Bioengineering protein gates by modification of channel porins

The key factors in the design of AOs with activity triggered by changes in environmental conditions are on-demand permeability of the compartment towards enzymatic substrates/products and structural integrity of the polymersome, which mimics that of natural organelles. Therefore, our biomimetic strategy aimed to equip PMOXA 6 -PDMS 44 -PMOXA 6 polymersome membranes with protein gates that are responsive to changes in glutathione (GSH) concentrations in intracellular environments, while preserving the structure of the nanocompartment (Fig. 1a).

Fig. 1 Engineering stimuli-responsive OmpF. a Schematic representation of modified OmpF acting as a gate in catalytic nanocompartments. b Molecular representation of the OmpF-M cysteine mutant37. c Chemical modification of OmpF-M cysteine mutant with the spin probe bis-(2,2,5,5-tetramethyl-3-imidazoline-1-oxyl-4-yl) disulphide. d Chemical modification of OmpF-M cysteine mutant with the fluorophore SAMSA-CF Full size image

It has been shown very recently that chemical modifications of amino acid residues at key locations of the OmpF porin backbone influence the translocation of substrates through the pore in a pH-responsive manner36. Here we go one step further by using a double mutant of OmpF37 to attach molecular caps to genetically introduced cysteine residues that serve to block/unblock the OmpF pore upon changes in redox potential, which occur when the system enters the intracellular microenvironment (Fig. 1b). In contrast to polymersomes with membranes containing OmpF genetically modified to release a payload in reductive conditions35, our system controls the overall functionality of the AOs. We chose a cysteine double mutant of OmpF (OmpF-M) because cysteine residues, replacing the amino acids K89 and R270, were expected to form reduction-sensitive disulphide bonds with molecules selected to serve as molecular caps. These molecular caps remain attached in mildly oxidising environments and block substrate diffusion through the pore, whereas in the presence of reducing agents, such as intracellular GSH, their cleavage restores normal passage of small molecular weight molecules (<600 Da) through the OmpF pores. This approach mimics pathways of metabolism regulation, where proteins within the membranes of natural cell organelles are irreversibly activated or deactivated on demand43,44. In addition, we were interested in developing an irreversible protein gate in order to be able to rapidly evaluate the functionality of the organelle in vivo.

The ability of the cysteine residues of OmpF-M to form disulphide bonds with thiol groups of small molecular weight molecules was examined by two complementary assays, one using a suitable spin probe (bis-(2,2,5,5-tetramethyl-3-imidazoline-1-oxyl-4-yl) disulphide) and the second using the fluorescent dye SAMSA fluorescein (SAMSA-CF) (Fig. 1c, d).

Coupling reaction of the molecular caps with the cysteine residues of OmpF-M resulted in the formation of OmpF conjugates (OmpF-S-S-CF for OmpF conjugated with SAMSA-CF, and OmpF-S-S-NO for OmpF conjugated with bis-(2,2,5,5-tetramethyl-3-imidazoline-1-oxyl-4-yl) disulphide), respectively.

Binding of the thiol reactive spin probe to the protein was evaluated by a combination of LC-MS-MS and electron paramagnetic resonance (EPR). Upon in-gel digestion of the porin45, LC-MS-MS analysis of the peptide fragments indicated a very high labelling efficiency of the spin probe to cysteine residues of the OmpF-M (96 ± 4%). Standard deviation is based on three measurements. The EPR spectrum of the bis-(2,2,5,5-tetramethyl-3-imidazoline-1-oxyl-4-yl) disulphide in phosphate-buffered saline (PBS) at 298 K consists of an isotropic triplet pattern (Supplementary Figure 1) with a hyperfine coupling a N value of 15.8 G that is similar to reported values for analogous nitroxide probes where no aggregation was present46,47. In contrast, OmpF-S-S-NO gave a broad anisotropic EPR spectrum with no isotropic component, and is similar to that reported for 5-DSA in lipid bilayers or cholesterol aqueous solutions48. This EPR spectrum indicates hindered rotation of the nitroxide probe49 after binding to the OmpF mutant (OmpF-S-S-NO), and demonstrates successful binding of the bis-(2,2,5,5-tetramethyl-3-imidazoline-1-oxyl-4-yl) disulphide to the modified OmpF mutant (Fig. 2a).

