Hollow Mesoporous Silica Nanocarriers with Multifunctional Capping Agents for in Vivo Cancer Imaging and Therapy

Article abstract of DOI:10.1002/smll.201503121

Efficient drug loading and selectivity in drug delivery are two key features of a good drug-carrier design. Here we report on such a drug carrier formed by using hollow mesoporous silica nanoparticles (HMS NPs) as the core and specifically designed multifunctional amphiphilic agents as the encapsulating shell. These nanocarriers combine the advantages of the HMS NP core (favorable physical and structural properties) and the versatility of an organic-based shell (e.g., specificity in chemical properties and modifiability). Moreover, both the properties of the core and the shell can be independently varied. The varied core and shell could then be integrated into a single device (drug carrier) to provide efficient and specific drug delivery. In vitro and in vivo data suggests that these drug nanocarriers are biocompatible and are able to deliver hydrophobic drugs selectively to target tumor cells. After the break of the pH-labile linkages in the shell, the drug payload can be released and the tumor cells are killed.

Full text of DOI:10.1002/smll.201503121

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Biocompatible Materials
Hollow Mesoporous Silica Nanocarriers with
Multifunctional Capping Agents for In Vivo Cancer
Imaging and Therapy
Shun Yang, Dongyun Chen,* Najun Li, Qingfeng Xu, Hua Li, Frank Gu, Jianping Xie,*
and Jianmei Lu*
E fficient drug loading and selectivity in drug delivery are two key features of a good
drug-carrier design. Here we report on such a drug carrier formed by using hollow
mesoporous silica nanoparticles (HMS NPs) as the core and specifically designed
multifunctional amphiphilic agents as the encapsulating shell. These nanocarriers
combine the advantages of the HMS NP core (favorable physical and structural
properties) and the versatility of an organic-based shell (e.g., specificity in chemical
properties and modifiability). Moreover, both the properties of the core and the shell
can be independently varied. The varied core and shell could then be integrated into
a single device (drug carrier) to provide efficient and specific drug delivery. In vitro
and in vivo data suggests that these drug nanocarriers are biocompatible and are able
to deliver hydrophobic drugs selectively to target tumor cells. After the break of the
pH-labile linkages in the shell, the drug payload can be released and the tumor cells
are killed.
Dr. S. Yang, Prof. D. Chen, Prof. N. Li, Prof. Q. Xu,
Prof. H. Li, Prof. J. Lu
1. Introduction
College of Chemistry
Chemical Engineering and Materials Science
Collaborative Innovation Center of Suzhou Nano
drug carriers with diagnostic and therapeutic modalities (or
Science and Technology
Soochow University
Current interest in the development of drug-delivery systems
has shifted toward multifunctional drug carriers, as hybrid
theranostic carriers) have shown significant improvement in
diagnostic accuracy and antitumor efficacy.[1–4] Drug carriers
Suzhou 215123, P. R. China
E-mail: dychen@suda.edu.cn; lujm@suda.edu.cn
of nanometer size (or nanocarriers) are ideal for this pur-
pose as they provide a particularly efficient platform to inte-
grate multiple functionalities, including targeting, imaging,
and therapy (or more), into a single entity. Efficient drug
loading and selective drug delivery are two key features for
a good design of drug nanocarriers. Here we report a unique
core–shell nanocarrier structure, where the complemen-
tary features of the two key components – inorganic nano-
particles (NPs, as the core) and functional organic moieties
(as the capping shell) can be independently designed and
integrated into a single nanocarrier to achieve efficient and
specific cancer therapy. By using inorganic nanoparticles,
Prof. J. Xie
Department of Chemical & Biomolecular Engineering
Faculty of Engineering
National University of Singapore
117576, Singapore
E-mail: chexiej@nus.edu.sg
Prof. F. Gu
Department of Chemical Engineering
Waterloo Institute for Nanotechnology
University of Waterloo
Ontario N2L 3G1, Canada
DOI: 10.1002/smll.201503121
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the nanocarriers could achieve a high drug loading content.
The incorporation of stimuli-responsive moieties in the
Moreover, the functional organic moieties could be designed capping layers is most attractive in this perspective.[20–25]
to be biodegradable/biocompatible, to specifically target dis- Biodegradable capping layers, which degrade in response to
ease sites, and to be sensitive to the environmental condi- a particular external stimulus, such as temperature, pH, and
tions. These core–shell nanocarriers could thus be employed enzymatic or photochemical reactions, have recently been
for efficient drug uptake, targeted drug delivery, and effective used in nanocarrier design to achieve a controlled release
cancer therapy, with improved performance in cancer man- of drug payloads within the affected cells.[26,27] For example,
agement compared with conventional therapeutics.[5,6]
Hollow mesoporous silica (hereafter referred to as HMS) drug payloads in a slightly acidic environment (pH 5.0–6.5)
NPs were chosen in this study as the drug carrier. Inor- of endosomes or lysosomes inside affected cells.[28–30]
pH-responsive moieties were used for a controlled release of
A
ganic NPs, such as gold, magnetic, and silica nanoparticles, variety of amphiphilic polymers with pH-labile linkages have
have been widely used as drug carriers owing to their facile been recently developed to achieve the controlled release of
synthesis, good colloidal stability, and excellent structural drug payloads within cells. The selective delivery of drug car-
rigidity.[7–11] In particular, mesoporous silica NPs, which also riers to cancer cells can be further realized by incorporating
feature a high porosity (or large surface area), tunable pore a cancer-targeting ligand, such as folic acid, arginylglycylas-
morphology (size and structure), and facile surface modifica- partic acid (RGD) peptides, sugars or antibodies, to the
tion, have recently emerged as an ideal nanocarrier for drug amphiphilic polymers.[31–37]
delivery.[12] By using a hollow structure for the mesoporous
Herein, we report a novel core–shell structured nanocar-
silica NPs (termed hollow mesoporous silica or HMS NPs), rier consisting of a hydrophobic fluorescent HMS NP as the
the drug-loading capacity and drug-releasing profile could be core and well-designed, multifunctional amphiphilic agents as
further improved, where the free volume in the hollow inner the capping shell for selective and controlled drug delivery.
