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Article Open Access Published: 25 February 2019
Polymeric Engineering of Nanoparticles for Highly Efficient Multifunctional Drug Delivery Systems
Beatrice Fortuni , Tomoko Inose, Monica Ricci, Yasuhiko Fujita, Indra Van Zundert,
Akito Masuhara, Eduard Fron, Hideaki Mizuno, Loredana Latterini, Susana Rocha &
Hiroshi Uji-i
Scientific Reports 9, Article number: 2666 (2019)
14k Accesses 91 Citations 6 Altmetric Metrics
Abstract
Most targeting strategies of anticancer drug delivery systems (DDSs) rely on the
surface functionalization of nanocarriers with specific ligands, which trigger the
internalization in cancer cells via receptor-mediated endocytosis. The
endocytosis implies the entrapment of DDSs in acidic vesicles (endosomes and
lysosomes) and their eventual ejection by exocytosis. This process, intrinsic to
eukaryotic cells, is one of the main drawbacks of DDSs because it reduces the
drug bioavailability in the intracellular environment. The escape of DDSs from
the acidic vesicles is, therefore, crucial to enhance the therapeutic performance
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4/27/23, 10:19 Page 1 of 51
at low drug dose. To this end, we developed a multifunctionalized DDS that
combines high specificity towards cancer cells with endosomal escape
capabilities. Doxorubicin-loaded mesoporous silica nanoparticles were
functionalized with polyethylenimine, a polymer commonly used to induce
endosomal rupture, and hyaluronic acid, which binds to CD44 receptors,
overexpressed in cancer cells. We show irrefutable proof that the developed
DDS can escape the endosomal pathway upon polymeric functionalization.
Interestingly, the combination of the two polymers resulted in higher endosomal
escape efficiency than the polyethylenimine coating alone. Hyaluronic acid
additionally provides the system with cancer targeting capability and
enzymatically controlled drug release. Thanks to this multifunctionality, the
engineered DDS had cytotoxicity comparable to the pure drug whilst displaying
high specificity towards cancer cells. The polymeric engineering here developed
enhances the performance of DDS at low drug dose, holding great potential for
anticancer therapeutic applications.
Introduction
Over the last few decades, the engineering of nanoparticles has given rise to
significant breakthroughs towards the employment of nanomaterials in
biomedical applications, such as cancer therapy, (bio-) chemical sensing, and
bio-imaging . In particular, mesoporous silica nanoparticles (MSNPs) have
been widely applied as promising anticancer drug nanocarriers thanks to their
biocompatibility, high loading capacity, chemical stability and straightforward
synthesis/surface functionalization . Unlike some other nanocarriers, MSNPs
have not been translocated into the clinical stage yet . However, the reasonable
biocompatibility accomplished in vivo is extremely promising for a proximate
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Food and Drug Administration (FDA-) approval .
To promote the specific internalization of nanoparticles to certain cancer cells
(cancer targeting), many strategies have been developed so far. These methods
are mainly based on the employment of specific ligands, which can bind to
receptors overexpressed in tumor cells and trigger particle internalization via
endocytosis . In this context, hyaluronic acid (HA) has gained increasing
attention as targeting ligand due to its high affinity with CD44, a glycoprotein
receptor overexpressed in many solid tumor cells (e.g. lung, breast, pancreatic,
renal tumor), in metastasis, as well as in cancer stem cells . As being one of the
main constituents of the extracellular matrix, HA exhibits high biocompatibility,
which has enabled its FDA-approval for medical and cosmetic use . The
harmlessness of HA, allied with its effective targeting capability, encouraged its
employment to selectively internalize HA-functionalized materials (HA-
materials) in CD44-overexpressing cancer cells via receptor-mediated
endocytosis .
In spite of the well-achieved cell-specific internalization, the control of the
particle fate after overpassing the plasma membrane remains challenging, and
existing strategies are still limited. In eukaryotic cells, external materials (such as
nutrients, protein and lipids, as well as nanoparticles), taken up via endocytosis,
are normally sorted out in endocytic vesicles (endosomes and lysosomes) and
can eventually be ejected to the extracellular matrix via exocytosis . Previous
reports have shown that non-coated MSNPs co-localize with the endo-
/lysosomes in the early stage of incubation , and are quickly
exocytosed, following this pathway . Similarly, HA-coated MSNPs are
internalized via CD44-mediated endocytosis and are subjected to same
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endocytic system, ending up in the acidic cellular compartments within few
hours of incubation , and being ejected via exocytosis within 48 h . The
endo-/exocytosis process represents one of the main hindrances of the DDSs in
light of the limited cargo release in the intracellular environment. The low
lysosomal pH (4.5–5.5 for normal cells and 3.5–5 for cancer cells) and the
strong enzymatic activity might lead to drug degradation, possibly inhibiting its
pharmaceutical activity . The therapeutic efficiency can be further decreased
by the fast exocytosis of the nanocarriers . As the drug release normally
occurs by slow diffusion, the DDS can be exocytosed to the extracellular matrix
before releasing all its cargo, contributing to the low therapeutic performance
(forcing the use of higher drug dose), as well as to chemotherapy side effects.
Despite the major consequences in terms of therapy efficiency, the intracellular
route of nanocarriers is often neglected in the development of novel DDS, and
strategies to enable the escape from this endocytic route are very limited. To
this end, the employment of cationic polymers, in particular polyethylenimine
(PEI), is a promising strategy, as it is non-immunogenic and easier to scale up,
compared to other agents, such as viral proteins and synthetic fusogenic
peptides . PEI is already widely used in DNA transfection for promoting the
release of genetic material from the acidic vesicles and, thus, facilitating the
incoming to the nucleus . The use of DNA-PEI polyplexes, instead of pure
DNA, was demonstrated to improve the gene expression efficiency up to 100-
fold . This enhanced gene expression can be associated to the so-called
“proton sponge effect” of PEI . Most specifically, thanks to the protonation of
tertiary amines, PEI exhibits high buffering capability at low pH, promoting an
influx of protons inside the acidic cellular compartments via ATPase proton
pumps and the consequent rupture of the organelle membrane due to an
osmotic imbalance. The proton sponge effect of PEI is a generally accepted
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hypothesis in literature, however, it is important to mention that this concept is
still heavily debated .