Fig. 2 Characterisation of stimuli-responsive OmpF Panel. a EPR spectra of bis-(2,2,5,5-tetramethyl-3-imidazoline-1-oxyl-4-yl) disulphide-labelled OmpF-M experimental (black) and simulated (blue) and b bis-(2,2,5,5-tetramethyl-3-imidazoline-1-oxyl-4-yl) disulphide-labelled OmpF-M in 1% OG incubated with 10 mM DTT experimental (black) and simulated (blue). c Normalised FCS autocorrelation curves for SAMSA-CF in PBS (black), SAMSA-CF in 1% OG (blue) and OmpF-S-S-CF in 1% OG (Red). Dotted line—experimental autocorrelation curves, full line—fit. d SAMSA-CF release kinetics from OmpF-M in 30 mM GSH, 1% OG, as measured by FCS and analysed with a two-component fit. Error bars show standard deviations from 60 measurements Full size image

After exposure of OmpF-S-S-NO to 10 mM DTT an isotropic EPR spectrum (a N value of 15.9 G) characteristic of the freely rotating spin probe was observed (Fig. 2b). This clearly demonstrates that the nitroxide spin probe that is bound to thiol groups of the OmpF-M under oxidative conditions is cleaved in a reductive environment.

SAMSA-CF (Thermo Fischer Scientific) was selected as a molecular cap because its size (molecular weight 521.49 Da) was expected to block the OmpF-M pore, and because of its ability to form cleavable disulphide bonds50. Thus, attachment of SAMSA-CF to OmpF-M introduces a stimuli-responsiveness to the pore, and therefore to the polymersome membrane when OmpF-S-S-CF is inserted. In addition, the fluorescent properties of SAMSA-CF allow pore modification to be analysed by a combination of sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) and fluorescence correlation spectroscopy (FCS).

LC-MS-MS analysis of the peptide fragments indicated a high labelling degree of OmpF-M (81 ± 31%). In addition, a fluorescent band appeared in the SDS-PAGE gel when SAMSA-CF was conjugated to OmpF-M, whereas the OmpF wild type did not interact with the fluorophore; this fluorescent band supports the formation of OmpF-S-S-CF (Supplementary Figure 2). To mimic the intracellular reductive environment, where the GSH concentration is kept at a constantly high level (10 mM GSH) by cytosolic enzymes51, such as glutathione reductase, we studied the behaviour of the reduction-responsive molecular caps in a similar environment. Because of the absence of a steady-state concentration and constant regeneration of GSH, we used 30 mM GSH to mimic the intracellular steady state of GSH. In SDS-PAGE the fluorescent band disappeared when the OmpF-S-S-CF was mixed with GSH, indicating successful cleavage of the molecular cap under reductive conditions (Supplementary Figure 2).

The binding of SAMSA-CF to OmpF-M cysteine residues was also evaluated by FCS, because it allows the determination of diffusion coefficients, which are correlated to possible interactions of the fluorescent molecules with supramolecular assemblies, such as polymersomes, liposomes and nanoparticles in the pico- to nanomolar concentration region9,21,47,48,49,50,51,52. We compared the molecular brightness and diffusion times of SAMSA-CF in PBS (pH 7.4), SAMSA-CF in 1% OG PBS (pH 7.4) and SAMSA-CF bound to OmpF (OmpF-S-S-CF) in 1% OG PBS (pH 7.4) (Fig. 2c). A labelling efficiency of an average of two SAMSA-CF molecules per monomer was calculated by comparing the molecular brightness (counts per molecule, CPM in kHz) of SAMSA-CF (2.2 ± 0.7 kHz) with that of protein bound to SAMSA-CF (4.8 ± 0.6 kHz) (Fig. 2c). Standard deviations in molecular brightness are based on individual measurements of the same probe (n = 60). In contrast, wild-type OmpF treated similarly to the cysteine mutant OmpF-M, did not present any fluorescence after purification, and there was therefore no binding of SAMSA-CF to OmpF-WT. To determine the kinetics of OmpF pore opening, we used FCS to evaluate the cleavage of SAMSA-CF from labelled OmpF-M upon addition of 30 mM GSH at pH 7.4 by FCS, and analysed the results by a two-component fit. Due to cleavage of the disulphide bonds between the dye and the OmpF-M, the percentage of the free dye increased over time to 85 ± 9%, with a plateau after 1 h (Fig. 2d, Supplementary Figure 3). Standard deviations in the percentage of the free dye are based on individual measurements of the same probe (n = 60) during a set time point.