of the HMS NPs can improve the drug loading capacity and Two key elements that can be independently designed and
the mesoporous layer of the NPs can provide a sufficient tailored have thus been incorporated in our nanocarrier
and controlled diffusion path for the drug payloads. Fur- design. As shown in Scheme 1, the first element is a hydro-
thermore, the size and shape of the hollow inner, and the phobic fluorescent HMS NP, which provides an excel-
thickness (or diffusion length) and pore morphologies of lent drug-loading capacity and drug-releasing profile. The
the mesoporous layer, can be independently fine-tuned to second element is an amphiphilic capping agent consisting
achieve an optimal drug loading and releasing profile in a of four functional motifs: a targeting segment (folic acid,
particular bio-setting. One key challenge in the use of HMS FA), a hydrophilic segment (4,7,10-trioxa-1,13-tridecanedi-
NPs as drug carriers is the potential toxicity of unmodi- amine, TDA), a pH-labile linkage, and a hydrophobic seg-
fied silica NPs. This challenge has been recently addressed ment (4-n-dodecyloxybenzalacetal, DBA), as illustrated in
by incorporating a polymer coating on the silica NP sur- Scheme 1. The as-designed multifunctional amphiphilic cap-
face.[13–17] However, using a thick polymer-coating layer on ping agents are hereafter referred to as FA-TDA-DBA or
the HMS NP surface may constrain the drug loading and FTD for short. HMS is easily encapsulated by FTD to form
releasing from the HMS nanocarriers. Therefore, there is a HMS@FTD because of the strong interaction between the
strong interest in the design of “smart” and efficient capping hydrophobic HMS NP and the hydrophobic segment of the
layers for the HMS nanocarriers to achieve a selective and FTD, and doxorubicin (DOX) can also be loaded to form
controlled drug release.[18,19]
DOX-loaded HMS@FTD in this process. The FTD capping
Scheme 1. Schematic illustration of the two key components in the design of the HMS@FTD drug nanocarriers: hydrophobic fluorescent HMS NPs
and multifunctional amphiphilic FTD capping ligands.
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agents can provide good protection for the HMS nanocar- Section (Figure S1 in the Supporting Information). The as-
riers in bio-fluids. They also make the specific targeting of designed amphiphilic capping agents provide a good protec-
cancer cells (because of the FA moieties) possible, as well as tion for the loaded drugs inside the HMS nanocarriers, which
the subsequent controlled release of drug payloads (from the could largely inhibit the premature release of the drug. The
pH-responsive moieties) in the affected cells.
core–shell drug nanocarriers were formed via self-assembly
of the hydrophobic fluorescent HMS NPs, hydrophobic drugs
(DOX), and amphiphilic FTD capping agents, based upon the
strong hydrophobic interactions between DOX and the alkyl
chains on the HMS NP surface (C18) and in the FTD capping
2. Results and Discussion
The fabrication of monodisperse HMS NPs (ca. 150 nm dia- agents (C12).[38–40] The hydrophobic drugs were encapsulated
meter) was carried out according to our previous procedure, inside the HMS NPs, and their release could be blocked by
where polystyrene NPs (125 nm), tetraethyl orthosilicate the FTD capping agents on the NP surface. It should be men-
(TEOS), and cetyltrimethylammonium bromide (CTAB) tioned that the as-fabricated drug nanocarriers with a high
were used as the hollow-generating NPs, silica precursors, and content of FA ligands showed a poor solubility in aqueous
pore-generating agents, respectively.[12] The pristine HMS solution because of the hydrogen bonding and π–π stacking
NPs were further incorporated with a fluorescent dye (e.g., between the FA ligands. To address this issue, an FA-free
red-emitting rhodamine B, RhB) to form fluorescent HMS amphiphilic ligand, TDA-DBA (or TD), was used as a co-cap-
NPs. A long-chain hydrophobic ligand octadecyltrimethoxysi- ping agent to the FTD ligands to improve the water solubility
lane (OTMS) was then crafted on the surface of the fluores- of the as-fabricated drug nanocarriers. The ideal mole ratio of
cent HMS NPs, which led to a remarkable wettability change FTD to TD ligands was determined to be 5:95. This optimized
of the NP surface—from highly hydrophilic to hydrophobic. ratio was used in our further study. The as-fabricated drug
Such surface modification could improve the hydrophobic nanocarriers are hereafter referred to as DOX-loaded HMS@
interactions between the HMS NPs and the amphiphilic FTD FTD nanocarriers. The as-fabricated drug carriers showed
capping agents (via their hydrophobic segments). The hydro- excellent solubility and stability in aqueous solutions (pH 7)
phobic nature of the NP surface can also increase the adsorp- and in various buffer solutions (e.g., phosphate buffer).
tion of the hydrophobic drugs, which could lead to a better
control of the drug release.
Transmission electron microscopy (TEM) was used to
examine the morphology and structure of the pristine HMS
The detailed synthesis and characterization of the amphi- NPs, hydrophobic fluorescent HMS NPs, and DOX-loaded
philic FTD capping agents are presented in the Experimental HMS@FTD nanocarriers. As shown in Figure 1a, nearly
Figure 1. a–c) Representative TEM images of pristine HMS NPs (a), hydrophobic fluorescent HMS NPs (b), and DOX-loaded HMS@FTD nanocarriers
(c); d) Dynamic light scattering analysis of the pristine HMS NPs and DOX-loaded HMS@FTD nanocarriers.