Since the action mechanism of most anticancer drugs, e.g. doxorubicin (Dox), is
based on its intercalation into DNA and complex formation with DNA-associated
enzymes , the same approach can be used to enhance the nuclear delivery of
anticancer drugs. The main hindrance for the application of PEI on DDSs is its
cytotoxicity, which can be, howbeit, drastically reduced by using a low molecular
weight (0.5–5 kDa) . So far, PEI has been used to functionalize MSNPs for
the successful delivery of either siRNA/DNA or siRNA/doxorubicin to HEPA-1
and KB-V1 cells, respectively . In these studies, the endosomolytic activity
of the PEI layer was assumed but not verified. On the other hand, Yanes et al.
demonstrated that the addition of a PEI layer can slow down the exocytosis rate
of MSNPs, although no investigation on the intracellular distribution of the
nanoparticles was performed . To the best of our knowledge, a study on the
intracellular sorting of PEI-coated nanocarriers, which provides an evidence of
their endosomal escape, has never been reported.
In this manuscript, we propose a facile method to functionalize mesoporous
silica nanoparticles with a polymeric bilayer, which simultaneously combines the
active targeting action of HA and PEI-mediated endosomal escape (Fig. 1). For
therapeutic applications, any anticancer drug can be loaded in the particles.
Here, we use Dox-loaded MSNPs and show that the combination of active
targeting, endosomal escape and controlled drug release results in high therapy
efficiency. The method presented paves the way for the development of the
next generation highly efficient DDSs.
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Figure 1
Multifunctional drug delivery system based on MSNPs: particle synthesis and
cellular trafficking. (a–c) Preparation of HAPEI-MSNP_Dox: (a) encapsulation of
doxorubicin (Dox) inside mesoporous silica nanoparticles (MSNP_Dox); (b) coating
with polyethylenimine (PEI) layer (PEI-MSNP_Dox); (c) surface grafting with
hyaluronic acid (HA) (HAPEI-MSNP_Dox). (d–f) Cellular uptake and intracellular
trafficking: (d) particle interaction with the plasma membrane via CD44-HA site-
specific binding; (e) HAPEI-MSNP_Dox uptake via receptor-mediated endocytosis
and wrapping in endosomes; (f) rupture of the endosomal membrane upon proton
sponge effect of PEI and drug release into the cytoplasm. A schematic illustration
representing functions and chemical interactions of each component is reported at
left-bottom of the figure.
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Results and Discussion
Preparation and characterization of multifunctional MSNPs
Due to their popularity as highly stable, low-cost and reasonably biocompatible
nanocarriers, mesoporous silica nanoparticles (MSNPs) were chosen as model
of nanoparticle for the application of the polymeric coating here proposed .
MSNPs were synthetized using the biphase stratification method developed by
Shen et al., that yields particles with a pore size of ~2.8 nm . Transmission
electron microscopy (TEM) images of uncoated MSNPs clearly show a uniform
mesoporous frame (Fig. 2a). The particles exhibit size and shape homogeneity,
with no observable aggregates. As depicted in Fig. 2b, the mean diameter was
estimated to be 120 nm. After the synthesis, MSNPs were loaded with
rhodamine B (RhodB) or fluorescein isothiocyanate (FITC) for monitoring cellular
uptake/trafficking, and with Dox for testing the drug release and the therapeutic
effect in cancer mammalian cells. The successful loading of dye/drug inside the
pores was verified by fluorescence microscopy (Supplementary Fig. S1a–c).
Figure 2
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Characterization of MSNPs and their surface modifications. (a) TEM image of bare
MSNPs. (b) Size distribution of the MSNPs (Gauss distribution in red fitting). (c) Zeta
potential measurements of MSNPs, PEI-MSNPs and HAPEI-MSNPs. (d–f) FE-SEM
images of MSNPs, PEI-MSNPs and HAPEI-MSNPs, respectively.
In order to provide the DDS with endosomal escape capability, MSNPs were
coated with PEI (~1.3 kDa). Besides their biocompatibility and high loading
capability, MSNPs offer a negatively charged surface which facilitates any kind
of electrostatic interaction-based functionalization. At physiological pH, primary
and secondary amines of PEI are protonated (pK 8–10, depending on the
molecular weight of the polymer) , whereas ~50% of hydroxyl groups on a
silica surface are deprotonated (pK ≈ 6.8) . This ionization percentage enables
the formation of a PEI shell on the MSNP surface via electrostatic interaction.
The presence of the PEI layer on MSNPs was demonstrated by the drastic
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change in the zeta potential of the particles after the coating (from −38.2 mV to
+37.7 mV, Fig. 2c).
Thanks to the abundance of amino groups, the presence of PEI on the surface
of MSNPs allowed for a straightforward binding of the targeting agent, HA,
without any extra chemical modification. The carboxylic group of HA was
covalently linked to the amino group of PEI via carbodiimide crosslinking
reaction . The decrease of the electrokinetic potential from +37.7 to +4.2 mV
after HA grafting onto the PEI coating indicates the successful functionalization
of the particles with HA (Fig. 2c). Considering such a change of the
electrokinetic potential upon HA grafting, an effect on the charge-based PEI
coating cannot be excluded. On the other hand, no attachment of HA would
occur without the presence of a PEI layer on the surface, suggesting that the
electrostatic interactions between PEI and the silanol groups endure the HA
grafting process.
The presence of the polymeric layers was confirmed using high resolution field-
emission scanning electron microscopy (FE-SEM). Representative images of
bare MSNPS, MSNPS coated with PEI (PEI-MSNPs) and MSNPs functionalized
with a bilayer of PEI and HA (HAPEI-MSNPs) are shown in Fig. 2d–f,
respectively. While the PEI layer is barely visible in the FE-SEM images of PEI-
MSNPs (Fig. 2e), after conjugation with HA, the edge contrast increases,
enabling an easier visualization of the polymeric layers in Fig. 2f. It is important
to note that during image acquisition the coating collapsed and detached from
the silica surface as a consequence of exposure to high accelerating voltages
(30 kV). Therefore, the thickness of the layers visible in Fig. 2e,f does not
correspond to the exact thickness of the shells. The halo displayed in Fig. 2e,f
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was never observed for bare MSNPs (additional FE-SEM images of bare MSNPs
and HAPEI-MSNPs in Supplementary Fig. S2).