Catalytic enzyme-polymersome nanocompartments with protein gates

The effect of different concentrations of individual components on the functionality of the final system has already been reported for the insertion of OmpF wild type into PMOXA-PDMS-PMOXA polymersomes and enzyme encapsulation within the inner cavity53. Here we used the optimised conditions and adapted them for the modified OmpF and our AOs. PMOXA 6 -PDMS 44 -PMOXA 6 copolymers spontaneously self-assembled in the presence of HRP, HRP and OmpF-S-S-CF, or HRP and OmpF-S-S-NO, and hollow spherical compartments were identified by cryo-TEM (Fig. 3a, Supplementary Figure 4). These spherical polymer assemblies were demonstrated by light scattering to be polymersomes with: R H of 99 ± 2 nm for HRP-loaded polymersomes containing OmpF-S-S-CF, R H of 89 ± 4 nm for polymersomes loaded with HRP and equipped with OmpF-SH, and R H of 101 ± 1 nm for HRP-loaded polymersomes (Supplementary Tables 1 and 2). Standard deviations were determined based on Pearson’s coefficient of the correlation function and the Guinier fitted one. The polymersome architecture was not affected by 30 mM GSH, with structural parameters ρ (ρ = R G /R H ) values in the range 0.90–0.96, which confirmed a hollow sphere morphology54 (Supplementary Tables 1 and 2). HRP-loaded polymersomes, HRP-loaded polymersomes equipped with OmpF-SH, and HRP-loaded polymersomes equipped with OmpF-S-S-CF all preserved their size and did not aggregate after 2 weeks storage at 4 °C in the dark (Supplementary Figures 5–7).

Fig. 3 Characterisation of stimuli-responsive catalytic nanocompartments. a Cryo-TEM micrographs of: (left) polymersomes loaded with HRP and equipped with OmpF-SH, (middle) polymersomes loaded with HRP and equipped with OmpF-S-S-CF, and (right) polymersomes loaded with HRP without OmpF. Scale bar = 100 nm. b Normalised FCS autocorrelation curves of SAMSA-CF in PBS (black) and OmpF-S-S-CF in the membrane of polymersomes (blue). Dotted line = experimental autocorrelation curves, solid line = fitted curve. Curves normalised to 1 to facilitate comparison. c Left panel: EPR spectrum of bis-(2,2,5,5-tetramethyl-3-imidazoline-1-oxyl-4-yl) disulphide-labelled OmpF reconstituted in PMOXA-PDMS-PMOXA polymersomes (black line). c Right panel: bis-(2,2,5,5-tetramethyl-3-imidazoline-1-oxyl-4-yl) disulphide-labelled OmpF reconstituted in PMOXA-PDMS-PMOXA polymersomes and incubated with 10 mM DTT experimental (black line) and simulated (blue line). d Amplex Ultra Red conversion of HRP-loaded polymersomes: immediately after addition of 30 mM GSH (left), and 1 h after addition of 30 mM GSH (right). OmpF-S-S-CF equipped HRP-loaded polymersomes (green) and OmpF-SH equipped HRP-loaded polymersomes (blue). Error bars present standard deviations in activity between three separately prepared catalytic nanocompartments (n = 3) Full size image

Insertion of channel proteins into enzyme-loaded PMOXA 6 -PDMS 44 -PMOXA 6 polymersomes is critical for in situ activity of the encapsulated enzyme, because the channels allow substrates and products of the enzymatic reaction to pass through the membrane. As OmpF is a pore protein, its functionality is independent of its orientation inside the membrane, and the channel porin mediates the flow of molecules up to 600 Da.

We evaluated OmpF-S-S-CF and OmpF-S-S-NO insertion into the polymersome membrane using FCS and EPR, respectively. A diffusion time of τ d = 2573 ± 960 µs was obtained by FCS for polymersomes with reconstituted OmpF-S-S-CF, indicating that the modified protein gates were successfully inserted into the polymer membranes (free OmpF-S-S-CF in 1% OG has τ d = 588 ± 261 µs). Standard deviation of the diffusion times is acquired from individual measurements (n = 60). By comparing the molecular brightness of the free fluorophore (CPM = 2.2 ± 0.7 kHz) and the OmpF-S-S-CF equipped polymersomes (CPM = 18.9 ± 11.1 kHz), it was calculated that there were five OmpF-S-S-CF porins/polymersome; these values are similar to those reported previously for wild-type OmpF36 (Fig. 3b).

