Magnetic resonance imaging-based quantification of endosomal escape using iron oxide nanoparticles-loaded lipid nanoparticles

doi:10.21203/rs.3.rs-4133978/v1

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Magnetic resonance imaging-based quantification of endosomal escape using iron oxide nanoparticles-loaded lipid nanoparticles

https://doi.org/10.21203/rs.3.rs-4133978/v1

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Lipid nanoparticles (LNPs) have attracted attention as a promising and advanced platform for the delivery of nucleic acid-based therapeutics. The therapeutic efficacy of LNP-based drugs depends heavily on endosomal escape. However, few methods are available for quantifying the efficiency of endosomal escape. In this study, we developed a novel method to quantify the endosomal escape efficiency using magnetic resonance imaging (MRI). We synthesized ultrasmall iron oxide nanoparticles (IONPs) and incorporated them into LNPs to produce IO@LNPs. After cells internalized the IO@LNPs, we observed a decline in the R₂ relaxation over time, suggesting that free IONPs were dispersed due to endosomal escape. Biological electron microscopy further corroborated this finding, showing a strong correlation between the R₂ relaxation and the number of intracellular vesicles harboring the intact IO@LNPs. Furthermore, in-vivo MRI experiments in mice demonstrated an initial drop and a gradual increase in the T₂ signal at the tissue site where IO@LNPs were injected, indicating the potential for in-vivo application of our method. Our findings could lead to advancements in LNP-based nucleic acid delivery by enhancing the understanding of endosomal escape dynamics.

Biological sciences/Biophysics/Nanoscale biophysics

Physical sciences/Nanoscience and technology/Nanobiotechnology/Nanoparticles

Biological sciences/Drug discovery/Drug delivery

Biological sciences/Biochemistry/RNA

Biological sciences/Biotechnology/Nanobiotechnology/Nanoparticles

Lipid nanoparticles (LNPs) have recently gained prominence as an optimal platform for delivering RNA molecules surpassing traditional drug delivery systems, as demonstrated by success of LNP-based mRNA vaccines during the COVID-19 pandemic¹. Possible components of RNA-loaded LNPs (RNA@LNPs) include cationic/ionizable lipids, structural helper lipids, polyethylene glycol (PEG)-anchored lipids, cholesterol, and anionic RNA cargo (Fig. 1a). The cargo can be small interfering RNA (siRNA), microRNA (miRNA), circular RNA (circRNA), and messenger RNA (mRNA)^{2, 3, 4, 5}. These RNAs of different structures and sizes could become integrated into LNPs through the electrostatic attraction between anionic RNA molecules and cationic or ionizable lipids, resulting in the formation of RNA@LNPs.

RNA@LNPs are generally taken up by cells through endocytosis and reside within endosomes in the cells afterwards. Among the various biological barriers that can hinder the therapeutic efficacy of RNA drugs, the endosome is the primary and most challenging intracellular barrier^{6, 7}. Currently, RNA@LNP-based therapeutics overcome the endosomal barrier by leveraging cationic or ionizable lipids within the LNPs. These lipids interact with the anionic endosomal membranes, enabling the release of RNA molecules into the cytoplasm through a process referred to as endosomal escape^{8, 9, 10}. Previous studies have demonstrated a direct relationship between the ability of a drug to escape from endosomes and its therapeutic effect^{1, 11}. However, existing endosomal escape strategies only have limited effectiveness: less than 1% and 2% of the administered siRNA and mRNA were reported to reach the cytoplasm, respectively^{12, 13}. Hence, recent research has focused on improving endosomal escape^{14, 15}, which in turn requires innovative methods to measure the endosomal escape efficiency and identify effective LNP-based delivery platforms. Traditional methods for evaluating endosomal escape, including fluorescent labeling assays and transfection assays, exhibit limitations¹⁶. Specifically, these methods are limited to microscopic analysis and unsuitable for multicellular contexts or in-vivo situations, which impedes the accurate quantification of RNA endosomal escape facilitated by LNPs.

Here, we report a novel method to evaluate the efficiency of endosomal escape in multicellular situations using magnetic resonance imaging (MRI), one of the most widely used in vivo imaging modalities. Our methodology integrates MRI with iron oxide nanoparticles (IONPs), the latter serve as an MRI contrast agent to amplify the MR signals. Using this approach, we synthesized IONP-loaded LNPs (IO@LNPs) and performed MRI scans in multicellular and in-vivo settings. We then analyzed the changes in MRI signals to estimate the efficiency of endosomal escape.

Our hypothesis is that a cluster of IONPs encapsulated within the LNPs would generate an MRI signal distinct from that of dispersed IONPs scattered throughout the cytoplasm following endosomal escape (Fig. 1b). Consequently, changes in the MRI signal may serve as an effective metric for determining the efficiency of endosomal escape.