Cellular uptake: HA-mediated active targeting
In order to evaluate the targeting efficiency and the cell specificity of the
external functionalization with HA, we monitored the cellular uptake of the
different particles into two mammalian cell lines. Most specifically, RhodB-
loaded MSNPs with different coatings were added to A549 cells (CD44-
overexpressing cells, derived from human lung carcinoma) and NIH3T3
(mouse embryonic fibroblasts, lowly expressing CD44 receptors, defined as
CD44-negative or CD44-inactive cells) .
Figure 3 shows typical fluorescence images of both cell lines after 3 h of
incubation with the different nanoparticles, with no coating (MSNPs), only a PEI
layer (PEI-MSNPs) or functionalized with both HA and PEI (HAPEI-MSNPs). In
order to quantify the cellular uptake, the plasma membrane was stained with a
membrane-incorporating fluorescent dye (DiO, shown in green in Fig. 3). While
there was a minimal amount of bare MSNPs detected inside the cells
(Fig. 3a,d,g), PEI-MSNPs show a 2-fold increase in cellular uptake,
independently of the cell line (Fig. 3b,e,g). This is in agreement with previously
published results and is linked to the positive charge of PEI, which boosts
electrostatic interactions with the electronegative plasma membrane and
facilitates particle internalization. The addition of HA minimizes the surface
charge of the PEI coated nanoparticles and reduces this effect. Consequently, in
NIH3T3 cells, the uptake of HAPEI-MSNPs is similar to that of bare MSNPs
(Fig. 3f). Remarkably, incubation of A549 cells with HAPEI-MSNPs results in a
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10-fold increase on the amount particles detected inside the cell (compared
with bare MSNPs, Fig. 3c and g). The drastic discrepancy in HAPEI-MSNP
uptake rate between NIH3T3 (Fig. 3f) and A549 cell lines (Fig. 3c) proves that
the HA functionalization provides our DDS (HAPEI-MSNP) with high specificity
towards CD44-overexpressing cancer cells.
Figure 3
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Influence of surface modification on the cellular uptake of MSNPs. (a–f)
Fluorescence images of A549 and NIH3T3 cells after incubation with
MSNPs_RhodB, PEI-MSNPs_RhodB and HAPEI-MSNPs_RhodB for 3 h. RhodB-
loaded particles are shown in orange; DiO-stained plasma membrane is colored in
green. The central panel displays an xy-plane within the cells, while the right and
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bottom panels show the yz and xz projections, respectively. (g) Mean intensity of
the RhodB signal per µm of cell (n = 4 for each condition), error bars indicate ± SD,
with ns = (p > 0.05), *(p < 0.05), **(p < 0.01) and ***(p < 0.001).
Intracellular trafficking: PEI-induced endosomal rupture
Previous reports have shown that bare and HA-functionalized MSNPs traffic
through the endocytic pathway, ending up into lysosomes and, eventually, being
exocytosed . In order to evaluate the effect of both PEI coating alone
and its combination with HA on the endosomal trafficking, A549 cells were
incubated with FITC-loaded particles for 3, 24 and 48 h. It is important to
mention that after 3 h of incubation, the medium was refreshed to discard the
excess of particles, preventing further internalization. After the incubation
period, lysosomes were stained using LysoTracker Red®, a fluorophore linked to
a weak base that is only partially protonated at neutral pH and is fluorescent
only in acidic environments. Cells were imaged by fluorescence microscopy and
the co-localization between the fluorescence signal of FITC-loaded
nanoparticles and LysoTracker Red® was determined using the Pearson’s
correlation coefficient (PCC) analysis (PCC threshold values of the current
study are reported in SI, Supplementary Fig. S3). Figure 4 displays
representative images of A549 cells incubated with MSNPs with different
coatings, after 3, 24 and 48 h. The particles are shown in green while the acidic
compartments are presented in red. As a consequence, MSNPs trapped in
endo- or lysosomes are displayed in yellow.
Figure 4
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figure 4
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figure 4
Influence of surface modification on the intracellular trafficking of MSNPs at
different time points. (a–i) Fluorescence images of A549 cells incubated with
MSNPs_FITC (a–c), PEI-MSNPs_FITC (d–f) and HAPEI-MSNPs_FITC (g–i) after 3,
24 and 48 h of incubation. The lysosomes were stained using LysoTracker Red®.
Green channel (FITC-loaded particles), red channel (LysoTracker Red-stained endo-
/lysosomes) and DIC merged images are shown. (j) Co-localization coefficient
between the fluorescence signal of FITC-loaded nanoparticles and the LysoTracker
Red (PCC ± SD plot over time, n = 5). PCC analysis was performed by using MATLAB
software.
After 3 h, MSNPs without any additional surface functionalization co-localized
with the endo-/lysosomes (Fig. 4a). Even after 48 h, all the particles detected
inside A549 cells were co-localized with acidic compartments, indicating that
none of the bare MSNPs taken up by the cell was able to escape the endocytic
pathway (Fig. 4c). Accordingly, the calculated PCC is constant over time (black
line in Fig. 4j). Note that, since the internalization rate of MSNPs is relatively low
comparted to that of HAPEI- and PEI-MSNPs, in order to get an appropriate
comparison study of the intracellular distribution, A549 cells with relatively
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higher MSNPs uptake were selected to perform confocal imaging and
subsequent PCC analysis.
Within a time span of 3 h, the PEI coating does not induce a clear effect on the
intracellular fate of the nanoparticles, with bare MSNPs and PEI-MSNPs
displaying similar intracellular distributions and co-localization coefficients
(Fig. 4a,d,j). In stark contrast, after 24 h, PEI-MSNPs are roughly equally
distributed between cytoplasm and acidic cellular compartments (Fig. 4e). The
associated mean PCC value drastically decreases from 0.64 (3 h) to 0.36 (24 h),
implying a reduced linear correlation between the fluorescence signal of PEI-
MSNPs and endo-/lysosomes. At this time point, a high heterogeneity in the
intracellular localization was observed between different cells, explaining the
large standard deviation (SD) of the mean PCC value. As depicted in Fig. 4f,
after 48 h the majority of PEI-MSNPs are excluded from the acidic
compartments, with PCC value dropping to 0.25. The ability of PEI-coated
particles to escape from the acidic vesicles is attributed to the proton sponge
effect of this polymer, which results in the rupture of the membrane
organelles . It is important to mention that the possible proton sponge effect of
PEI does not change the pH of the endo-/lysosomes , and has no effect on the
staining of these organelles with LysoTracker probes. Consequently, a lower co-
localization with the LysoTracker Red® can be directly linked to endo-/lysosomal
damage and/or rupture.