HRP-loaded polymersomes containing OmpF-S-S-NO produced a broad EPR spectrum (Fig. 3c), indicative of low mobility, a result similar to that reported for 5-DSA and 16-DSA inserted in polymersomes membranes55. However, when these HRP-loaded polymersomes containing OmpF-S-S-NO were exposed to reductive conditions (10 mM DTT), an isotropic EPR spectrum (a N = 15.9 G) was observed superimposed on the broad peak, indicating successful cleavage of some of the nitroxide spin probe from the OmpF (Fig. 3c).

Stimuli-responsiveness of the catalytic nanocompartments

The effect of an external stimulus on the functionality of the HRP-loaded polymersomes equipped with OmpF-S-S-CF was evaluated by their response to the addition of 30 mM GSH. The fluorescent signal associated with formation of a resorufin-like product (RLP) during the in situ enzymatic reaction in the presence of Amplex Ultra Red (AR) as a substrate for HRP was measured spectroscopically56. Enzymatic turnover of the AR substrate was significantly lower with HRP-loaded polymersomes equipped with OmpF-S-S-CF (by up to 36±4%) compared to HRP-loaded polymersomes equipped with OmpF-SH, suggesting that the molecular cap is sufficient to reduce the passage of small molecules through the pore. Note that the very low activity of HRP-loaded polymersomes without inserted OmpF was taken into account for background correction. Standard deviation is based on three measurements of separately prepared catalytic nanocompartments. Addition of 30 mM GSH to the system increased the activity of HRP-loaded polymersomes equipped with OmpF-S-S-CF up to that of HRP-loaded polymersomes equipped with OmpF-SH. This indicates that reduction of the disulphide bridge between the attached SAMSA-CF cap and cysteine residues of the OmpF-M successfully restored the OmpF-M pore permeability for the substrate of the enzyme by releasing the molecular cap (Fig. 3d, Supplementary Figures 8 and 9).

Nanocompartments as stimuli-responsive AOs

Here we have gone a step further by developing stimulus-triggered AOs, whose functionality is modulated by the responsiveness of modified OmpF porins inserted in the membrane of the catalytic nanocompartments. Previously designed AOs successfully overcame the first barrier of cell membranes and escaped from endosomes17. As PMOXA-PDMS-PMOXA polymersomes are stable at acidic pH27,36, we consider that this will favour a successful lysosomal/endosomal escape during the recycling of lysosomes and endosomes.

Possible internalisation mechanisms of various PMOXA-PDMS-PMOXA-based polymersomes and their high cytocompatibility in various cell lines have already been reported17,57,58,59. Here, we evaluated the cytocompatibility of the biomimetic AOs by testing their cellular toxicity using the 3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulphophenyl)-2H-tetrazolium (MTS) assay before studying their intracellular activation and enzymatic activity. Notably, their biocompatibility at the cellular level was shown by the absence of any decrease in viability in HeLa cells even after 48 h (i.e. polymer concentration ranging from 0.25 to 0.75 mg ml−1) (Supplementary Figure 10).

In order to study cellular internalisation and intracellular localisation, we first conjugated HRP with Atto488 (HRP-Atto488) and Atto647 (HRP-Atto647), respectively (Supplementary Figure 11). Then we encapsulated labelled-HRP inside the cavity of polymersomes, polymersomes equipped with OmpF-S-S-CF, and polymersomes equipped with OmpF-SH. Cellular uptake assays in HeLa cells indicated successful internalisation resulting in a particulate intracellular staining pattern with increasing intensity in a time-dependent manner from 8 to 24 h (Fig. 4a, Supplementary Figures 12 and 13). The quantitative analysis indicates that after 24 h AOs did not co-localise with early endosomes or lysosomes, confirming successful intracellular endosomal escape (Supplementary Figure 14)17. Localised HRP-Atto488 signals confirmed the intracellular integrity of the polymersomes. In sharp contrast, if cells were treated with a membrane disrupting agent (i.e. 0.1% saponin) (Supplementary Figure 15), polymersome membranes were affected and resulted in an intracellular cytoplasmic distribution of HRP-Atto488.