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monodisperse spherical particles with a diameter of around
Confocal laser scanning microscopy (CLSM) was further
used to confirm the successful encapsulation of DOX inside
150 nm were observed for the pristine HMS NPs.[41,42]
A
strong contrast between the edge (darker, ca. 20 nm in thick- the HMS@FTD nanocarriers. The as-prepared DOX-loaded
ness) and the center (lighter, ca. 110 nm in diameter) could HMS@FTD was used without any further treatment after
also clearly be observed in the NPs, implying the hollow centrifugation in this study. DOX shows red fluorescence
structure of the pristine HMS NPs. The incorporation with with a respective excitation and emission wavelength of 485
the fluorescent dyes and the modification with the hydro- and 592 nm. To differentiate the fluorescence of the loaded
phobic ligands had negligible effects on the morphology and DOX (red emission) from that of the fluorescent HMS nano-
structure of the HMS NPs, as evidenced by TEM (Figure 1b). carriers, the nanocarriers were labeled with a green-emitting
However, the particle size of the HMS@FTD nanocarriers organic dye, fluorescein isothiocyanate (FITC). Both green
increased from around 150 nm (the pristine HMS NPs) to (Figure 2a, λem = 518 nm) and red (Figure 2b, λem = 592 nm)
around 155 nm, as shown in Figure 1c. The increase in par- fluorescence was observed in the DOX-loaded HMS@FTD
ticle size was also supported by dynamic light scattering nanocarriers under an excitation wavelength of 485 nm. This
(DLS, Figure 1d) analyses of the pristine HMS NPs and data provides more supportive evidence for the successful
HMS@FTD nanocarriers. In addition, compared to the pris- loading of DOX in the HMS@FTD nanocarriers.
tine HMS NPs (Figure 1a), where the pore structure can be
The loading efficiency and capacity of DOX in the
clearly identified, the pore structure in the HMS@FTD nano- HMS@FTD nanocarriers were determined by UV-vis spec-
carriers (Figure 1c) was very difficult to detect. This change troscopy to be 89.4% and 158.9 µg DOX per mg of nano-
in pore structure suggests the successful capping of the FTD carriers, respectively. The drug-loading efficiency was much
ligands on the HMS NPs. The blocking of the pores by the higher when compared to traditional drug-delivery systems
FTD capping ligands could therefore provide a better pro- (e.g., mesoporous silica nanoparticles and micelles).[44] The
tection for the loaded drugs inside the HMS nanocarriers. loaded DOX in the HMS@FTD nanocarriers was very stable
The successful capping of the FTD ligands on the HMS NPs in PBS buffer at physiological pH (7.4). As shown in Figure 3,
was also substantiated by thermogravimetric analysis (TGA) the release of the loaded DOX was negligible at pH 7.4 even
of the HMS@FTD nanocarriers, where the capping ligands after a long (100 h) incubation time. This result also confirms
contributed about 13% of the weight of the nanocarriers. In that DOX was loaded into the nanoparticles instead of on the
contrast, the organic moieties contributed to only 4% of the surface of the nanoparticles. Moreover, at pH 5.0 (Figure 3),
weight of the nanocarriers in the pristine HMS NPs.
a fast release of DOX was triggered in the HMS@FTD nano-
Fourier-transform infrared spectroscopy (FTIR) pro- carriers over the first 10 h, implying a fast hydrolysis of the
vided yet another line of evidence for the successful cap- amphiphilic capping ligands (FTD) at the slightly acidic pH.
ping of the FTD ligands and loading of the hydrophobic The TEM images of the degraded HMS@FTD nanoparticles
drugs (DOX) in the HMS@FTD nanocarriers. As shown at pH 5.0 are shown in Figure S3 (Supporting Information).
in Figure S2 (Supporting Information), a distinct peak at It can be seen that the FTD capping ligands have detached
around 1750 cm−1 was observed in the FTIR spectrum of from the HMS, and the pores have become uncovered, which
DOX-loaded HMS@FTD nanocarriers (bottom line). This further confirms that the release of DOX was caused by the
peak could be assigned to the vibration stretching of the pH-induced removal of folic acid from the particles. The
amide group in folic acid, which implied the successful cap- strongly pH-dependent release profile of the HMS@FTD
ping of the FTD ligands on the HMS NPs. In addition, three nanocarriers provides an ideal platform for selective and con-
absorption peaks at 1405, 1612, and 1070 cm−1, which corre- trolled drug delivery, where the drug release can be largely
sponded well to the carbonyl groups in quinine and ketone inhibited during circulation in the blood at physiological pH,
of DOX,[43] were enhanced significantly in the FTIR spec- and the loaded drugs will only be unloaded when the nano-
trum of DOX-loaded HMS@FTD nanocarriers compared carriers reach the targeted cells and will then be transported
to that of OTMS modified fluorescent HMS (OFHMS). This into the cells. In addition, a more sustained drug release pro-
data confirmed the successful loading of DOX in the HMS@ file was observed in the HMS@FTD nanocarriers compared
FTD nanocarriers.
to that of pristine HMS nanocarriers, further suggesting the
Figure 2. CLSM images of DOX-loaded HMS@FTD nanocarriers under different emission wavelengths (λex = 485 nm): a) λem = 518 nm for
green-emitting FITC; b) λem = 592 nm for red-emitting DOX; and c) physically overlaid image of the panels (a) and (b). Scale bar: 5 µm.