A similar trend was observed with the multifunctionalized HAPEI-MSNPs. After
48 h the majority of the particles with a HA-PEI bilayer were not co-localized
with acidic cellular compartments (Fig. 4i).
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However, the initial uptake and endosomal escape rate is very different. At 3 h of
incubation, a considerable fraction of HAPEI-MSNPs had already escaped the
acidic vesicles (Fig. 4g, mean PCC = 0.45), indicating an effect of the polymeric
bilayer in the endosomal escape rate (PCC similar to that of PEI-MSNPs after 24
h, Fig. 4j). The fraction of particles co-localizing the acidic compartments
markedly decreased after 24 h, when most HAPEI-MSNPs were found to be no
longer entrapped inside the endo-/lysosomal vesicles (Fig. 4h). After 48 h,
practically all HAPEI-MSNPs were distributed in the cytosol (Fig. 4i, mean PCC =
0.10), indicating a highly effective escape of HAPEI-MSNPs from the acidic
compartments.
The results obtained with fluorescence imaging were further validated by
electron microscopy. For the TEM measurements, cells were incubated with
differently functionalized particles for 3 h and fixed after 24 h (more info in SI,
Supplementary Figs S4 and 5). In agreement with the fluorescence images
acquired at this time point, TEM images show that bare MSNPs were clearly
trapped inside the lysosomes, MSNPs coated with PEI alone were found to be
distributed either in the cytoplasm or inside the endo/lysosomes, and MSNPs
with a polymeric bilayer (PEI and HA) were detected mainly outside of the
lysosomes (Supplementary Fig. S5).
These results constitute the first irrefutable evidence that coating of
nanoparticles with specific polymers induces the rupture of the endo-
/lysosomes and further escape to the cytosol. Both fluorescence and electron
microscopy images demonstrate an evident enhancement/acceleration of the
endosomal escape efficiency with HA-PEI coating compared to using PEI alone.
Further research is necessary to assess the mechanism behind this effect,
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although we speculate that it might be associated either to a faster uptake rate
of the HAPEI-coated particles, thanks to the HA targeting, or, more generally, to
the presence of an additional polymer. At low pH, the inclusion of an extra
polymeric layer can, indeed, increase both the buffering capacity and the
polymeric swelling, contributing to the destabilization of the endo-/lysosomal
membrane .
Drug release in vitro
Thanks to the therapeutic effectiveness towards a wide range of cancers
(carcinomas, sarcomas and hematological cancers) and to its fluorescent
properties , doxorubicin (Dox) was selected as anticancer drug model for the
current work. To be efficient, DDSs should guarantee a stable encapsulation of
the drug, combined with a controlled release at the specific target. For bare
MSNPs, the environmental pH plays a crucial role on the drug release kinetics.
Further information about the mechanism of Dox uptake and release in/from
MSNPs is reported in SI (Supplementary Fig. S9). Gao et al. have shown that the
release rate of Dox in vitro is accelerated at acidic pH, although a relatively
smaller amount can be released at neutral pH as well . In the particle design
proposed here, in addition to confer to the MSNPs active targeting towards
cancer cells and the capability to induce endosomal rupture, the HA-PEI
polymeric bilayer will function as a capping agent, preventing the leakage of the
drug before reaching the intracellular environment. At neutral pH, according to
the pK values of PEI and silica hydroxyl groups , the electrostatic
interactions guarantee a stable attachment of the PEI shell to the particles,
impeding the discharge of Dox in blood circulation. At acidic pH, instead, as the
majority of the hydroxyl groups of the silica particles are protonated, the
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electrostatic interactions are minimized, reducing the capping effect of the
polymeric coating and facilitating the drug release in the cellular acidic
compartments. In this context, Meng and co-workers reported that PEI coating
does not hinder the Dox release at acidic pH .
In order to evaluate the capping effect of the polymeric HA-PEI bilayer proposed
here, the release kinetics of Dox from HAPEI-MSNPs was estimated in vitro. As
depicted in Fig. 5a, functionalization of MSNPs with a HA-PEI bilayer resulted in
particles with no drug release in both neutral (pH 7) and acidic environments
(pH 6 and 4.5). This suggests that the stability of the shell is enhanced by the
external HA layer, which likely hinders the polymer detachment, making the
coating more compact and stable, even at acidic pH, thanks to the amide bond
links to the HA.
Figure 5
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figure 5
Drug release in vitro and in cellulo. (a) Time-dependent in vitro release profile of Dox
from HAPEI-MSNPs_Dox (0–72 h) at pH 4.5 (red circle), pH 4.5 + Hyal-1 (red
triangle), pH 6 (blue circle), pH 6 + Hyal-2 (blue triangle) and pH 7.4 (green triangle)
(each point consists of mean ± SD, n = 3). (b–d) Fluorescence images of Dox
released from HAPEI-MSNPs_Dox inside A549 cells after 3, 24 and 48 h (b-d,
respectively). Dox channel (in red), DIC (gray) and merged images are shown from
left to right, respectively. The contrast of the red channel was kept constant in all
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images.
In the cellular environment, the external HA shell can be degraded by digestive
intracellular enzymes, thereby promoting the discharge of the drug exclusively
within the target cell. The main HA digestive enzymes are Hyaluronidase-1
(Hyal-1), which is normally located inside endosomes and lysosomes, and
Hyaluronidase-2 (Hyal-2), mainly present on the plasma membrane . While
most degradation occurs in the acidic compartments, Hyal-2 can already
degrade the high molecular weight HA into smaller fragments during the ligand-
receptor binding, immediately prior to endocytosis .
Enzyme-mediated HA degradation and subsequent drug release was evaluated
by incubating the Dox loaded HAPEI-MSNPs in different solutions at 37 °C. MES
buffer (pH 6) with Hyal-2 was selected to mimic the extracellular matrix in tumor
tissue, and acetate buffer (pH 4.5) containing Hyal-1 was used to simulate the
late endosomes and lysosomes.
The amount of Dox released at different incubation times (3, 12, 24, 48, 72 h) is
shown in Fig. 5a. While in absence of enzymes and independently of the pH the
percentage of Dox released was negligible, upon enzymatic digestion by Hyal-1
(pH 4.5) or Hyal-2 (pH 6), the release profiles were similar to those of bare
MSNP (Supplementary Fig. S6). Similarly to bare MSNPs, Dox release kinetics
were faster at more acidic pH, which is in agreement with previous reports .