Fig. 4 Cellular uptake and intracellular activation of AOs. a Confocal fluorescence micrographs of HeLa cells showing cellular uptake of fluorescently labelled HRP-loaded polymersomes and AOs loaded with fluorescently labelled HRP. Scale bar: 10 µm. b Cellular uptake and intracellular activation of fluorescently labelled HRP-loaded polymersomes and fluorescently labelled HRP-loaded AOs. Blue signal: Hoechst 33342 nucleus stain. Grey signal: CellMask Deep Red Plasma membrane stain. Green signal: Atto488 HRP. Red signal: resorufin-like product (RLP). Scale bar 20 µm Full size image

The capacity of the AOs to act within target cells in a stimuli-responsive manner was investigated by using a combination of confocal laser scanning microscopy (CLSM) and flow cytometry to evaluate their potential to respond to increased intracellular GSH levels. HeLa cells were incubated with HRP-loaded polymersomes without OmpF or with HRP-loaded polymersomes equipped with either OmpF-S-S-CF (AOs) or with OmpF-SH. Extracellular polymersomes were removed by washing before imaging the intracellular activity of AOs. Cells were incubated with a 1:1 substrate mixture of H 2 O 2 and AR to allow the intracellular deposition, and finally conversion of AR into its RLP by AOs. Note that both hydrogen peroxide and AR pass through the cellular membrane via passive partitioning, while they do not penetrate the membrane of polymersomes (Supplementary Figure 9). In contrast to untreated cells, or those incubated with HRP-loaded polymersomes without OmpF, a significant increase of intracellular fluorescence was observed with AOs equipped with OmpF-S-S-CF or OmpF-SH (Fig. 4b, Supplementary Figure 16). A similar trend was observed when AR turnover was quantified by flow cytometry (Supplementary Figure 17). The strong fluorescent signal for AOs based on HRP-loaded polymersomes equipped with OmpF-S-S-CF confirmed successful intracellular cleavage of the molecular cap attached to OmpF-M, and subsequent activation of the AOs within the intracellular environment of the HeLa cells (Supplementary Figure 17).

In vivo activity of stimuli-responsive AOs

As a step further to obtaining insight into their safety, tolerability and performance in vivo, AOs were studied in a ZFE model. ZFEs were selected, because of their recognition as a complementary vertebrate animal model for applications, such as compound screening in drug discovery, toxicological studies and recombinant disease models60,61,62. Compared to rodent in vivo models, the ZFE offers unique advantages: (i) high reproducibility, (ii) low costs, (iii) high level of genetic homology to humans, (iv) availability of transgenic lines and (v) most importantly for the evaluation of AO, optical transparency. Due to their optical transparency, ZFE provide the possibility of imaging fluorescently-tagged objects and fluorescent processes in vivo at a high resolution over time63 (Supplementary Figure 18). Our approach offers the possibility of gaining detailed insight into the circulation behaviour of AOs and subsequent enzymatic reactions as we reported recently for nano-particulate drug delivery systems in vivo64. In order to follow the biodistribution of AOs, we injected intravenously via the duct of Cuvier HRP-Atto488-loaded polymersomes with membranes equipped with OmpF-S-S-CF or with OmpF-SH, respectively. No acute toxicity, such as change in behaviour i.e. mobility, seizures, heart failure or other toxic effects such as malformations, denaturation of tissue fluids or yolk mass was observed in ZFE injected with AOs after 24 h. ZFE analysed 2 h post intravenous injection of all types of AOs containing Atto488 conjugated HRP showed a distinct fluorescent staining pattern (Supplementary Figure 19) in the posterior cardinal vein region, and we hypothesise that polymersomes are recognised by the ZFE early immune system and are subsequently taken up by macrophages65. The remarkable recognition of polymersome-based AOs by the ZFE immune system was confirmed by the colocalisation of AOs loaded with Atto647-conjugated HRP (Atto647-HRP) injected into transgenic ZFE specifically expressing eGFP in macrophages (Fig. 5a, Supplementary Figure 20). In strong contrast to AOs loaded with Atto647-HRP, the free Atto647-HRP enzyme did not show significant macrophage colocalisation after 24 h, even when Atto647-HRP was injected at concentrations of 0.2 mg ml−1 (Supplementary Figure 21). Notably, only macrophages in circulation were targeted and not tissue resident macrophages (i.e. star shaped).