Characterization of IONPs and IO@LNPs

Ultrasmall IONPs (< 8 nm) have been reported to penetrate the glomerular filtration barrier of the kidneys, resulting in high renal clearance without toxicity. In a previous study, citrate-stabilized ultrasmall IONPs were synthesized using a solvothermal method without surface coating such as PEG modification¹⁷. The surface of IONPs is generally modified with a polymer coating for prolonged blood circulation and long-term colloidal stability in cells^{18, 19}. However, unmodified IONPs carry a more negative surface charge, making it easier for them to become encapsulated in LNPs²⁰. This process is similar to the integration of RNA with cationic/ionizable lipids. Hence, bare IONPs were selected in this study over the conventional IONPs. The peaks observed in the Fourier-transform infrared (FTIR) spectrum confirm the successful synthesis of the IONPs (Fig. 1c)^{17, 20}. The peaks at 1,587 and 1,372 cm^− 1 correspond to the stretching vibrations of conjugated C = O and C-O bonds, and those near 2,900 and 1,100 cm^− 1 indicate the stretching vibrations of -CH₂ and -OH groups of sodium citrate, respectively. The presence of these peaks assigned to key functional groups suggests that the surface of IONPs was successfully stabilized with citrate. A vibrating sample magnetometer (VSM) test was conducted to measure the magnetic properties of the IONPs, which are pertinent to their contrast effect in MRI (Fig. 1d). The normalized saturation moment of the IONPs was approximately 3 emu/g, which corresponds to the literature regarding its value^{17, 20}. In addition, the ability of IONPs to respond sensitively to an external magnetic field in the hysteresis loop indicates that these nanoparticles are either ferromagnetic or superparamagnetic. However, the near-zero coercivity implies that the IONPs are not ferromagnetic but solely paramagnetic.

IO@LNPs (IONP:lipid = 0.2 w/w) of a homogenous size were prepared with 1,2-di-O-octadecenyl-3-trimethylammonium propane (DOTMA; cationic lipid), 1,2-distearoyl-sn-glycero-3-phosphocholine (DSPC; structural helper phospholipid), 1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N-[methoxy(polyethylene glycol)-5000] (mPEG-DSPE); PEGylated lipid), and cholesterol using a fluidic-based mixture (see Methods). The zeta potential (surface charge) of IO@LNPs was 6.11 ± 7.18 mV (mean ± SD), which is more positive than that of IONPs (-18.0 ± 5.18 mV)²⁰ but less than that of empty LNPs (11.9 ± 6.01 mV) (Fig. 1e). The hydrodynamic size of IONPs, measured by dynamic light scattering (DLS), was 2.289 ± 0.641 nm (mean ± SD) with a polydispersity index (PDI) of 0.235 (Fig. 1f). Meanwhile, these values for IO@LNPs and empty LNPs were 93.13 ± 30.66 nm (PDI 0.166) and 129.7 ± 31.18 nm (PDI 0.040), respectively. The surface charge and size distribution results indicated that the negatively charged IONPs attracted the positively charged lipids in the LNPs, reducing the effective number of cationic lipids on the surface of IO@LNPs²¹.

The structures of IONPs and LNPs were observed using transmission electron microscopy (TEM) with negative staining and cryo-electron microscopy (cryo-EM), respectively. The images in Fig. 1g show the morphology of IONPs, IO@LNPs, and empty LNPs. The negative-staining TEM image shows that the IONPs were ultrasmall in size and uniformly dispersed. The cryo-EM images of IO@LNPs demonstrate the integration of IONPs into LNPs, which is clearly distinct from the vacant core observed in empty LNPs. In addition, electrophoresis was performed based on the polarity determined from the zeta potentials, and the results also verified that most of the added IONPs were successfully incorporated into the LNPs (Supplementary Fig. 1).

Furthermore, stability tests were carried out on IONPs and IO@LNPs using distilled water (DW), phosphate-buffered saline (PBS), and Roswell Park Memorial Institute (RPMI) 1640 growth medium containing 10% fetal bovine serum (FBS) and 1% antibiotic-antimycotic. The IONPs were stable in various media for over 7 h (Supplementary Fig. 2a). In addition, the size and PDI of IO@LNPs remained stable for a week in these media (Supplementary Fig. 2b).