The addition of Hayl-2 to the solution mimicking the extracellular matrix (pH 6)
led to a total release of Dox from HAPEI-MSNPs of 58 ± 3% after 72 h. Notably,
after only 3 h, 15% of the drug had been already released, suggesting that a
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partial digestion of HA on the plasma membrane can facilitate some Dox
release. The addition of Hyal-1 at 4.5 pH (similar to the endo-/lysosomes) turned
out to be the condition with the higher amount of Dox released, reaching a
percentage of 68 ± 1% in 72 h. The Dox release profile from HAPEI-MSNPs in the
presence of hyaluronidase demonstrates that only enzyme-mediated
degradation of the polymeric coating, which occurs exclusively in the cellular
environment, triggers drug release from the particles.
Drug release in cellulo
In order to evaluate drug release kinetics in cellulo, HAPEI-MSNPs loaded with
Dox were added to A549 cells and intracellular Dox release was monitored using
fluorescence microscopy (Fig. 5b–d). When adding pure Dox to cells,
fluorescence could be detected uniformly in the cytoplasmic region after 24 h,
with no signal coming from the cell nucleus (Supplementary Fig. S7). The
absence of fluorescence in the cell nucleus is associated to Dox intercalation
between the DNA base pairs. As reported by several research groups, nuclear
penetration causes a drastic quenching of Dox fluorescence , up to 95%
of its intrinsic emission .
Similar to the pure drug, after 3 h of incubation with Dox-loaded HAPEI-MSNPs,
fluorescence could be detected in the cytoplasm of A549 cells. The weak
dispersed signal in the cytoplasmic area was attributed to a small ratio of Dox
release within the 3 h of incubation (which is in agreement with the results
obtained in vitro in the presence of HA-degrading enzymes). While cells
incubated with the pure drug only show a disperse fluorescence over the whole
cytoplasmic region (Supplementary Fig. S7), when Dox-loaded nanoparticles
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are used, it was possible to observe bright dots in the intracellular environment
(Fig. 5b). These bright dots were attributed to the HAPEI-MSNPs containing
Dox. At longer time intervals (24 and 48 h), the fluorescence signal from Dox
was more intense over the cytoplasm, while the bright spot-like signals arising
from the particles became dimmer (Fig. 5c,d). This suggests that during time
Dox was released from the particles into the intracellular environment (note that
after 3 h the cells were washed, stopping further uptake of any drug and/or
particles). This change in the distribution of Dox fluorescence signal was
observed in all cells (Supplementary Fig. S8) and is in agreement with the
enzyme-mediated release profile obtain in the in vitro experiments.
Anticancer efficiency: cell viability tests
In order to evaluate the efficiency of the newly developed polymer-coated
particles as anticancer DDSs, we monitored the cell viability 72 h after treatment
with free Dox, Dox-loaded HAPEI-MSNPs and empty HAPEI-MSNPs, at different
concentration of drug/particles (Fig. 6).
Figure 6
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figure 6
Anticancer efficiency of Dox-loaded HAPEI-MSNPs. Viability tests of A549 cells
incubated with HAPEI-MSNPs_Dox (wine red line column), free Dox (violet column)
and empty HAPEI-MSNPs (light gray) for 72 h. Aliquots of 2, 4, 6, 8, 10 µL
corresponding to final Dox concentrations of 80, 160, 240, 320, 400 nM and particle
concentration of 20, 40, 60, 80, 100 µg/mL, respectively, were added to 1 mL of cell
culture medium. All data are shown as mean ± SD (n = 3) with ns = (p > 0.05), *(p <
0.05), **(p < 0.01) and ***p < 0.001).
While at low concentration (<40 µg/mL) empty HAPEI-MSNPs are not toxic,
higher concentration leads to a lower cell viability, reaching a minimum of 85%
at 80–100 µg/mL. A similar effect of empty HA-coated MSNPs on the cell
viability was reported by previous studies . In our experiments, it can be23
4/27/23, 10:19 Page 24 of 51
associated either to the massive uptake of HA-coated particles into A549 cells
(as shown in Fig. 3c, above) or to the intrinsic toxicity of PEI itself . Note that
the HAPEI-MSNP uptake is highly specific towards CD44-overexpressing
cancer cells and, consequently, the limited toxicity of the empty particles cannot
be considered of negative impact on the functionality and effectiveness of the
DDS.
The cytotoxicity of Dox-loaded HAPEI-MSNPs is at least as high as the one of
free Dox, indicating an efficient intracellular release and trafficking of the drug.
For free and Dox-encapsulated particles, the cell viability decreases with the
increase of the drug concentration, reaching a mean value of 42 and 20% at
400 nM, respectively. As a general trend, Dox encapsulated in HAPEI-MSNPs
seems to induce higher cell mortality rate in comparison to free Dox, with a
sharper discrepancy at high drug/particle concentration. This enhanced killing
capability might be explained by the well-engineered properties of the DDS
(high uptake, endosomal escape and controlled drug released) allied with the
toxicity co-effect of the nanocarrier itself.
Taken together these results demonstrate that our DDS has great therapeutic
potential, specifically towards CD44-overexpressing cancer cells, having a
comparable, or better, efficiency than free Dox. The high therapy efficiency
achieved at low drug concentrations is strictly related to the fast internalization
rate (HA coating targeting effect), to the improved endosomal release, with
consequent retention of the particles in the cytoplasm (HA-PEI shell inducing
endosomal rupture), and to a controlled drug release overtime (enzymatic
polymeric digestion).
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4/27/23, 10:19 Page 25 of 51
In conclusion, in this work, a polymeric bilayer functionalization of mesoporous
silica nanoparticles was designed for drug delivery-based tumor therapy.
Hyaluronic acid and polyethylenimine layers provide the drug delivery system
with active targeting and endosomal escape capability, simultaneously,
enhancing the therapeutic efficiency. The as-obtained nanoparticles (HAPEI-
MSNPs) turned out to possess an excellent active targeting capability towards
CD44-overexpressing cell (A549). Furthermore, the presence of PEI was
demonstrated to trigger endosomal escape of both PEI-MSNPs and HAPEI-
MSNPs. Unlike previous reports, the endosomal breakout of PEI- and HAPEI-
coated mesoporous silica nanoparticles was unambiguously shown.