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To further evaluate the targeting capability of the HMS@
FTD nanocarriers, KB cells with over-expressed folate recep-
tors were treated with DOX-loaded HMS@FTD nanocarriers,
DOX-loaded HMS@TD nanocarriers (without the FA moie-
ties in the capping agents), and free DOX in different concen-
trations (5, 10, and 20 µg DOX mL−1) for 24 h. As shown in
Figure 4b, the viability of the KB cells depended strongly on
the DOX dosage. The DOX-loaded HMS@FTD nanocarriers
(Figure 4b, middle columns) showed nearly the same cytotox-
icity against the KB cells as free DOX (right-hand columns)
at different levels of DOX dosage. This value was much
higher than that of the DOX-loaded HMS@TD nanocarriers
without the FA ligands (left-hand columns). The obvious dif-
ference in cytotoxicity between the HMS@FTD nanocarriers
and HMS@TD nanocarriers suggests the high selectivity of
the FA ligands toward the KB cells. The HMS@FTD nano-
carriers (with the FA ligands) were able to specifically target
the KB cells with over-expressed folate receptors, and they
can be uptaken by the cells via a direct folate-receptor-medi-
ated endocytosis pathway.[45–47] This active transport pathway
is more efficient than the non-specific and passive endocytosis
Figure 3. DOX release profiles of DOX-loaded pristine HMS NPs and
HMS@FTD nanocarriers at pH 5.0 and 7.4.
crucial role of the hydrophobic surface modification and the pathway, leading to an improved cytotoxicity to the KB cells.
as-designed FTD capping ligands on the NP surface. Even The targeting efficacy of the FA ligands was further
though the HMS@FTD did not fully release the loaded DOX confirmed by CLSM and flow cytometry studies. Human
within 100 h as opposed to the pristine HMS nanocarriers, hepatoma 7402 cells without over-expressed folate receptors
the therapeutic efficacy of the HMS@FTD nanocarriers were chosen as a negative cell line. The red fluorescence of
would be much higher than that of the pristine HMS nano- DOX was used to monitor the intracellular activity of the
carriers due to the pH-controlled release. The as-designed drug nanocarriers. The KB cells and 7402 cells were treated
HMS@FTD nanocarriers provide an excellent platform for with DOX-loaded HMS@FTD nanocarriers (FA-conjugated)
well-controlled drug release, which could further be used to and HMS@TD nanocarriers (FA-free) at the same DOX
achieve good long-term antitumor efficacy.
concentration of 10 µg mL−1. The red fluorescence of the
The HMS@FTD nanocarriers also showed good biocom- nanocarriers inside the cells was monitored by CLSM over a
patibility. KB cells were chosen as the model cancer cell line period of 3 h. As shown in Figure 5a and 5b (top panel), no
to evaluate the potential cytotoxicity of the drug carriers. obvious difference in the fluorescence intensity was detected
The KB cells were first treated with DOX-free HMS@FTD for the DOX-loaded HMS@FTD nanocarriers in the KB and
nanocarriers in different concentrations for 24 h, and the 7402 cells at 0.5 h. The possible reason is that, at the early
viability of the cells was measured using a sulforhodamine B stage of incubation (ca. 0.5 h), the KB and 7402 cells might
(SRB) assay. As shown in Figure 4a, a very low cytotoxicity uptake the drug nanocarriers via a similar non-specific and
(cell viability >90% after 24 h incubation) was observed for passive endocytosis pathway. Similarly, some red fluorescence
DOX-free HMS@FTD nanocarriers even at a high dosage intensity was also observed for the DOX-loaded HMS@TD
concentration (500 µg mL−1). This good biocompatibility of nanocarriers in the KB cells (Figure 5c, top panel). However,
the HMS@FTD nanocarriers can be ascribed to the non- after 3 h of incubation, the fluorescence intensity of the DOX-
toxic feature of the as-designed amphiphilic FTD capping loaded HMS@FTD nanocarriers (FA-conjugated) in the KB
agents.
cells (folate receptor positive, Figure 5a, bottom panel) was
Figure 4. The viability of KB cells after incubation with a) DOX-free HMS@FTD nanocarriers, and b) DOX-loaded HMS@TD (left), DOX-loaded HMS@
FTD nanocarriers (middle), and free DOX (right) for 24 h.
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Figure 5. a,b) Representative CLSM images of incubation of DOX-loaded HMS@FTD nanocarriers in a) KB cells and b) 7402 cells for 0.5 h (top) and
3 h (bottom). c,d) CLSM images of incubation of DOX-loaded HMS@TD nanocarriers in c) KB cells and d) 7402 for 0.5 h (top) and 3 h (bottom).
e) Mean fluorescence intensity (MFI) of DOX-loaded HMS@FTD nanocarriers in KB cells (blue) and 7402 cells (green); and DOX-loaded HMS@TD in
KB cells (red) and 7402 cells (cyan) for 0.5 and 3 h.
significantly higher than that in the 7402 cells (folate receptor the FA-conjugated nanocarriers (HMS@FTD) in the folate
negative, Figure 5b, bottom panel). Similarly, after 3 h of receptor positive cells (KB and HeLa).
incubation, the FA-conjugated nanocarriers (HMS@FTD,
The in vitro studies showed that the DOX-loaded HMS@
Figure 5a, bottom panel) in the KB cells also showed a much FTD nanocarriers were able to selectively deliver the drugs
stronger intensity compared to that of the FA-free nanocar- to the cancer cells with over-expressed folate receptors. To
riers (HMS@TD, Figure 5c, bottom panel) in the same cancer determine the maximum tolerance dose (MTD) of the DOX-
cell line. These results further confirm that the direct internal- loaded HMS@FTD nanocarriers, the nanocarriers were intra-
ization via folate-receptor-mediated endocytosis has greatly venously injected into healthy Balb/c mice at doses of 0 (used
enhanced the uptake of the drug nanocarriers. Similar obser- saline solution as the control), 20, 40, or 60 mg DOX per kg
vations were observed in the flow cytometry studies, where body weight. The mice showed morbidity if the highest dose
the mean fluorescence intensity (MFI) was used to evaluate of DOX (60 mg DOX per kg body weight) was intravenously
the uptake of the drug nanocarriers by the cells. As shown in administrated. In contrast, the other mice groups, receiving
Figure 5d, after 3 h of incubation, the FA-conjugated nano- lower doses of nanocarriers, were healthy and <10% weight
carriers (HMS@FTD) showed a much higher MFI in the loss was detected over two weeks. Therefore, the MTD of
folate receptor positive cells (KB cells) than that in the other the HMS@FTD nanocarriers was around 40 mg DOX per kg
two systems [FA-conjugated nanocarriers (HMS@FTD) in body weight. This value is much higher (also an indicative of
the folate receptor negative cells (7402 cells) and FA-free lower cytotoxicity) than that of free DOX (8 mg DOX per kg
nanocarriers (HMS@TD) in the folate receptor positive cells body weight). Figure S5 (Supporting Information) presents
(KB cells)]. HeLa cells were also used in the flow cytometry the in vivo pharmacokinetics of the HMS@FTD nanocar-
studies, and the results are shown in Figure S4 (Supporting riers after their intravenous administration to mice. A slow
Information), which also confirm the targeting properties of decrease in the DOX concentration from the DOX-loaded
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HMS@FTD nanocarriers was detected in the blood over 24 h the HMS@FTD nanocarriers, a control experiment was car-
after injection, which suggests that the HMS@FTD nanocar- ried out in which the mice were pre-dosed with excess folic
riers had a long systemic circulation and slow elimination acid (10 mg kg−1) prior to the injection with the nanocarriers,
in the blood. Such features are pivotal in designing efficient which annulled almost all folate receptors in the tumor cells.