MR properties of IONPs and IO@LNPs

In MRI technology, the R₂ relaxation rate is the reciprocal of the transverse relaxation time (T₂) that represents the progression of the transverse component of proton magnetization from 100% (pulse flip) to 37%. Similarly, the R₁ relaxation rate is the reciprocal of the longitudinal relaxation time (T₁) that represents the progression of the longitudinal component of proton magnetization from 0% (pulse flip) to 63%. Graphical MRI data obtained in the time domain can be converted to images expressed as R₁ or R₂ values in the frequency domain. The r₁ and r₂ relaxivities represent the change in R₁ and R₂ values divided by the change in IONP concentration, respectively. In this study, the units of r₁ and r₂ are assigned as s^− 1 [mg/ml]⁻¹.

To examine the MR signals produced by the lipid components of LNPs, the MR signals of empty LNPs were analyzed. The signal from the empty LNPs (r₁ = -0.017; r₂ = 0.031) showed a constantly low level with a marginal deviation from 0 (Supplementary Fig. 3). Thus, we regarded the MR signals from the lipid components of LNPs as zero. The IO@LNPs samples were prepared at varying IONP-to-lipid ratios (w/w). In the same batch experiment, the r₂ value showed no significant difference between IO@LNPs with lower and higher IONP concentrations (r₂ = 6.97 for 0.1 w/w and r₂ = 7.24 for 0.2 w/w, respectively) (Supplementary Fig. 4). Thus, IO@LNPs with a lipid-to-IONP ratio of 0.2 w/w were used in subsequent experiments.

MRI phantom maps were obtained with varying concentrations of IONPs to acquire MR signals from free IONPs and IO@LNPs. The map images showed that while there were slight R₁ signals observed with IONPs (r₁ = 0.33) and IO@LNPs (r₁ = 0.016), the R₂ values were more prominent with IONPs (r₂ = 2.70) and IO@LNPs (r₂ = 5.69) (Fig. 2a–b). These results demonstrate that the IONPs and IO@LNPs may serve as T₂ contrast agents^{22, 23}.

In addition, the IO@LNPs showed lower r₁ and higher r₂ values than the dispersed IONPs. These findings are expected from the principles of MR. The R₁ relaxation rate can be regarded as an indicator of how quickly the protons in water return to equilibrium after excitation by radio frequency (RF) energy during MRI scan. The dispersed IONPs have greater mobility and therefore promote the dissipation of RF energy, leading to an increase in the R₁ value compared with that of IO@LNPs. In contrast, the R₂ relaxation rate increases as the local magnetic field inhomogeneities increase in proportion to the square of the magnetic moment^{18, 24}. When the IONPs are clustered within the LNPs, their magnetic moments align in an additive manner, thereby increasing the R₂ value. Thus, IO@LNPs have a larger R₂ value compared to the dispersed IONPs. These properties were similarly demonstrated in a previous study that compared the relaxation rates of ultrasmall IONPs and their clusters²⁵.

To confirm the reduction in R₂ value due to the dispersion of IONPs from LNPs, the IO@LNPs were combined with Triton X-100 detergent to induce the dissociation of LNPs. The sample added with Triton X-100 showed a higher r₁ value (0.58) and a lower r₂ value (1.63) than that without the detergent (r₁ = 0.047 and r₂ = 5.74, respectively) (Fig. 2c). These results show that detergent treatment led to the dissociation of IO@LNPs and release of IONPs. We further verified that the IONPs dispersed from LNPs have very similar MR properties to those of the original free IONPs (r₁ = 0.40 and r₂ = 2.74), thus supporting our proposed approach of detecting endosomal escape (Fig. 2d).

In-vitro MRI for evaluating endosomal escape

To determine the observation window for the endosomal escape of IO@LNPs in vitro, the temporal trends of LNP uptake in cells (4T1 cell line) were assessed by labeling the IO@LNPs with a fluorescence dye (DiD) (Supplementary Fig. 5). The cellular uptake of IO@LNP/DiD was then observed using confocal laser scanning microscopy (CLSM) imaging. The cells were treated with the same amount of IO@LNP/DiD and washed after different treatment times (Supplementary Fig. 6a). As expected, the intracellular uptake of LNPs was rare at 0 min (Supplementary Fig. 6b). At 20 min, the LNPs primarily appeared on the cellular surface, and by 40 min they started to move further into the cytoplasmic region. Therefore, endosomal escape seemed to occur within an hour, as evidenced by the previously reported rapid endosomal movement during endosomal escape²⁶. Taken together, we observed a continuous onset of endosomal escape starting from 40 min.