Fluorescence data were consistent with the results obtained by electron
microscopy. A time-lapse fluorescence-based investigation showed that after
48 h, multifunctionalized particles were mainly localized in the cytoplasm (data
confirmed by PCC analysis). Additionally, the system enabled Dox release upon
HA degradation by specific enzymes, proving the capping effect of the
polymeric shell and the enzyme-responsive intracellular drug release. Dox-
loaded HAPEI-MSNPs exhibit great killing efficiency at low drug concentrations
(nM range), which was comparable with that of pure Dox, but with specificity
towards CD44-overexpressing cancer cells. These results provide evidence that
the DDS here developed, supplied with targeting capability towards cancer
cells, endosomal escape capacity, controlled drug release and, consequently,
high therapeutic effect, holds great potential for tumor therapy applications. The
polymeric functionalization proposed can be applied to a wide range of
nanocarriers towards the increase of the therapeutic power at low drug dose
and the decrease of exocytosis rate, drastically reducing the side effects of
anti-cancer drugs.
4/27/23, 10:19 Page 26 of 51
Methods
Materials
Tetraethyl orthosilicate (TEOS, 98%), cetyltrimethylammonium chloride solution
(CTAC, 25% in H O), triethanolamine (TEA, 99%), hydrochloric acid (HCl, 1 N),
rhodamine B basic violet 10 (RhodB, 93%), fluorescein 5(6)-isothiocyanate
(FITC, ≥90% HPLC), polyethyleneimine solution (PEI, 50% w/v in H O), N-(3-
dimethylaminopropyl)-N′-ethyl-carbodiimide (EDC, 97%), N-
hydroxysulfosuccinimide sodium salt (sulfo-NHS, ≥98% HPLC), doxorubicin
hydrochloride (Dox, suitable for fluorescence, 98–102%, HPLC), sodium acetate
buffer solution, MES hydrate (titration, ≥99.5%), hyaluronidase type I-S (Hyal-1,
from bovine testes), hyaluronidase Type II (Hyal-2, from sheep testes) were
purchased from Sigma Aldrich. Sodium hyaluronate (HA, research grade, 289
kDa) was obtained from LifeCore BioMedical. Dulbecco’s modified eagle
medium (DMEM), and Lysotracker RED DND-99 were purchased from Molecular
Probes. Gentamicin, Dulbecco’s phosphate buffered saline (PBS, no calcium, no
magnesium), Hank’s balanced salt solution (HBSS, no phenol red), GlutaMaxi
supplement, fetal bovine serum (FBS, South America origin), Ethanol (absolute,
99.9%), Vybrant DiO cell-labeling solution were purchased from ThermoFisher
Scientific. Trypan blue solution (0.4%, TC grade) was purchased from Life
Science. All the chemicals were used without further purifications.
HAPEI-MSNPs preparation
The basic synthesis of MSNPs was performed by mixing 0.18 g of TEA with a
solution containing 24 mL of CTAC and 36 mL of milli-Q. The mixture was then
heated to 60 °C. After 1 h, 20 mL of TEOS (20 v/v % in 1-octadecene) was gently
2
2
4/27/23, 10:19 Page 27 of 51
added. The reaction was kept on going overnight at 60 °C under magnetic
stirring. After cooling to room temperature, the particles were washed with a
solution of HCl 1.1 M in water/ethanol (v/v = 1.25:10) using centrifugation-
dispersion-sonication cycles in order to remove CTAC from the pores.
Subsequently, two washing cycles were performed with milli-Q to bring the
solution to neutral pH. At this step, the concentration of particles in the colloidal
solution was estimated to be 10 mg/mL. The loading of RhodB and FITC were
performed in milli-Q, while for the Dox soaking, MSNPs were first dispersed in
phosphate buffer (pH 9) to increase the loading efficiency, which was otherwise
minimized at neutral pH. The three mixtures were left under magnetic stirring
(500 rpm) overnight. After centrifugation, the supernatants were then replaced
with milli-Q, obtaining MSNPs_RhodB/FITC/Dox as pellet (MSNPs_X). A solution
0.75% PEI in H O was adjusted at pH 7 and consequently added dropwise to
loaded-MSNPs in milli-Q water (1:1 v/v). PEI layer formation takes about 3 h
under magnetic stirring. In the meanwhile, HA were dissolved in 10 mL of MES
buffer (0.1 M, pH 6) (to be 0.4 mM of the final HA concentration) and stirred for
few hours. Afterwards, EDC and NHS-sulfo were simultaneously added to the
HA solution (with a final concentration of 1 mg/mL of EDC and NHS-sulfo) and
kept under stirring for 30 minutes for the activation of the carboxylic groups. On
the other hand, PEI functionalized particles (PEI-MSNPs_X) were washed via
centrifugation and dissolved in MES buffer as well. As a final step, 1 mL of
carboxyl-activated HA was slowly added to 3 mL of PEI-MSNPs_X (1 drop/5 s)
and kept under stirring overnight. The products (HAPEI-MSNPs_X) were then
washed and dispersed in milli-Q. Supernatants of each washing step were
collected and analyzed with UV-VIS spectrometer (Lambda 950, PerkinElmer) in
order to estimate the soaked cargo concentrations via absorbance. Final
concentrations of soaked RhodB and FITC inside HAPEI-MSNPs were
2
4/27/23, 10:19 Page 28 of 51
determined to be 276 and 165 µM, respectively. Dox concentration in HAPEI-
MSNPs used for the viability experiments was 40 µM, while for the release
experiments (in vitro and in cellulo) a higher concentration was used for
facilitating the absorbance and emission detection (150 µM).
HAPEI-MSNPs_X characterization
MSNPs obtained by biphase stratification method were first characterized for
size, shape and porosity by transmission electron microscopy (TEM). The
colloidal solution was deposited on an amorphous carbon-coated copper grid
and measured by JEOL-JEM 2100 TEM (200 kV). The loading was verified by
collecting wide-field images and the relative emission spectrum of
MSNPs_Dox/RhodB/FITC. These measurements were recorded by using an
inverted optical microscope (TiU, Nikon). Argon krypton ion laser (488 nm) was
used for FITC and Dox excitation, while Nd:YAG laser (532 nm) was applied on
RhodB samples. Lasers were focused on the samples by a 100x oil-objective
(N.A. 1.3, plan fluo Nikon) for RhodB and Dox samples, while a 40x objective
(N.A. 0.6, plan fluo Nikon) was used for the FITC sample. Emission images were
recorded by a charge-coupled device (CCD) camera (ImagEM, Hamamatsu)
operating at −85 °C. The spectra were collected by a CCD camera (DU920P,
Andor), equipped with a spectrograph (iHR320, Horiba), operating at −85 °C. A
pinhole (100 µm) was placed before the entrance of the spectrograph.