drug-delivery nanocarriers with a high clinical efficacy.
As expected, after the intravenous injection of the HMS@
In vivo studies of the as-fabricated HMS@FTD nano- FTD nanocarriers (24 h after injection), no obvious fluores-
carriers were carried out using female athymic nude mice cence (Figure 6c) was observed in the tumor site. This data
bearing HeLa (folate receptor positive) tumors with a tumor further confirms the crucial role of the targeting FA ligands
size of around 0.5 cm (after 3 weeks post inoculation of about in the HMS@FTD nanocarriers.
1 × 106 cells on the right foreleg) as the animal model. The
To study the blood circulation of the nanocarriers, Blood
fluorescence image (from the red-emitting RhB labeled was drawn at different time points post injection and solu-
on the HMS@FTD nanocarriers) was used to evaluate the bilized by a lysis buffer. The HMS levels in the blood sam-
uptake of the drug nanocarriers. As shown in Figure 6b , a ples could be measured by UV-vis spectroscopy due to the
strong fluorescence was observed in the tumor at around 24 h modified FITC, according to previous reports. The blood cir-
after the intravenous injection of the DOX-loaded HMS@ culation data of the HMS@FTD nanocarriers are shown in
FTD nanocarriers. In contrast, no visible fluorescence was Figure S6 in the Supporting Information, which confirms a
detected in the tumor of the mice after 24 h if the fluorescent long blood circulation half-life.
HMS NPs (without the FTD capping ligands) were injected
The in vivo distribution of the DOX-loaded HMS@
intravenously (Figure 6a). This data provides supporting FTD nanocarriers and the pristine fluorescent HMS NPs
evidence for the crucial role of the multifunctional FTD (without the amphiphilic FTD capping agents) is presented
capping ligands in the HMS@FTD nanocarriers. To further in Figure 7. The fluorescence intensity of the HMS@FTD
affirm the in vivo targeting capability of the FA ligands in nanocarriers in the tumor was significantly higher than
that in other major organs. This intensity
was also much stronger than that of the
HMS NPs in the tumor, further demon-
strating the high targeting efficiency of
the designed FTD capping ligands. The
HMS@FTD nanocarriers were also barely
found in normal organs, indicating that
the nanocarriers had a prolonged and suf-
ficient blood circulation for the efficient
targeting and delivery of the drugs to the
tumor. The in vivo distribution of Si from
the HMS@FTD nanocarriers was further
examined by inductively coupled plasma
atomic emission spectrometry (ICP-AES).
As shown in Figure S4 (Supporting Infor-
mation), the results were also consistent
with the biodistribution data from the flu-
orescence images (Figure 7, and Figure S8,
Supporting Information), further con-
firming that the as-designed nanocarriers
were mainly accumulated in the tumor.
The suitable size of the as-designed nano-
carrier in combination with its biocom-
patible PEG-like capping shell, may have
led to its prolonged blood circulation,
reduced nonspecific binding, and the final
preferential deposition in the tumor. To
further evaluate whether the improved
biodistribution can enhance the anti-
tumor efficacy, the changes in tumor size
after intravenous injection of saline, pris-
tine fluorescent HMS NPs, free DOX, and
DOX-loaded HMS@FTD nanocarriers
were investigated. As shown in Figure 8,
the best tumor growth inhibition was a
shrinkage to about 5.12% of the control
group tumor, which was achieved in the
Figure 6. In vivo fluorescence images of the subcutaneous HeLa tumor-bearing athymic
nude mice (right foreleg, indicated by the red circle) after an intravenous injection of
500 µL of a) fluorescent HMS NPs (1 mg mL−1) and b) HMS@FTD nanocarriers (1 mg mL−1).
c) A blocking dose of folic acid was first injected into the mouse to neutralize the folate
receptors in the tumor cells followed by an injection of 500 µL HMS@FTD nanocarriers
(1 mg mL−1). The fluorescence images were achieved at 24 h after injection.
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ligands showed complementary properties (the HMS NPs
provided a physical and structural stability and the FTP
ligands provided the chemical specificity for tumor cell tar-
geting). These properties were then independently tailored
and integrated into a single HMS@FTD nanocarrier to
achieve efficient and targeted drug delivery. Both in vitro
and in vivo data suggested that the as-fabricated HMS@FTD
nanocarriers are biocompatible and are able to deliver the
hydrophobic drug to the target tumor cells. The nanocarriers
could also unload the drug payload directly into the affected
cells, where the weakly acidic environment inside the cells
triggered the hydrolysis of the pH-labile linkages in the FTD
capping ligands. The smart drug carrier in this study is a good
prospect for further clinical studies.
Figure 7. Fluorescence intensities of the dissected organs of sacrificed
mice bearing the HeLa tumor 24 h after the intravenous injection
of pristine fluorescent HMS NPs (green) or DOX-loaded HMS@FTD
nanocarriers (blue).