To measure MRI signal of cells treated with IO@LNPs, the cells were continuously treated with IO@LNPs for 40 min, followed by thorough washing to remove any IO@LNPs remaining from the media. The cells were then incubated for different additional durations (0, 20, 40, and 80 min) to allow endosomal escape (Fig. 3a). Cells harvested from each group were fixed with 4% paraformaldehyde (PFA), and cell-pellets were prepared for MR signal acquisition (see Methods). Between 40 and 80 min, the in-vitro MR results confirmed that a constant R₁ value along with a declining R₂ value, implying the endosomal escape at the time period (Fig. 3b,c). The R₁ value in the in-vitro MR experiments showed no statistical difference from 40 to 80 min, as expected from the mild R₁ value difference between IO@LNPs and free IONPs (Fig. 2a,b). This finding reinforces our approach of prioritizing the R₂ over the R₁ signal. However, at the 120 min time point, both the R₁ and R₂ values decreased. This could be linked to the rapid exocytosis of ultrasmall IONPs²⁷ and/or the biodegradation of IONPs stemming from their unmodified exposed surface²⁸.

To confirm that the changes in MR signals were due to endosomal escape, cells were treated with chloroquine, a substance renowned for enhancing endosomal escape. Although there was no significant difference in the R₁ value between the treated and untreated groups, the group treated with chloroquine exhibited reduced R₂ values compared to the untreated one (Fig. 3d,e). This decrease in R₂ value might be due to an immediate endosomal escape following the IO@LNP treatment.

MRI-based in-vitro quantification of endosomal escape

Using the MR theory and a simple application of the chain rule from calculus, we were able to calculate the efficiency of endosomal escape in %/min (Supplementary Methods). The efficiency of endosomal escape over the entire duration of 40–120 min was determined to be 0.93%/min. In the first half of this duration (40–80 min), the efficiency was 1.98%/min. Previous studies have consistently confirmed that protein expression in HeLa cells began approximately 1.5 h following RNA@LNP treatment¹². Additionally, as the major cellular organelle related to endosomal escape, the early endosomes began to encompass the nanoparticles 1 h after treatment^{8, 9, 12, 26}. Therefore, our observation that endosomal escape actively occurred at approximately 1 h is consistent with the previous studies. The slightly earlier onset of endosomal escape in our study (by approximately 20 min) may be due to differences in the nanoparticle characteristics and cell line used.

Earlier studies have also shown a compromise between directly evaluating the endosomal escape efficiency and quantifying it in multicellular or in-vivo settings (Table 1). High-resolution microscopy is commonly used for direct measurement of endosomal escape. This involves calculating the subcellular spatial relationship between the drug delivery systems (DDSs) and endosomes at various stages (Method 1).

Table 1

Comparison of different methods for evaluating the endosomal escape of delivered molecules.
Method	Imaging modality	Multicellular evaluation	In-vivo applicability	Association	Quantification	Mechanism	Ref
1	Confocal microscope	No	No	Direct	Correlation with super-resolution microscope	Co-localization and correlation	^{12, 33, 41, 42}
2	Confocal microscope	No	No	Indirect	Fluorescence pattern	Diffusion of escaped material	^{16, 43, 44, 45}
3	Microscope	No	No	Indirect	Super-resolution microscope	Membrane-impermeable dye	^{46, 47, 48, 49, 50, 51}
4	Microscope	No	No	Indirect	Super-resolution microscope	A signal upon contact with the escaped component and a protein in the cytosol	⁵²
5	Microscope	Yes	No	Indirect	Fluorescence intensity	Membrane damage marker	^{53, 54, 55}
6	Microscope	Yes	Yes	Indirect	ROS intensity	pH-sensitive marker under acidic endosome	^{56, 57}
7	Microscope	Yes	Yes	Indirect	FRET-based fluorescence intensity	e.g., Intracellular Dicer reaction, glutathione reaction	^{58, 59, 60}
8	Microscope	Yes	Yes	Indirect	Fluorescence intensity	Transfection of a reporter gene (e.g., luciferase)	^{7, 16}
This study	MRI	Yes	Yes	Direct	R₂ change	IONP dispersal

Methods 2–4 in Table 1 also use microscopy to assess the efficiency. Method 2 utilizes the changes in the DDS fluorescence pattern before and after the escape. However, fluorescent labeling may affect the escape efficiency and, in turn, reduce the accuracy of efficiency estimates. Method 3 utilizes membrane-impermeable dyes such as calcein that produce unique fluorescent patterns after escape. Nevertheless, this approach poses a challenge as the efficiency of endosomal escape can vary between small molecules and the DDS. Method 4 assumes a constant signaling protein level throughout the cytoplasm, which may not be true. Thus, unlike Method 1, Methods 2–4 only indirectly measure the efficiency.

It is important to note that Methods 1–4 are not applicable to multicellular or in-vivo settings because the amount of materials in the cytosol is usually low²⁹. However, using fluorescence with sufficient fluorophores may enable multicellular or in-vivo quantification (Methods 5–8). Flow cytometry has been successfully applied for multicellular estimation using these methods^{30, 31}.