Fig. 5 Internalisation and activity of AOs in macrophages in vitro and in vivo. a Localisation of AOs in ZFE. Lateral view of the ZFE injected with HRP-Atto647-loaded AOs equipped with OmpF-S-S-CF. Arrowheads: Localisation of AOs. Blue signal: ZFE melanocytes. Green signal: GFP macrophages. Red signal: Atto647-loaded AOs. b Phagocytosis of AOs by human macrophage differentiated THP-1 cells in vitro. Qualitative (inset) and quantitative analysis of macrophage differentiated THP-1 cells incubated with AOs loaded with Atto488-labelled HRP without addition of inhibitors (left), and in the presence of phagocytosis inhibitor cytochalasin B (Cyto B) (right). Blue signal: Hoechst 33342 nucleus stain. Green signal: Atto488 HRP. Scale bar inset 20 µm. c Quantification of AOs in the presence of different pharmacological pathway inhibitors by flow cytometry: polyinosinic acid (Poly(I:C)), colchicines, cytochalasin B (Cyto B) and sodium azide (NaN 3 ). d In vivo ZFE biodistribution and activity of AOs—lateral view of the ZFE cross-section. Blue signal: ZFE melanocytes. Green signal: HRP-Atto488. Red signal: Resazurin-like product (RZLP). Arrows show regions of enzymatic activity of AOs Full size image

Once cellular internalisation of AOs by the early immune system of ZFE was successful in vivo, we explored the uptake rate, exact intracellular localisation and internalisation mechanisms of AOs in immune cells in vitro by using human macrophage differentiated THP-1 cells. AOs internalisation started as early as 30 min, and a strong internalisation by immune cells was achieved after 3 h (Supplementary Figure 22), with increasing uptake rates at higher time points. As THP-1 cells are immature macrophages with reduced phagocytotic capacity, a higher uptake rate of AOs is possible for mature (primary) macrophages in vitro and in vivo.66 Importantly, all macrophage uptake studies were performed in the presence of serum proteins to mimic physiological conditions in vivo because opsonisation of nanoparticles by serum proteins can highly influence their interaction with cells.67 To obtain a mechanistic understanding of the internalisation process, THP-1 macrophages were pre-treated with different pharmacological pathway inhibitors.67 We used inhibitors with specific inhibition profiles: (i) polyinosinic acid to block scavenger receptors, (ii) colchicine to inhibit pinocytosis, (iii) cytochalasin B as phagocytosis inhibitor and (iv) sodium azide to inhibit all energy-dependent uptake processes. Cells not incubated with Atto488 HRP-loaded AOs served as a control. A 1.28-fold increase in the mean fluorescence intensity (MFI) was observed by flow cytometry analysis of the cells incubated with Atto488 HRP-loaded AOs for 6 h, which indicates internalisation of AOs by THP-1 macrophages. The uptake of AOs by macrophages was significantly inhibited by cytochalasin B (a 0.13-fold increase in MFI) and in a lower degree by sodium azide (0.43-fold increase in MFI), which indicates an energy-dependent phagocytotic internalisation process (Fig. 5b, c). On the contrary, polyinosinic acid did not inhibit the AOs uptake, suggesting little or no involvement of the scavenger receptor in the internalisation mechanism of AOs (Fig. 5c).

The internalisation process analysed by CLSM using LysoTracker™ Red DND-99 as a reporter for the lysosomal compartments indicates that AOs co-localise with lysosomal compartments during their internalisation process (Supplementary Figure 23). Interestingly, we could not detect a lysosome signal (lysotracker) 24 h after incubation of macrophages with AOs, suggesting the presence of an intracellular lysosomal escape mechanism once the AOs are taken-up by macrophages (Supplementary Figure 23). After internalisation in macrophages, the signals associated with Atto488-HRP-loaded AOs in lysosomal compartments changed to larger intracellular vesicular signals. This suggests an expansion of the AO-bearing lysosomal compartments before the AOs are released into the cytosol. For an exact mechanism by which AOs escape the lysosomal compartment and interact with cellular membranes, further investigations are planned but they are beyond the scope of this study.