4. Experimental Section
Chemicals: 1-bromododecane, p-toluenesulfonic acid (PTSA),
anhydrous potassium carbonate, glycerol, di-t-butyl dicarbonate
(BOC)-anhydride, folic acid, dicyclohexacarbodiimide (DCC) and
N-hydroxysuccinimide (NHS) were purchased from Sinopharm
Chemical Reagent Co. Ltd. (2-(Acryloyloxy)ethyl) trimethylammonium
chloride (AETAC, 80 wt% in water), tetraethylorthosilicate (TEOS), hex-
adecyltrimethylammonium bromide (CTAB, >99.0%), 2,2-azobis(2-
mice group injected with DOX-loaded HMS@FTD nanocar-
riers. In contrast, no obvious inhibition was observed in the
mice group injected with the pristine fluorescent HMS NPs.
Free DOX showed an inhibitory effect to about 54.43%
shrinkage of the tumor as compared to that of the control
group. The amount of shrinkage is thus much lower than
that of the DOX-loaded HMS@FTD nanocarriers. Taken
together, these data confirmed that the incorporation of the
FTD moieties in the as-designed drug carrier has significantly
enhanced the antitumor efficacy on the systemic level.
methylpropionamidine)
dihydrochloride
(V-50,
>97.0%),
fluorescein isothiocyanate (FITC), rhodamine B isothiocyanate
(RhB), 3-aminopropyltriethoxysilane (APTES), 18-crown-6, octade-
cyltrimethoxysilane (OTMS), 4,7,10-trioxa-1,13-tridecanediamine
(TDA), and cyanuric chloride were obtained from Sigma-Aldrich Co
(Shanghai, China). Styrene (St, >99.0%) was washed through an
inhibitor remover column and then distilled under reduced pressure
prior to use. Other reagents were used as received. All of the animal
procedures were performed in compliance with Soochow University
Guidelines for the Care and Use of Laboratory Animals.
3. Conclusions
Synthesis of Hollow Mesoporous Silica (HMS) Nanoparticles:
The templating polystyrene nanoparticles (PS NPs) were synthe-
sized by an emulsifier-free emulsion polymerization. In a typical
synthesis of 120 nm PS NPs, 500 mg of AETAC (80 wt% in H2O)
was dissolved in 180 g of water in a 500 mL round-bottomed flask,
followed by a slow addition of 20 g of styrene. The mixed solu-
tion was stirred for 30 min, purged with nitrogen for 20 min, and
heated up to 90 °C. A 5 mL aqueous solution containing 500 mg of
V-50 was then introduced. The emulsion was kept at 90 °C for 24 h
under nitrogen protection for the complete polymerization. The PS
NPs were collected by centrifugation and washed copiously with
ethanol for several times.
A smart core–shell structured drug nanocarrier was devel-
oped in this study using HMS NPs as the core and specifi-
cally designed multifunctional amphiphilic FTD ligands as
the encapsulating shell. The HMS NPs and amphiphilic FTD
In a typical synthesis of the HMS NPs, 0.8 g of CTAB was
first dissolved in a mixed solvent containing 80 g of water, 60 g
of ethanol, and 1.5 mL of ammonium hydroxide solution (28%).
930 mg of PS NPs was then dispersed in water (10.0 g) by soni-
cation. The dispersed PS NPs were added dropwise to the above
CTAB solution at room temperature and under vigorous stirring.
The mixed solution was sonicated for 10 min. The milky mixture
was then stirred for 30 min followed by the addition of 4.0 g of
TEOS. The mixture was kept stirring at room temperature for 12 h.
The precipitate was washed copiously with ethanol and dried at
room temperature. The product was then calcined in air at 600 °C
for 6 h, which completely removed the templating PS NPs and the
Figure 8. Changes in tumor volume after intravenous injection of
saline, pristine fluorescent HMS NPs, free DOX, and DOX-loaded HMS@
FTD nanocarriers in Hela tumor-bearing nude mice.
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pore-generating agents CTAB. The HMS NPs were then collected was stirred for 12 h at room temperature in the dark. The solution
and stored in a desiccator. was then filtered off and precipitated in diethyl ether. The resulting
Synthesis of Hydrophobic FITC or RhB-Doped Fluorescent HMS yellow powder was washed several times with anhydrous ether
NPs: The preparation and modification of the organic dye (FITC or and used immediately for the next step. FA-NHS (1.2 g, 2.2 mmol)
RhB) labeled HMS NPs were performed at room temperature. In was completely dissolved in 100 mL of dry pyridine, followed by
brief, 190 mg of APTES was introduced into an ethanol solution a slow addition of BOC-TDA (0.73 g, 2.27 mmol) over 30 min. The
(10 mL) containing 37 mg of FITC or RhB. The mixed solution was mixture was stirred for 12 h at room temperature in the dark. After
stirred for 10 h, and the resultant solution was added to 5 mL of evaporating the pyridine, the resulting compound was dissolved
ethanol solution containing 100 mg of HMS NPs. The reaction was in 5 mL of trifluoroacetic acid (TFA) to remove the BOC group. The
allowed to proceed overnight, leading to the formation of fluores- deprotection step was carried out at room temperature for 4 h. TFA
cent HMC NPs. To modify the surface of the fluorescent HMS NPs was then removed under vacuum. The resulting compound was
from hydrophilic to hydrophobic, 5 mL of OTMS was introduced loaded on a DEAE Sephadex A25 column packed with potassium
into the HMS NP solution. The mixture was then stirred overnight. tetraborate, and the compound was eluted with 10–50 mM ammo-
The product was collected by centrifugation, washed several times nium bicarbonate. All fractions were collected and lyophilized.
with acetonitrile and ethanol, and then dried under vacuum.