Nevertheless, Methods 5–8 are all indirect measures of endosomal escape efficiency. For example, in Method 5 the model cellular membrane does not fully represent the actual endosomal membrane. Method 6 is indirect in the following two senses. If a DDS is coupled to pH-sensitive moieties, these moieties should ideally remain consistently attached to the DDS, a scenario that may not be true. On the other hand, if the pH-sensitive moieties are introduced separately from the DDS, their efficiency may differ from those of the DDS. Method 7 is also indirect because the surface modifications used for Förster resonance energy transfer (FRET) could affect the endosomal escape efficiency.

Method 8, often referred to as transfection assay, is generally regarded as a standard effectiveness measure of RNA therapeutics by directly evaluating the transfected outcomes that are closely tied to therapeutic efficacy. However, due to the difficulty in measuring transfection efficiency and its potential lack of reproducibility, there is a need for LNP quality assessment methods based on single factors closely linked to the therapeutic efficacy such as endosomal escape. For example, additional steps after RNA escape from endosomes (e.g., RNA transportation to the nucleus, efficiency of transfection, capability of RNAs to be translated, and degradation of nucleic acids) could complicate the distinction between other factors and LNP effectiveness. Also, variations in the size and chemical composition of nucleic acids may affect transfection efficiency and hinder the comparison of different studies. For example, the endosomal escape efficiency can be affected by the charge on RNA, since RNA molecules carrying a more negative charge experience a stronger mutual repulsive force that facilitates their escape from the LNPs³². The negative charge on RNA in turn is determined by a variety of factors, such as the RNA length, nucleotide sequence, single- or double-stranded structures, and the number of attached phosphate groups. Thus, a direct quantification of endosomal escape solely focusing on DDS is preferable than transfection assay.

In contrast, our approach here offers standardization because of the homogeneity and reproducibility of the IONPs and IO@LNPs. Furthermore, since the change in MR signal is in theory solely due to endosomal escape, our method can be categorized as a direct measurement technique. In addition, the MRI technique enables the application in multicellular and in-vivo contexts.

Validation of MRI-based method for evaluating endosomal escape

To validate the in-vitro MR results, biological transmission electron microscopy (Bio-TEM) imaging was carried out to observe intracellular vesicles (endosomes) containing IO@LNPs. There was no detectable endosome in the IONP-treated or control group without IO@LNPs (Fig. 4a). However, the IO@LNPs were visible in the TEM images (Fig. 4b).

Based on this observation, we could calculate the number of endosomal structures containing IO@LNPs in single cells (Fig. 4c). The frequency of endosomes decreased as the total incubation time increased. The coefficient of determination (R²) between the average frequency of endosomes and the R₂ relaxation rate from the in-vitro MRI was 0.95 (Fig. 4d). This result supports that the change in R₂ values observed in the in-vitro MRI maps was due to endosomal escape. Some cells released many extracellular vesicles in the Bio-TEM images taken at 120 min. This suggests that the decrease in R₂ value in in-vitro MRI data between 80 and 120 min (Fig. 3b-c) might be affected by active exocytosis²⁷.

MRI-based in-vivo quantification of endosomal escape

We also applied our technique to evaluate endosomal escape in vivo using a mouse model. IO@LNP stained with DiR (IO@LNP/DiR) was injected intramuscularly, and fluorescence signals at the injection site were quantified using an In Vivo Imaging System (IVIS) (Fig. 5a). In in vivo imaging, the longer-wavelength DiR was more advantageous, so it was used as a fluorescent dye instead of DiD. The radiant efficiency remained stable for 3 h in the muscle but was not observed in other organs (Fig. 5b and Supplementary Fig. 7). This result indicates that the majority of the injected LNPs remained without diffusion to other organs within 3 h after administration.

Additionally, IO@LNPs and empty LNPs were intramuscularly injected into mouse hindlimbs, and T₂-weighted images were acquired at different time points (Fig. 5c). Following the injection of IO@LNPs, two distinct signals were observed from the muscle tissue: intense bright areas and dark regions. The bright areas, which were also observed in the mice injected with empty LNPs, were attributed to the signals caused by PBS, as a similar area was observed in mice treated with PBS only. In contrast, the dark regions were exclusive to the IO@LNP-treated samples, suggesting that they were unique signals associated with IO@LNPs. These dark regions became gradually brighter after administration, implying endosomal escape (Supplementary Fig. 8).

