SWOT ANALYSIS APPLIED
TO THE LIFE SCIENCES
BIOLOGY 495
SHARON BROWN, PH.D.
WHAT IS A SWOT ANALYSIS?
• SWOT
stands for Strengths, Weaknesses, Opportunities,
Threats. A SWOT analysis is a simple but useful framework
for analyzing your organization's strengths and weaknesses,
and the opportunities and threats that you face.
• A SWOT analysis is one of the most commonly used strategic
planning techniques.
• The primary use of a SWOT analysis is to provide structure to,
or summarize, your innovative analysis. This technique can
also be used in decision making, to help determine which of
several options is better.
WHY USE A SWOT ANALYSIS
IN THE LIFE SCIENCES?
• A SWOT analysis will help summarize your understanding of the major issues
identified in your innovative analysis
• Your innovative analysis will focus on identifying the strengths, weaknesses,
opportunities and threats in three key strategic environments.
Three strategic environments:
• Your internal environment
• Your industry environment
• Your macroenvironment
STRENGTHS
Strength refers to a core competency of your business
where your business has a competitive advantage
TANGIBLE INTERNAL STRENGTHS:
These tend to be strengths that can be
precisely identified, measured or realized
•Physical assets, including plant or
facilities and equipment
•Assets you have in your team, such as
knowledge, education, network, skills,
and reputation
•Long-term rental or business contracts
•Unique or market-leading products
•Financial resources to fund change or
change management of your business
•Competitive cost advantages
•Advanced technologies or information
systems
•High volume business production
•Scalability of business or products
INTANGIBLE NTERNAL STRENGTHS: These tend
to be strengths that cannot be physically touched
or physically measured
•Brand or product recognition
•Location
•Business reputation—Are you a market leader in
your industry?
•Goodwill and customer relationship management
•Business/Supplier relationships
•Strong employee relationships
•Strategic business alliances or partnerships
•Intellectual property rights or patents
•Advertising strategy or process
•Level of experience in your field
•Level of management experience in your
workforce
•Industry knowledge superiority
•Industry associations that give competitive
advantage
•Innovative practices within your business
WEAKNESSES
A weakness refers to a core competency of your business where your
competitor has a competitive advantage when it comes
to customer value propositions
TANGIBLE INTERNAL WEAKNESSES: These
tend to be weaknesses that can be
precisely identified, measured or realized
• Old or outdated plant and
equipment
• Lack of qualified personnel
• Narrow product line
• Insufficient financial resources to
fund necessary changes
• High operating costs
• Inferior technology
• Low volume and restricted
scalability
TANGIBLE EXTERNAL WEAKNESSES: These
tend to be weaknesses that cannot be
physically touched or physically measured
•
•
•
•
•
•
•
•
•
•
Weak or unrecognizable brand
Weak or unrecognizable image
Poor relationships with your customers
Poor relationships with your suppliers
Poor relationships with your employees
Marketing failing to meet objectives
Manager inexperience
Ineffective research & development
Insufficient industry knowledge
Meager innovative skills
OPPORTUNITIES
An opportunity is an environmental condition in your macro or industry environments
that can improve your organization's competitive position relative to that of your
competitors. When completing your analysis, you will find that your opportunities
generally fall under two categories.
INDUSTRY OPPORTUNITIES: These
are opportunities in your industry
environment and generally reduce
the level of price competition in your
industry.
•Expansion of your product range
•Diversification of your business
interests
•Growth in your customer's field
•Growth in your supplier's field
•Expansion of your customer base
•Placid competition
•Export opportunities
•Products or service growth
MACRO
OPPORTUNITIES:
These
opportunities are in the broader
environment that generally impacts
all businesses in your region.
•Favorable changes to legislation
•Favorable changes to any import/export
constraints
•Favorable economic outlook
•Favorable cultural shifts
•Technology that your business can embrace
and utilize, such as Ecommerce or Internet sales
THREATS
A threat is a forecasted environmental condition that is out of your control
and has the potential to harm your business‘s profitability.
INDUSTRY THREATS: These threats
are related to an increase in the
competition in your industry or a
reduction in market size reduce your
business‘s profitability.
•
•
•
•
•
Low cost imports: The threat of low-cost
imports affects almost any manufacturer
in the developed world
Consumer ability to shift to a substitute
product and changing demand for
substitute products
Slow market growth or decline in market
size
Shifts in customer or supplier buying
power:
The changing needs of buyers (customers)
MACRO THREATS: These threats affect all
industries in your region and result in
risk of reduced profitability.
•
•
•
Shifts in foreign exchange rates that impact
imports or exports:
Demographic changes: The aging workforce
makes it difficult to hire skilled workers in many
developed countries.
Industry Regulations: increasing regulations
and increased costs needed to administer these
new regulations.
SWOT SUMMARY
CONCLUSION: WHAT’S NEXT?
•
•
•
•
With your SWOT analysis complete, you’re ready to convert it
into real strategy. After all, the exercise is about producing a
strategy that you can work on during the next few months to
begin work your innovation.
You will need to look at your strengths and figure out how
you can use those strengths to take advantage of your
opportunities. Then, look at how your strengths can combat
the threats that are in the biotechnology market. Use this
analysis to produce a list of actions that you can take.