Longpass filters (HQ500LP for 488 nm, HQ545LP for 532 nm excitation,
Chroma) were employed either in front of the imaging CCD or in front of the
spectrograph to block the excitation light. The zeta potential of nanoparticles
was measured in milli-Q water by Delsa Nano HC (Beckman Coulter) and FE-
SEM images were collected by Quanta FEG250 FEI.
4/27/23, 10:19 Page 29 of 51
Cell culture
All cells were cultured in 25 cm cell culture flasks at 37 °C and under humidified
5% CO atmosphere. The cell passage was performed via trypsinization every
2–3 days, when the confluency reached 80%. NIH3T3 and A549 cell lines were
maintained in DMEM medium containing 10% FBS, 1% L-glutammax and 0.1%
gentamicin. For confocal imaging, cells were cultured in 35-mm glass bottom
dishes (MatTeK). When the confluency reached about 60%, the medium was
replaced with fresh medium (1 mL) and particles were added into the dishes,
which were then incubated at 37 °C under humidified 5% CO for different time
intervals.
Cellular uptake
For verifying the effect of different functionalization on the uptake, RhodB-
loaded particles (MSNPs_RhodB, PEI-MSNPs_RhodB and HAPEI-
MSNPs_RhodB) were incubated with NIH3T3 and A549 cells for 3 h at 37 °C.
Before the fluorescence measurements, the dishes were washed three times
with PBS to remove the residual extracellular particles; the cells were then kept
in HBSS. The plasma membrane was stained with DiO (1 µM) in HBSS for 15 min
and the dishes were visualized under a confocal fluorescence microscope
(FV1000, Olympus). High magnified images were obtained with 100x oil
objective (N.A. 1.40). The RhodB-encaspulated particles were visualized with
561 nm (20 µW, power density ~15.8 kW/cm ) excitation wavelength, while DiO
was imaged by using a 488 nm laser (5 µW, power density ~5.3 kW/cm ). A DM
405/488/559/635 was chosen as the main dichroic mirror; emissions were
detected through bandpass filters (BA 500–520 and BA570–670, respectively).
2
2
2
2
2
4/27/23, 10:19 Page 30 of 51
A SDM-560 was placed in front of the DiO detection channel as sharp-cut
dichroic mirror for splitting the emissions. The Z-stack method, changing the
focal length from bottom to top of a single cell, was used to collect a set of
images. The Z-reconstruction then offers an orthogonal view of the cell
thickness cross-section on the x/y-axis as a clear proof of the particles
internalization. Images were processed using FV10-ASW Viewer Software.
Fluorescence intensity analysis was performed by MATLAB software in order to
estimate the mean intensity per µm of cell. After background subtraction, a
single-cell area was manually selected and the cell volume was calculated over
a constant height of 1 µm (starting from the bottom of the cell).
Intracellular trafficking: fluorescence microscopy
In order to track the intracellular fate of the particles over time, FITC was
employed as cargo. MSNPs_FITC, PEI-MSNPs_FITC, HAPEI-MSNPs_FITC were
incubated individually with A549 cells for 3 h. Before the measurements, the
dishes were washed with PBS three times to remove the residual extracellular
particles; the cells were kept in HBSS during the measurements. Confocal
fluorescence measurements were performed immediately after the washing,
obtaining data of 3 h of incubation. On the other hand, after washing with PBS,
other copies of the same samples were suspended in fresh medium and placed
back in the incubator for measurements after 24 and 48 h. For the acidic
compartment imaging, endo/lysosomes were stained with Lysotracker Red (50
nM) in HBSS for 15 mins. Confocal images were obtained with FV 1000 Olympus
microscope, using 100x oil objective (N.A. 1.40). 488 nm (5 µW, ~5.3 kW/cm )
and 561 nm (2 µW, power density ~1.6 kW/cm ) excitation wavelengths were
applied for detecting FITC-loaded particles and Lysotracker-stained endo-
3
2
2
4/27/23, 10:19 Page 31 of 51
/lysosomes emissions, respectively. A DM 405/488/559/635 was chosen as
main dichroic mirror; emissions were detected through bandpass filters (BA
510–530 and BA575–675, respectively). Short-cut dichroic mirrors were located
in the emission pathways to split the emissions. A SDM 510 was placed before
the detection channels to reject scattered and excitation light. A SDM 530 was
applied to selectively reflect the FITC emission towards the 488-channel.
Intracellular location of the particles was verified collecting high magnification
2D-confocal fluorescence images. Images were processed using FV10-ASW
Viewer Software. Pearson correlation coefficient (PCC) analysis was carried
out using MATLAB software. After background subtraction and single cell area
selection, PCC values were estimated by the following equation (Eq. 1)
where R and G are the intensities per pixel i of the red and green channel,
respectively, and \(\bar{R}\) and \(\bar{G}\) are the corresponding mean
intensities over the cell area.
PCC values near 0 indicate that the fluorescence intensities of the two channels
are uncorrelated, while PCC values are close to 1 when the two fluorescence
intensities are perfectly linearly related. Threshold PCC values of the current
study, related images and descriptions are reported in SI (Supplementary
Fig. S3).
56
$$PCC=\frac{{\sum }_{i}({R}_{i}-\bar{R})\cdot ({G}_{i}-\bar{G})\,}{\sqrt{{\sum
}_{i}{({R}_{i}-\bar{R})}^{2}\cdot {\sum }_{i}{({G}_{i}-\bar{G})}^{2}\,}},$$
(1)
i i
4/27/23, 10:19 Page 32 of 51
Drug release in vitro
In order to perform the in vitro follow-up of the Dox release, three different pH
buffers were selected: acetate buffer (pH 4.5), MES buffer (pH 6) and PBS (pH
7.4). Dox-incorporated particles were suspended in the different buffers
individually and kept in a thermomixer (thermomixer comfort, eppendorf) under
stirring (300 rpm) at 37 °C. In order to check the enzymatic digestion of HA
layer, Hyal-1 and Hyal-2 (150 U/mL) were added to HAPEI-MSNPs_Dox in
acetate and MES buffer, respectively. Aliquots of the suspensions were taken at
different incubation time (0–72 h), centrifuged and analyzed in the micro-
volume spectrophotometer (BioDrop µLITE, BioDrop). The concentration of the
Dox released was estimated by collecting the absorbance at 490 nm.