After treating muscle tissue with PBS, empty LNPs, and IO@LNPs, the tissues were obtained from mice at different time points (0, 1, and 3 h), and Bio-TEM images were acquired for validation study (Fig. 5d and Supplementary Fig. 9). Unlike PBS and empty LNPs, treatment with IO@LNPs produced a distinctive pattern caused by the aggregated IONPs (black dots) within IO@LNPs. Immediately after treatment (i.e., at 0 h), a pattern of IO@LNPs was observed along the spaces between muscle fibers. At 1 h, the IO@LNPs penetrated the muscle fibers, and the nanoparticle pattern was not visible in many muscle fibers. After 3 h, the IO@LNP pattern was difficult to discern. This indicates that IONPs loaded in IO@LNPs were gradually ingested by muscle cells and became dispersed, leading to a significant decrease in the amount of loaded IONPs after 1 h. Therefore, the bio-TEM results well support the in-vivo MR findings.

Efficient RNA delivery by LNPs cannot be therapeutically effective unless the RNA molecules escape from the endosome to the cytoplasm to perform their functions. Endosomal escape is a bottleneck for intracellular delivery, and its efficiency is crucial for determining the efficacy of RNA-based therapies. Therefore, LNPs with high endosomal escape efficiency are sought after as promising drug vehicles for RNA delivery³³. In this study, we proposed using MRI to measure the efficiency of LNP-mediated endosomal escape. Ultrasmall and uncoated IONPs with negative charges were used to explore the benefits of electrostatically mediated encapsulation within the LNPs. We observed a dramatic decrease in the r₂ value of the IONPs after they escaped from the LNPs and endosomes. The in-vitro endosomal escape efficiency of the IO@LNPs was successfully measured, and the method’s potential for in-vivo measurements was presented.

Most DDS studies on mRNA delivery assess the extent of translation from mRNA to protein using luciferase mRNA^{34, 35, 36}. However, our method, which employs IONPs with reproducible sizes and charges, may act as an optimal reference for the evaluation of endosomal escape. Notably, our results obtained in the presence of chloroquine, a chemical known to enhance endosomal escape, suggested that the change in the R₂ signal was largely dependent on endosomal escape. This indicates that our technology can play a critical role in comparing endosomal escape efficiencies across diverse LNP formulations in RNA@LNP research. Moreover, this technique for screening endosomal escape may be extended to different DDS types aimed at nucleic acid delivery.

Note that the proposed method has certain limitations. First, because this study only considered one IONP formulation and one LNP composition, it is necessary to compare the results with those of other compositions to check if the endosomal escape efficiency is truly linked to therapeutic efficacy. Secondly, given the probable exocytosis of cells after 120 min (following a 40 min incubation with IO@LNPs and an 80 min observation period), it is crucial to examine the impact of IONPs’ exocytosis and degradation on our approach.

Solvothermal synthesis of IONPs. Citrate-stabilized ultrasmall IONPs were synthesized using the solvothermal method, in accordance with a previous study¹⁷. First, FeCl₃·6H₂O (270.25 mg, 1.00 mmol) was dissolved in diethylene glycol (DEG, 10 ml) under stirring at 1,000 rpm. This solution was mixed with Na₃Cit·2H₂O (117.75 mg, 0.40 mmol) and heated at 80°C for 2 h, added with anhydrous sodium acetate (328 mg, 4.00 mmol), and then heated at 200°C for 4 h. After cooling to 100°C, the reaction mixture was added with ethanol and centrifuged at 8,500 rpm for 30 min. The supernatant was discarded. The iron oxide sediment was washed with ethanol, separated by centrifugation at 7,000 rpm for 30 min, and dried overnight at room temperature.

Self-assembly of IO@LNPs. The chemical composition of our LNPs was based on a previous study³⁷. It is well documented that cationic (e.g., DOTMA) and ionizable lipids can encapsulate negatively charged molecules including nucleic acids³⁸. Cationic lipids were used in this study to enhance the loading efficiency. The mixed lipids (DOTMA/DSPC/cholesterol/PEG = 50/10/38.5/1.5 mol%) were dissolved in ethanol to a final total lipid concentration of 10.4 mg/ml. The lipids were mixed with 25 mM sodium acetate buffer (pH 4.0) containing the IONPs (IONP:lipid = 0.2 w/w), using a T-junction system and a Gemini 88 Plus Dual Rate syringe pump (KD Scientific Inc., USA) at a constant flow rate. This ethanol-water formulation was then enclosed in Spectra/Por 2 dialysis membrane (MWCO: 12–14 kD; Spectrum Labs, USA) and dialyzed in 1,000-fold volume of 1X PBS overnight to remove ethanol.

A lipophilic fluorescent dye (DiD) was applied to IO@LNPs to allow visualization of the LNPs under a fluorescence microscope. IO@LNP/DiD was prepared following the same procedure as IO@LNPs, except for the addition of 4 µl of DiD into the lipid-ethanol phase.