You will also want to analyze how external opportunities
might help you combat your own, internal weaknesses. Can
you also minimize those weaknesses so you can avoid the
threats that you identified?
The next step is to conduct a futuring analysis to see how
your proposed innovation fits into the future of
biotechnology.
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OPEN
Received: 16 July 2018
Accepted: 16 January 2019
Published: xx xx xxxx
Polymeric Engineering of
Nanoparticles for Highly Efficient
Multifunctional Drug Delivery
Systems
Beatrice Fortuni1, Tomoko Inose2, Monica Ricci1, Yasuhiko Fujita3, Indra Van Zundert1,
Akito Masuhara4, Eduard Fron1, Hideaki Mizuno1, Loredana Latterini5, Susana Rocha1 &
Hiroshi Uji-i1,2
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 at low drug dose. To this end, we developed a multifunctionalized DDS
that combines high specificity towards cancer cells with endosomal escape capabilities. Doxorubicinloaded 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.
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-imaging1–5. 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 functionalization6–8. Unlike some other nanocarriers, MSNPs have not
been translocated into the clinical stage yet9. However, the reasonable biocompatibility accomplished in vivo is
extremely promising for a proximate Food and Drug Administration (FDA-) approval10.
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 endocytosis11–14. 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),
1
KU Leuven, department of Chemistry, Celestijnenlaan 200G-F, Heverlee, 3001, Belgium. 2RIES Hokkaido University,
Research Institute for Electronic Science, N20W10, Kita-Ward Sapporo, 0010020, Japan. 3Toray Research Center,
Inc., 3-3-7, Sonoyama, Otsu, Shiga, 520-8567, Japan. 4Yamagata University, department of Engineering, Yonezawa,
Yamagata, 992-8510, Japan. 5University of Perugia, department of Chemistry, Biology and Biotechnology, via Elce
di sotto 8, Perugia, Italy. Correspondence and requests for materials should be addressed to B.F. (email: beatrice.
fortuni@kuleuven.be) or S.R. (email: susana.rocha@kuleuven.be) or H.U. (email: hiroshi.ujii@kuleuven.be)
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in metastasis, as well as in cancer stem cells15. 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 use16–18. 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
endocytosis19–26.
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
exocytosis27. Previous reports have shown that non-coated MSNPs co-localize with the endo-/lysosomes in the
early stage of incubation28–31, and are quickly exocytosed, following this pathway32. Similarly, HA-coated MSNPs
are internalized via CD44-mediated endocytosis and are subjected to same endocytic system, ending up in the
acidic cellular compartments within few hours of incubation23,33, and being ejected via exocytosis within 48 h34.
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
activity35. The therapeutic efficiency can be further decreased by the fast exocytosis of the nanocarriers36. 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 peptides37,38. 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 nucleus39,40. The use of DNA-PEI polyplexes, instead of pure DNA, was demonstrated to improve the gene
expression efficiency up to 100-fold40,41. This enhanced gene expression can be associated to the so-called “proton
sponge effect” of PEI42. 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 hypothesis in literature, however, it is important to mention that this concept
is still heavily debated43.
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 enzymes44, 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)45,46. 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, respectively45,47. 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
performed48. 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.
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 proposed10,49. MSNPs were synthetized
using the biphase stratification method developed by Shen et al., that yields particles with a pore size of ~2.8 nm50.
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).
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 (pKa 8–10, depending on the molecular weight of the polymer)51,52, whereas ~50%
of hydroxyl groups on a silica surface are deprotonated (pKa ≈ 6.8)53. 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 change in the zeta potential of the particles after the coating (from −38.2 mV to
+37.7 mV, Fig. 2c).
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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.
Figure 2. 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, PEIMSNPs and HAPEI-MSNPs. (d–f) FE-SEM images of MSNPs, PEI-MSNPs and HAPEI-MSNPs, respectively.
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Figure 3. 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 bottom panels show the yz and xz
projections, respectively. (g) Mean intensity of the RhodB signal per μm3 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).
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 reaction23. 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 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)54 and NIH3T3 (mouse embryonic
fibroblasts, lowly expressing CD44 receptors, defined as CD44-negative or CD44-inactive cells)55.
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
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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 results45 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 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.
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 exocytosed23,28,30,31,33. 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)56 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.
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 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 organelles42. It is important to mention that the possible
proton sponge effect of PEI does not change the pH of the endo-/lysosomes57, 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).
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, 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 membrane43.
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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.
®
Drug release in vitro. Thanks to the therapeutic effectiveness towards a wide range of cancers (carcinomas,
sarcomas and hematological cancers)58 and to its fluorescent properties59, 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 well60. 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 pKa
values of PEI and silica hydroxyl groups53,61, 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 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 pH47.
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Figure 5. Drug release in vitro and in cellulo. (a) Time-dependent in vitro release profile of Dox from HAPEIMSNPs_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 images.
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.
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 membrane62,63. 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 endocytosis63.
Scientific Reports |
(2019) 9:2666 | https://doi.org/10.1038/s41598-019-39107-3
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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).
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 reports43. 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 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 fluorescence64–66, up to 95% of its intrinsic emission67.
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 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).
While at low concentration (
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