1H NMR (400 MHz, DMSO-d6), δ [ppm]: 8.60 (s, 1H), 8.08–7.76
Synthesis of 4-n-dodecyloxybenzaldehyde (DBD): 1-bromodo- (m, 6H), 6.75 (d, 2H), 4.56 (s, 2H), 4.40–4.76 (m, 1H), 3.68–3.37
decane (29.9 g, 120 mmol) was added dropwise into the mixture (m, 14H), 3.37 (d, 2H), 2.88–2.82 (m, 2H), 2.53 (s, 2H), 2.39–2.21
of p-hydroxybenzaldehyde (12.2 g, 100 mmol) and anhydrous (m, 2H), 1.76 (t, 2H), 1.60–0.92 (m, 4H). 13C NMR (75 MHz, DMSO-
potassium carbonate (20.7 g, 150 mmol) in 150 mL of acetone. d6), δ [ppm]: 175.0, 172.9, 166.9, 161.3, 158.5, 153.6, 151.8,
After heating under reflux with stirring for 14 h, the mixture was 148.7, 130.0, 128.8, 121.8, 111.9, 70.3–69.3, 68.8, 67.5, 46.2,
filtered off and acetone was removed by a rotary evaporator. The 36.9, 35.7, 30.2, 29.3, 27.2.
crude product was purified by column chromatography (ethyl
Synthesis of 2,4-dichloro-dodecyloxybenzalacetal-1,3,5-tria-
acetate/petroleum ether, 1:10). Anal. Calcd. for C19H30O2: C, zine (TsT-DBA) and FA-TDA-DBA: 423 mg of potassium carbonate
78.57%, H, 10.41%, O, 11.02%. Found: C, 78.48%, H, 10.53%, and 25 mg of 18-crown-6 were added to 10 mL of a toluene solu-
1
O, 10.99%. H NMR (400 MHz, CDCl3), δ [ppm]: 9.88 (s, 1H), 7.83 tion containing 568 mg of cyanuric chloride. DBA (1.1 g) in 5 mL of
(d, 2H), 6.99 (d, 2H), 4.04 (t, 2H), 1.86–1.76 (m, 2H), 1.46 (m, toluene was then added dropwise under nitrogen. The reaction was
2H), 1.26 (m, 16H, CH2), 0.88 (t, 3H). 13C NMR (75 MHz, CDCl3) allowed to proceed under refluxing for around 18 h. The product
δ [ppm]: 191.12, 164.50, 132.24, 129.91, 114.96, 68.65, 32.16, was collected, passed through a plug of Celite, concentrated in a
29.29–29.88, 26.19, 22.94, 14.39. High resolution mass spectro- rotary evaporator, and dried overnight under vacuum. The FA-TDA
meter (HRMS) calcd. for C19H30O2 [M+H]+ 291.2206, found 291.2211. was conjugated following the procedures described above to
1
Synthesis of 4-n-dodecyloxybenzalacetal (DBA): In a typical obtain FA-TDA-DBA. H NMR (400 MHz, DMSO-d6), δ [ppm]: 8.60
synthesis, DBD (8.7 g, 30 mmol) was allowed to react with glyc- (s, 1H), 7.82–6.95 (m, 10H), 6.65 (d, 2H), 5.21 (s, 2H), 4.40–3.58
erol (2.76 g, 30 mmol) in 50 mL of toluene using PTSA (0.5 g) as (m, 10H), 3.48–2.90 (m, 16H), 2.88 (d, 2H), 2.49–2.36 (m, 2H),
a catalyst. The solution was refluxed under vigorous stirring for 1.73 (d, 2H), 1.70–1.41 (m, 4H), 1.61–0.90 (m, 24H). Anal. Calcd.
14 h, and the water formed by the dehydrogenation reaction was for C54H75ClN12O12: C, 57.92%, H, 6.75%, Cl, 3.17%, N, 15.01%,
removed by the oil/water separator. The mixture was then evapo- O, 17.15%.
rated and washed with an aqueous solution of potassium carbonate
Loading and Release Profiles of Drugs in the As-fabricated
(1 wt%, 80 mL) to remove the acid catalyst and the remaining HMS@FTD Nanocarriers: Doxorubicin (DOX) was chosen as the
glycerol. Afterwards, the precipitate was filtered off and purified model drug to evaluate the loading and release profiles in the as-
by column chromatography (ethyl acetate-petroleum ether, 1:2). fabricated HMS@FTD nanocarriers. The water-insoluble DOX was
Anal. Calcd. for C22H36O4: C, 72.49%, H, 9.95%, O, 17.56%. Found: extracted from doxorubicin hydrochloride (DOX·HCl).[48] The DOX
1
C, 72.36%, H, 10.04%, O, 17.60%. H NMR (400 MHz, CDCl3), δ solution (5 mg mL−1) was added to 0.8 mL of the as-prepared HMS
[ppm]: 7.41 (d, 2H), 6.90 (d, 2H), 5.51 (s, 1H), 3.57–4.38 (m, 8H), and FA-TDA-DBA in tetrahydrofuran, followed by a slow addition of
1.86–1.76 (m, 2H), 1.50–1.39 (m, 2H), 1.26 (m, 16H), 0.88 (t, 3H). 10 mL of phosphate buffer (0.02 M, pH 7.4). The mixed solution
13C NMR (75 MHz, CDCl3), δ [ppm]: 191.15, 132.25, 127.36, 114.48, was shaken for 24 h to allow the diffusion of DOX into the NPs. The
101.93, 72.48, 68.28, 64.21, 32.17, 29.29–29.88, 26.22, 22.95, DOX-loaded HMS@FTD nanocarriers were then centrifuged, and
14.39. HRMS calcd. for C22H36O4 [M+H]+ 365.5126, found 365.5121. the free DOX was removed. The concentration of free DOX in solu-
Synthesis of BOC-4,7,10-trioxa-1-tridecaneamine (BOC-TDA): tion was determined by measuring the UV absorbance at 485 nm.