In-solution MRI phantom of IO@LNPs. To compare the MR characteristics of IONPs, IO@LNPs, and empty LNPs, MRI phantom experiments were performed using a Powerscan MRS 7024 instrument (MR SOULUTIONS, UK). An MRI scanner with a high magnetic field of 7.0 T was used to improve the resolution and contrast. The samples were prepared by serial dilution with saline (concentration levels: 1/1, 1/2, 1/3, 1/4, and 1/5) and placed in 0.2-ml polymerase chain reaction (PCR) tubes. The IONP, IO@LNP, and empty LNP samples at the concentration level of 1/1 contained 0.87 mg ml^− 1 IONP + 0 mg ml^− 1 lipid, 0.87 mg ml^− 1 IONP + 4.33 mg ml^− 1 lipid, and 0 mg ml^− 1 IONP + 4.33 mg ml^− 1 lipid, respectively. The MRI sequence parameters for the R₁ and R₂ relaxation rates are presented in Supplementary Table 1.

In-solution MRI phantom of dissolved IO@LNPs. Changes in the MR signal according to the dispersion of adjacent IONPs within the LNPs were evaluated by dissolving the IO@LNPs using a detergent. The IO@LNP solution was mixed with 2% Triton X-100, incubated at 37°C for 10 min, and cooled at room temperature for 5 min. Samples were prepared at concentration levels ranging from 1/1 to 1/5. The IO@LNP samples treated with Triton X-100 at the concentration level of 1/1 contained 0.52 mg ml^-1 IONP + 2.59 mg ml^-1 lipid. Finally, the MR signal was measured using a 0.7 T MRI scanner, with the MRI sequence parameters presented in Supplementary Tables 2 and 3.

In-vitro MRI. The protocol for in-vitro MRI has been summarized in the literature^{39, 40}. The 4T1 cells were cultured in T-flasks, washed with Dulbecco's phosphate-buffered saline (DPBS), and initially incubated for 40 min in fresh RPMI medium containing IO@LNP/DiD. After collecting and changing the medium, the samples were further incubated for 0, 20, 40, and 80 min. Then, the cells were washed with DPBS and detached from the flasks by 1X Trypsin-EDTA treatment. Cell pellets obtained by centrifugation were resuspended in 1 ml DPBS, and the total cell density and viability for each sample were calculated using an automatic cell counter. The cells (ranging from 1.7 × 10⁷ to 2.2 × 10⁷ cells/ml for each experimental group) were transferred to a microtube, and cell pellets were collected by centrifugation for 10 min at 500 g and 4°C. The pellets were then resuspended with 4% PFA, incubated for 15 min at room temperature, centrifuged again for 10 min at 500 g, and rinsed with DPBS. The cells were additionally centrifuged, resuspended in DPBS (50 µl), and transferred over 4% agarose-PBS gel in PCR tubes. Agarose gel was used for the equal height of the cell pellets. After waiting for the cells to completely precipitate, the supernatant was removed. The resultant samples were stored at 4°C. R₁ and R₂ values were acquired as described in Supplementary Table 4.

During analysis of the MRI map, five different concentrations were employed to create a regression curve linking the IONP concentration with the MR relaxation rate. The MRI experiments were repeated thrice for each of the three biological replicates. To determine whether the regression slope was zero, F-statistics were calculated using GraphPad Prism 8.4.2. In addition, to compare the regression slopes between different groups, one-way ANOVA test was performed using the aov function in R. Standard statistical conventions, including star symbols, were applied unless stated otherwise.

Preparation of biological TEM samples. Samples of 4T1 cells treated with IONPs or IO@LNPs were prepared using the same methods as in-vitro MRI until cell fixation. The cell pellets collected in each microtube were treated with Karnovsky’s Fixative to preserve the biological samples for electron microscopy. During primary fixation, the cells were incubated overnight at 4°C in 0.05 M sodium cacodylate buffer (pH 7.4), 2% PFA, and 2% glutaraldehyde. Afterwards, the cells were washed with 0.05 M sodium cacodylate buffer, added with 1% osmium tetroxide diluted with 0.1 M sodium cacodylate buffer, incubated for 2 h at 4°C, washed with DW, and incubated for En Bloc staining in 0.5% uranyl acetate overnight at 4°C. Afterward, the samples were washed with DW and gradually dehydrated with ethanol (30%, 50%, 70%, 80%, 90%, and 100%). Spurr's Resin was applied for transition and embedding, and then the samples were incubated overnight at 70°C for polymerization. The solidified samples were sectioned into thin films using an EM UC7 ultramicrotome (Leica, Germany). Finally, Bio-TEM images were acquired to observe the intracellular IO@LNPs.

