cells. Open bars; DMPC–SMA complexes, closed bars; DMPC/DMPG–SMA complexes, hatched bars; DMPC/DMTAP–SMA complexes. *p < 0.05 vs DMPC–SMA complexes.
nanoparticles due to electrostatic interaction with cells. However, surface charges of nanoparticles optimum for the cellular uptake virtually depend on the type of the nanoparticles [31]. Especially, molecular modeling study suggests that cellular uptake mechanism of discoidal lipid nanoparticles is quite different from traditional nanoparticles [32]. The addition of positively charged DMTAP into SMA nanodisks enhanced its uptake by both RAW264 and MDA-MB-231 cells, despite DMPC/DMTAP–SMA particles showing an equivalent zeta potential to either DMPC–SMA or DMPC/DMPG–SMA. Since these three nanodisks are discoidal in shape and SMA molecules are assumed to wrap around the circumference of bilayer disks, the cells seemed to recognize the positive charge enriched in the planar lipid membranes of the nanodisks. In contrast, the uptake of SMA nanodisks with negatively charged DMPG was enhanced in RAW264 cells, but not in MDA-MB-231. Scavenger receptors are usually involved in the uptake of negatively charged liposomes by macrophages [33], thus, it is conceivable that the cellular uptake mechanism for DMPC/DMPG–SMA and DMPC/ DMTAP–SMA particles by RAW264 cells is distinct. Indeed, the amount of uptake for DMPC/DMPG–SMA particles was increased from 30 min to 2 h whereas that for DMPC/DMTAP–SMA particles remained constant, as shown in Fig. 4, although we did not fully quantify these changes. Cellular uptake of nanoparticles can also be modulated by the presence of serum [34,35]. When administered intravenously, the surface of nanoparticles would be covered with serum proteins, which may alter the cellular uptake process. Thus, the effect of the protein corona formed around the nanoparticles will need further consideration in future experiments. It is well known that positively charged nanoparticles can exhibit greater cytotoxicity [36]. Our preliminary experiments suggested that DMPC/DMTAP liposomes with a diameter of approximately 100 nm formed at a ratio of 8:2 caused more evident cytotoxic effects in RAW264 cells compared with DMPG-containing liposomes (data not shown). However, as shown above, the effect of DMTAP addition into SMA nanodisks on the cytotoxicity was negligible possibly because the presence of SMA maintained the net negative charge and might have compensated for the toxic effect. Such reduced cytotoxic activity even when in the presence of positively charged lipids can be an advantage of SMA nanodisks. The cytotoxicity results in the present study did not vary between particles, but rather varied in a cell dependent manner, similarly to the subcellular distribution pattern visualized by fluorescence microscopy. These results suggest that the potency of the cytotoxic effects may be correlated with the degree of internalization by the cells. The mechanistic basis for such cell specificity is still unclear. Because SMA is an artificial polymer with no postulated physiological functions, SMA nanodisks are assumed to be biologically inert. Thus, surface modifications may be required in future pharmacologic applications. In the present study, we proposed that the cellular uptake of SMA nanodisks is modulated simply by changes in the lipid composition. Ideally SMA nanodisks would specifically accumulate and exert toxic effects in cancer cells but not in normal cells, and this could be achieved by combining SMA with specifically designed lipid
SMA nanodisks, but notably, they all showed significantly greater cytotoxicity in MDA-MB-231 cells than in RAW264 cells. These results may be due to SMA nanodisks' internalization by MDA-MB-231 cells as observed by the fluorescence microscopy (Fig. 5E). In the cellular uptake assay, no apparent cell damage was observed even at the PC concentration of 40 μg/mL (corresponding to approximately 60 μM). This was likely due to the differences in the incubation times between the experiments, 120 min in the cell uptake assay versus 6 h for toxicity assay.
4. Discussion SMA required the presence of NaCl to solubilize vesicles with charged lipids, unlike in the case of vesicles composed of DMPC alone. The addition of NaCl facilitated the vesicle solubilization both in the presence of positively and negatively charged lipids. This result is consistent with the previous observation that there is no significant preference of SMA to solubilize homogeneous lipid mixtures [22]. We have previously reported that SMA was not able to solubilize 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC) vesicles in phosphate buffer [17]. We have, however, confirmed the turbidity clearance of POPC vesicles in phosphate buffer containing NaCl (Fig. S2). Indeed, Scheidelaar et al. demonstrated that increased salt concentrations could promote vesicle solubilization [23]. Even in the presence of NaCl, the degree of the clearance decreased with increasing amount of DMTAP, which may have been due to the shift of transition temperature upon DMTAP enrichment [24]. Alternatively, the capacity of nanodisks to accommodate positively charged lipids may be limited. However, 30% of DMTAP (by weight) could be incorporated into nanodisks composed of apoA-I and DMPC as the major lipids, in phosphate buffered saline [15]. Liposomes composed of phospholipids are rapidly trapped in the reticuloendothelial system (RES) when administered systemically. To evade RES uptake and prolong the blood circulation time, the inclusion of polyethylene glycol (PEG) is most commonly used [25]. However, PEGylation often hampers the functionality of the affinity ligand attached to liposomes [26] and also affects the uptake of liposomes containing phosphatidylserine by macrophages [27]. In contrast, SMA nanodisks mimicking nascent HDL are thought to be biocompatible and consequently it is anticipated that PEGylation may not be required. We have previously reported that POPC–SMA complexes are not recognized as foreign substances [28]. Furthermore, immunogenicity induced by PEGylation is one of the major issues in liposomal formulations. For example, acceleration of blood clearance (ABC phenomenon) upon repeated injection of PEGylated liposomes is typically observed [29]. Although there is a possibility that SMA, being an artificial polymer similar to PEG, could exert an immunogenic response, drug-conjugated SMA has so far demonstrated no immunogenicity, in a previous study [30]. Several reports have shown that positively charged nanoparticles can have a higher cellular uptake when compared to negatively charged 5
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Fig. 5. Fluorescence microscopy images of (A, B) RAW264 and (C, D) MDA-MB-231 cells treated with DMPC/DMPG–SMA complexes. A and C; differential interference contrast image, B and D; Rhodamine-PE fluorescence image. (E) Confocal fluorescence image of MDA-MB-231 cells showing internalization of DMPC–SMA complexes.