Drug release in cellulo
10 µL of HAPEI-MSNPs_Dox (Dox: 150 µM) were added to 1 mL of medium
containing A549 cells on a glass-bottom dish. The dish was then placed in the
incubator for 3 h. Subsequently, the un-internalized particles were washed away
with PBS washing (x3) and the cells were measured by using a confocal
microscope (FV 1000 Olympus microscope) without any staining, obtaining data
of 3 h of incubation. On the other hand, after the PBS washing, other copies of
the same samples were suspended in fresh medium and placed back in the
incubator for measurements after 24 and 48 h. Differential interference contrast
(DIC) images were collected to visualize the cells and Dox emission was
detected by using a 488 nm laser (5 µW, power density ~5.3 kW/cm ), a 100x oil
objective (N.A. 1.4) and a bandpass filter 600–670 nm.
2
4/27/23, 10:19 Page 33 of 51
Anticancer efficiency
In order to carry out viability tests, the cells were seeded in TC Dish 35 Standard
dish (SARSTEDT) with 1 mL of medium and growth until density reached about 5
× 10 cells/cm . After removing the dead cells and placing fresh medium,
aliquots of 2, 4, 6, 8 and 10 µL solutions containing HAPEI-MSNPs_Dox, free
Dox and empty HAPEI-MSNPs were added individually to the culture medium.
The corresponding concentrations of Dox and particles in the medium were 80,
160, 240, 320 and 400 nM and 20, 40, 60, 80, 100 µg/mL, respectively. The
particle solutions were incubated with the cells overnight, then extracellular
particles were removed by PBS washing and a fresh medium was replaced. A
further incubation was executed, reaching 72 h in total. The viability was
estimated comparing the number of viable cells in the dishes after the treatment
(HAPEI-MSNPs_Dox, free Dox and empty particles incubation) with number of
viable cells in a control dish (no treatment). The cell counting was performed by
trypsinazing the cells and depositing aliquots of the cell suspension in glasstic
slide 10 with grids (KOVA). The estimation of viable cell number was
accomplished by following KOVA system protocol; non-viable cells were stained
with Trypan blue solution to be excluded from the counting. The viability data
were expressed as mean percentage of viable cell compared to control.
Statistical analysis
Data were shown as mean ± standard deviation. Each experiment was repeated
at least 3 times (n ≥ 3, specific n values indicated in figure captions). One-way
ANOVA test was performed to compare differences among groups, followed by
post-hoc t-test analysis. Results were considered statistically significant at p <
5 2
4/27/23, 10:19 Page 34 of 51
0.05.
Data Availability
The datasets generated during the current study are available from the
corresponding authors on reasonable request.
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Acknowledgements
This work was supported by KAKENHI (JP17H03003, JP17H05244, and
JP17H05458), FWO (G0B5514N, G081916N, G056314N, and G025912N) and
ERC (#280064 to H. U.). Financial support from the KU Leuven (C14/15/053,
OT/12/059, and IDO/12/008), JST PRESTO, and BELSPO (IAP VII/05) is greatly
acknowledged. A. M. and H. U. are indebted to the Cooperative Research
Program of “NJRC Mater. & Dev”. M. R. and B. F. acknowledge the support from
the Research Foundation-Flanders for FWO PhD and Postdoctoral
fellowships, respectively (1S33117N and 12X1419N). A. M. acknowledges the
Yashima Environment Technology Foundation and the MIKIYA Science and
Technology Foundation for the support. L. L. acknowledges the support from
the University of Perugia (Fondo d’Ateneo per la Ricerca di base 2014). A part of
this work was supported by the “Nanotechnology Platform” in Hokkaido
University. B. F. and T. I. acknowledge the RIES International Exchange Program
of “Dynamic Alliance for Open Innovation Bridging Human, Environment and
Materials” from MEXT. The authors would like to thank Prof. Peter Peters
(Maastricht University) and his research group for helping the TEM
4/27/23, 10:19 Page 46 of 51
measurements and the related sample preparation.
Author information
Authors and Affiliations
KU Leuven, department of Chemistry, Celestijnenlaan 200G-F, Heverlee, 3001, Belgium
Beatrice Fortuni, Monica Ricci, Indra Van Zundert, Eduard Fron, Hideaki Mizuno, Susana Rocha & Hiroshi Uji-i
RIES Hokkaido University, Research Institute for Electronic Science, N20W10, Kita-Ward Sapporo, 0010020, Japan
Tomoko Inose & Hiroshi Uji-i
Toray Research Center, Inc., 3-3-7, Sonoyama, Otsu, Shiga, 520-8567, Japan
Yasuhiko Fujita
Yamagata University, department of Engineering, Yonezawa, Yamagata, 992-8510, Japan
Akito Masuhara
University of Perugia, department of Chemistry, Biology and Biotechnology, via Elce di sotto 8, Perugia, Italy
Loredana Latterini
Contributions
B.F. and H.U. conceived and designed all experiments, B.F., T.I., M.R., I.Z., A.M.
and E.F. conducted the experiments; B.F., S.R. and Y.F. analyzed the data; H.M.
supervised the biological aspects of the project; H.U., S.R., H.M., L.L. and E.F.
critically reviewed and revised the scientific content of the manuscript,
providing crucial suggestions; B.F. wrote down the manuscript, H.U. supervised
the project.
Corresponding authors
Correspondence to Beatrice Fortuni, Susana Rocha or Hiroshi Uji-i.
4/27/23, 10:19 Page 47 of 51
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Fortuni, B., Inose, T., Ricci, M. et al. Polymeric Engineering of Nanoparticles for Highly
Efficient Multifunctional Drug Delivery Systems. Sci Rep 9, 2666 (2019).
https://doi.org/10.1038/s41598-019-39107-3
Received
16 July 2018
Accepted
16 January 2019
Published
25 February 2019
DOI
https://doi.org/10.1038/s41598-019-39107-3
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Subjects Chemical engineering • Drug delivery • Nanoparticles • Synthesis and processing
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