3.7 g of TDA was dissolved in 1,4-dioxane (80 mL) and treated with To determine the amount of drugs encapsulated inside the nano-
BOC-anhydride (1.8 g). The mixture was stirred at room temperature carriers, a standard plot was prepared under identical conditions.
overnight, followed by the removal of the solvent. The resulting yellow
The free DOX concentration was studied after the centrifuga-
oil was purified by silica gel chromatography (methanol-dichlo- tion and the loading efficiency was calculated as follows:
romethane, 1:10). 1H NMR (400 MHz, CDCl3), δ [ppm]: 5.00 (s, 1H),
3.52–3.48 (m, 12H), 3.20 (d, 2H), 2.79 (t, 2H), 1.71–1.65 (m, 4H),
(1)
loading efficiency =(C0V0 −CtVt )/C0V0
1.58 (s, 2H), 1.42 (s, 9H). 13C NMR (75 MHz, CDCl3), δ [ppm]: 155.6,
69.5–68.9, 66.2, 48.5, 36.9, 30.2, 28.5, 27.2.
Synthesis of FA-4,7,10-trioxa-1-tridecaneamine (FA-TDA): 0.62 g
of DCC and 0.51 g of NHS were mixed with 50 mL of dry dimethylfor-
mamide solution containing 2.0 g of folic acid. The mixed solution
where C0 and V0 were the concentration and volume of the added
DOX, and Ct, Vt were the concentration and volume of the free DOX
after centrifugation. To assure the accurate determination of this
value, the experiments were carried out three times and a mean
value was achieved.
© 2015 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
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The drug-release test was performed by suspending the DOX- were then centrifuged, washed three times with cold PBS buffer,
loaded HMS@FTD nanocarriers in phosphate-buffered saline (PBS) and resuspended in PBS buffer. Flow cytometry was performed on
buffer (pH 7.4). The mixed solution was shaken in a water bath at a BD FACS Calibur instrument.
a constant temperature of 37 °C. To determine the release amount
Tumor Xenografts and In Vivo Imaging: HeLa tumor cells were
of the drug at a particular time, 1.0 mL of the solution was with- harvested by centrifuging, and resuspending in sterile PBS buffer.
drawn after centrifugation, and the same volume of PBS buffer was The tumor cells (1 × 106 cells/site) were then implanted sub-
introduced to keep a constant volume. To determine the release cutaneously into the right foreleg of female athymic nude mice
amount at a particular pH, the obtained colloid was adjusted to a (4 weeks old). Biodistribution and imaging studies were performed
certain pH by acetate buffer. The drug concentration in the with- when the tumor size reached around 0.5 cm (3 weeks after post-
drawn solution was analyzed by measuring the UV absorbance inoculation). In vivo fluorescence imaging was performed on a
at 485 nm. The experiments were conducted in triplicate and Kodak DXS 4000 PRO system. The fluorescence intensities were
the results were presented as an average value with standard analyzed using Kodak Molecular Imaging Software.
deviations.
Cell Culture and Preparation: Human epidermoid carcinoma
(KB cells, with over-expressed folate receptors or FR positive)
and hepatoma 7402 (without over-expressed folate receptors
or FR negative) cell lines (purchased from Shanghai Cell Institute
Country Cell Bank, P. R. China) were cultured at 37 °C in a humidi-
Supporting Information
fied incubator (5% CO2 in air, v/v) as a monolayer in the RPMI-
1640 medium supplemented with 10% of heat-inactivated fetal
bovine serum.
Supporting Information is available from the Wiley Online Library
or from the author.
In Vitro Cytotoxicity: Sulforhodamine B (SRB) assay was used to
assess the potential cytotoxicity of the as-fabricated drug carriers.
In brief, hepatoma 7402 or KB cells were seeded in 96-well plates
(1.3 × 104 cells per well). Four duplicate wells were set up for each
sample. The culture medium was replaced with the medium con-
taining different concentrations of DOX-free or DOX-loaded HMS@
FTD nanocarriers, and then cultured at 37 °C in a humidified incu-
bator (5% CO2 in air, v/v). After 72 h, the medium was poured
away and 10% (w/v) trichloroacetic acid in Hank’s balanced salt
solution (100 µL) was added, and stored at 4 °C for 1 h. The sta-
tionary liquid was then discarded, and the cells were washed with
copious amounts of water for five times before air drying and they
were then stained with 0.4% (w/v) SRB solution (100 µL per well)
for 30 min at room temperature. After the removal of free SRB, the
cells were washed with 0.1% acetic acid solution for five times.
The bound SRB dye was solubilized in a 10 mmol L−1 tris-base
buffer (150 µL, pH 10.5). The optical density (OD) was determined
by the optical absorbance at 531 nm of individual wells by a
spectrophotometer.
Cellular Uptake of DOX-Loaded HMS@FTD Nanocarriers: The
KB cells and Hepatoma 7402 cells were seeded in 96-well plates
(1.3×104 cellsperwell)andincubatedovernightat37°Cinahumidi-
fied incubator. The dispersion was prepared in RPMI-1640 medium.
The DOX concentration in the HMS@FTD (FA-conjugated) and
HMS@TD (FA-free) nanocarriers was identical, namely 10 µg mL−1.
The cells were washed twice with PBS buffer and incubated with
the above solutions for 0.5 or 3 h. The culture media were removed
and the cells were washed three times with PBS buffer before
testing. The cells were observed under a Nikon AY laser scanning
confocal microscope. All images were gathered at the same set-
tings and processed with Nikon AY software.
Acknowledgements
This work was supported by the Collaborative Innovation Center
of Suzhou Nano Science and Technology. The authors grate-
fully acknowledge the financial support provided by the National
Natural Science Foundation of China (21336005, 21301125), the
Natural Science Foundation of Jiangsu Province (BK2012625), and
the Suzhou Nano-project (ZXG2013001, ZXG201420).
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Received: October 15, 2015
Published online: November 30, 2015
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