Fig. 6. Viability of (A) RAW264 and (B) MDA-MB231 cells incubated with SMA nanodisks for 6 h. DMPC–SMA complexes (closed circles), DMPC/ DMPG–SMA complexes (closed triangles), DMPC/ DMTAP–SMA complexes (closed square). No significant differences existed between all three SMA nanodisks, but significant differences were observed between RAW264 and MDA-MB-231 cells at 6 h.
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compositions. More studies will be required to further our understanding of the cellular uptake mechanisms and the immunogenic properties of SMA nanodisks for the development of novel cell specific and biocompatible nano carriers for use in pharmacological settings.
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Declarations of competing interest The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper. CRediT authorship contribution statement Masafumi Tanaka: Conceptualization, Writing - original draft. Hisayasu Miyake: Investigation. Satoko Oka: Investigation. Shintaro Maeda: Investigation. Kenji Iwasaki: Resources, Investigation. Takahiro Mukai: Supervision, Writing - review & editing. Acknowledgments TEM observation was supported by the Platform Project for Supporting in Drug Discovery and Life Science Research (Platform for Drug Discovery, Informatics, and Structural Life Science) from the Japan Agency for Medical Research and Development (AMED), Japan. We would like to thank Dr. Kohei Sano for the help with fluorescence microscopy imaging, Ms. Ayano Yuge for technical assistance, and Enago for the English language review. Appendix A. Supplementary data Supplementary data to this article can be found online at https:// doi.org/10.1016/j.bbamem.2020.183209. References [1] C.J. Fielding, P.E. Fielding, Molecular physiology of reverse cholesterol transport, J. Lipid Res. 36 (2) (1995) 211–228. [2] P.G. Frank, Y.L. Marcel, Apolipoprotein A-I: structure-function relationships, J. Lipid Res. 41 (6) (2000) 853–872. [3] K.A. Rye, P.J. Barter, Formation and metabolism of prebeta-migrating, lipid-poor apolipoprotein A-I, Arterioscler. Thromb. Vasc. Biol. 24 (3) (2004) 421–428. [4] C.G. Brouillette, G.M. Anantharamaiah, Structural models of human apolipoprotein A-I, Biochim. Biophys. Acta 1256 (2) (1995) 103–129. [5] R.O. Ryan, Nanodisks: hydrophobic drug delivery vehicles, Expert Opin. Drug Deliv. 5 (3) (2008) 343–351. [6] K.K. Ng, J.F. Lovell, G. Zheng, Lipoprotein-inspired nanoparticles for cancer theranostics, Acc. Chem. Res. 44 (10) (2011) 1105–1113. [7] M.C. Orwick, P.J. Judge, J. Procek, L. Lindholm, A. Graziadei, A. Engel, G. Grobner, A. Watts, Detergent-free formation and physicochemical characterization of nanosized lipid-polymer complexes: Lipodisq, Angew. Chem. Int. Ed. Engl. 51 (19) (2012) 4653–4657. [8] R. Zhang, I.D. Sahu, L. Liu, A. Osatuke, R.G. Comer, C. Dabney-Smith, G.A. Lorigan, Characterizing the structure of lipodisq nanoparticles for membrane protein spectroscopic studies, Biochim. Biophys. Acta 1848 (1 Pt B) (2015) 329–333. [9] J.M. Dorr, S. Scheidelaar, M.C. Koorengevel, J.J. Dominguez, M. Schafer, C.A. van Walree, J.A. Killian, The styrene-maleic acid copolymer: a versatile tool in membrane research, Eur. Biophys. J. 45 (1) (2016) 3–21. [10] S. Lund-Katz, P.M. Laplaud, M.C. Phillips, M.J. Chapman, Apolipoprotein B-100 conformation and particle surface charge in human LDL subspecies: implication for LDL receptor interaction, Biochemistry 37 (37) (1998) 12867–12874. [11] D.L. Sparks, C. Chatterjee, E. Young, J. Renwick, N.R. Pandey, Lipoprotein charge and vascular lipid metabolism, Chem. Phys. Lipids 154 (1) (2008) 1–6. [12] T. Yamamoto, R.O. Ryan, Anionic phospholipids inhibit apolipoprotein E–lowdensity lipoprotein receptor interactions, Biochem. Biophys. Res. Commun. 354 (3)
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