Semi-quantitative analysis for Bio-TEM. To analyze the Bio-TEM results, we defined two criteria for identifying IO@LNPs in intracellular vesicles (endosomes). First, the signal should be significantly lower than that in the surrounding cytoplasm. Second, the vesicle should include at least three black dots (IO@LNPs). To semi-quantitatively analyze the selected cells containing endosomes, only unique cells were considered. The number of endosomes in these cells was counted and corrected based on the exposed surface area relative to the total surface area of the cells. Also, only the top 50% of cells based on the number of endosomes were considered from a minimum of 30 cells in each group, as a quality control (QC) process. Subsequently, a regression curve was generated between the time points and the frequency of endosomes. The frequency of endosomes per cell was plotted against the R₂ value at the corresponding time point.

In-vivo experiments. Female BALB/c mice (6 weeks old) were injected with 50 µl of DiR fluorescent IO@LNP (1.8999 mg IONP/9.5657 mg lipid/kg mouse) intramuscularly in their right hindlimbs. The in-vivo distribution of LNP lipid components was then investigated using an In Vivo Imaging System (IVIS) pre-injection and at 0, 1, 2, and 3 h post-injection. To acquire the MR signal of IO@LNP and empty LNP in vivo, BALB/c mice were properly fixed in a mold and underwent breath gating before acquiring the T₂-weighted images both pre-injection and at 0, 1, 2, and 3 h post-injection. The number of tissue slices was set to 13, and the median slice was selected for subsequent quantitative measurements. The MR contrast effect in a given tissue was normalized using neighboring tissue where this effect was not observed. The muscle tissue was treated with PBS, empty LNP, and IO@LNPs using the same administration route, and after 0, 1, and 3 h the same Bio-TEM process was applied, including Karnovsky's Fixation.

Acknowledgements

This research was supported by the National Research Foundation of Korea (NRF) (NRF-2019M2D2A1A01058210, NRF-2020R1C1C1009000, NFR-2021M2E8A1039564, and BK21 FOUR Program No. 5120200513755), Creative-Pioneering Researchers Program through Seoul National University (SNU), Korea Evaluation Institute of Industrial Technology (KEIT) grant funded by the Korea government (MOTIE) (No. 20018522), and Korea Drug Development Fund funded by Ministry of Science and ICT, Ministry of Trade, Industry, and Energy, and Ministry of Health and Welfare (HN22C0632).

Corresponding Author

Hyung-Jun Im, MD, PhD

Associate Professor

Department of Applied Bioengineering,

Graduate School of Convergence Science and Technology

Seoul National University

Office: 031-888-9187

Homepage: http://tmtl.snu.ac.kr

E-mail: [email protected], [email protected]

Author Contributions

Jeongbin Park and Somin Lee contributed equally to this project, and they conceived and designed all experiments. Jeongbin Park and Somin Lee performed the nanoparticle synthesis, characterization, and in-vitro and in-vivo experiments. Jeongbin Park interpreted and analyzed the magnetic resonance images. Han Na Jung, Shengjun Lee, and Woojin Lee performed the in-vivo experiments. Jeongbin Park, Somin Lee, and Hyung-Jun Im wrote the manuscript. All authors reviewed the final version of the manuscript.

Disclosure

Hyung-Jun Im is the co-founder and Chief Scientific Officer of Portrai, Inc. Jeongbin Park is a researcher at Portrai Inc. Otherwise, the authors have no competing financial interests to declare.

Ethics declarations

Animal experiments and MRI phantom studies were approved by the Seoul National University Bundang Hospital (SNUBH). The approval code is BA-2102-314-012-05.

Competing interests

Jeongbin Park, Somin Lee, and Hyung-Jun Im were listed as inventors in patent applications for the technology presented in this manuscript (International Patent Application No. PCT/KR2022/021076 and priority claims based on Korean patent application no.10-2022-0181667). Otherwise, the authors declare no competing interests.

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Yes there is potential Competing Interest. Hyung-Jun Im is the co-founder and Chief Scientific Officer of Portrai, Inc. Jeongbin Park is a researcher at Portrai Inc. Jeongbin Park, Somin Lee, and Hyung-Jun Im were listed as inventors in patent applications for the technology presented in this manuscript (International Patent Application No. PCT/KR2022/021076 and priority claims based on Korean patent application no.10-2022-0181667). Otherwise, the authors declare no competing interests.

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Magnetic resonance imaging-based quantification of endosomal escape using iron oxide nanoparticles-loaded lipid nanoparticles

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Abstract

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Introduction

Characterization of IONPs and IO@LNPs

MR properties of IONPs and IO@LNPs

In-vitro MRI for evaluating endosomal escape

MRI-based in-vitro quantification of endosomal escape

Validation of MRI-based method for evaluating endosomal escape

MRI-based in-vivo quantification of endosomal escape

Discussion

Methods

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References

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