Horseradish Peroxidase-Encapsulated Hollow Silica Nanospheres for Intracellular Sensing of Reactive Oxygen Species
Chen et al. Nanoscale Research Letters
Horseradish Peroxidase-Encapsulated Hollow Silica Nanospheres for Intracellular Sensing of Reactive Oxygen Species
Hsin-Yi Chen 1
Si-Han Wu 0 2
Chien-Tsu Chen 4
Yi-Ping Chen 0 2
Feng-Peng Chang 1
Fan-Ching Chien 3
Chung-Yuan Mou 1
0 Graduate Institute of Nanomedicine and Medical Engineering, College of Biomedical Engineering, Taipei Medical University , No. 250 Wuxing St., Taipei 11031 , Taiwan
1 Department of Chemistry, National Taiwan University , No. 1, Sec. 4, Roosevelt Rd., Taipei 10617 , Taiwan
2 International PhD Program in Biomedical Engineering, College of Biomedical Engineering, Taipei Medical University , No. 250 Wuxing St., Taipei 11031 , Taiwan
3 Department of Optics and Photonics, National Central University , No. 300, Zhongda Rd., Taoyuan City 32001 , Taiwan
4 Department of Biochemistry, Taipei Medical University , No. 250 Wuxing St., Taipei 11031 , Taiwan
Reactive oxygen species (ROS) have crucial roles in cell signaling and homeostasis. Overproduction of ROS can induce oxidative damage to various biomolecules and cellular structures. Therefore, developing an approach capable of monitoring and quantifying ROS in living cells is significant for physiology and clinical diagnoses. Some cellpermeable fluorogenic probes developed are useful for the detection of ROS while in conjunction with horseradish peroxidase (HRP). Their intracellular scenario is however hindered by the membrane-impermeable property of enzymes. Herein, a new approach for intracellular sensing of ROS by using horseradish peroxidase-encapsulated hollow silica nanospheres (designated HRP@HSNs), with satisfactory catalytic activity, cell membrane permeability, and biocompatibility, was prepared via a microemulsion method. These HRP@HSNs, combined with selective probes or targeting ligands, could be foreseen as ROS-detecting tools in specific organelles or cell types. As such, dihydrorhodamine 123-coupled HRP@HSNs were used for the qualitative and semi-quantitative analysis of physiological H2O2 levels in activated RAW 264.7 macrophages. We envision that this HSNs encapsulating active enzymes can be conjugated with selective probes and targeting ligands to detect ROS in specific organelles or cell types of interest.
Reactive oxygen species; Hollow silica nanospheres; Horseradish peroxidase; Enzyme delivery; Porous materials
Reactive oxygen species (ROS) consisting of radical and
non-radical molecules, such as superoxide anions,
hydrogen peroxide, hydroxyl radical, singlet oxygen, and
peroxynitrite, are continuously produced during aerobic
metabolism. Cellular ROS are mainly generated from the
mitochondrial electron transport chain (mETC) and are
normally counterbalanced by enzymatic (such as
superoxide dismutases, catalases, and peroxidases) and
nonenzymatic (e.g., vitamins A, C, and E; urate; and bilirubin)
antioxidant defenses [
]. However, imbalances in ROS
production can lead to oxidative stress and subsequent damage
to DNA, fatty acids, proteins, and other cellular
components, potentially contributing to diabetes [
], cancer [
and cardiovascular disorders [
], and neurodegenerative
] such as Alzheimer’s disease and Parkinson’s
disease. Direct imaging and ROS quantification in living
cells are highly desirable but very challenging.
Advances in fluorescent microscopy [
] have allowed
the development of the noninvasive measurement and
imaging of ROS evolution at the single-cell level. To detect
ROS, most probes are designed to measure changes in the
fluorescence intensity or shifts in the emission wavelength
(i.e., ratiometric methods) following oxidization of
profluorescent aromatic molecules or deprotection of masked
compounds to fluorescent products . Specificity to a
particular type of ROS is of importance when designing
successful probes; for example, boronate oxidation is
utilized as a bioorthogonal reaction approach for studying
the chemistry of hydrogen peroxide in living systems [
To explore the spatio-temporal dynamics of ROS, several
boronate-based probes conjugated with a positively
charged phosphonium moiety were generated for
mitochondrial targeting [
]. However, their potential for
in vivo imaging is limited by their instability in biological
milieu, low penetration of tissue barriers, and rapid
elimination from the body through the urinary system [
]. To overcome such problems, some strategies have
been developed either by chemically grafting an additional
stabilizing structure onto the probe  (e.g., a triethylene
glycol chain), developing genetically encoded fluorescent
protein-based indicators [
], or applying reaction-based
bioluminescent reporters [
] or positron emission
tomography (PET) probes for the molecular imaging of ROS
]. Moreover, several comprehensive studies highlighted
nano-formulations as an important design consideration
and demonstrated that nanoparticle-based probes can
provide mechanistic insights and innovative strategies to
image ROS in living organisms with high specificity and
]. Enzymes with high catalytic activity
and distinct substrate selectivity have also been utilized as
clinical diagnostic tools for identifying target analytes.
However, the lack of lasting stability and the difficulty in
permeating through biological membranes of free
enzymes have often limited their applications in complex
biological milieu. Even though applying electrode is not
suitable for intracellular assays or in vivo imaging,
considerable efforts have been devoted to developing horseradish
peroxidase (HRP)-incorporated biosensors to determine
H2O2 based on electrochemical methods [
In this work, enzymatic nanoreactors, composed of
HRP encapsulated in 45-nm hollow silica nanospheres,
were synthesized by a water-in-oil (w/o) microemulsion
route followed by a mild etching process [
Previously, we demonstrated that such hollow nanomaterials
can maintain stable activity of the encapsulated enzymes
and nanocatalysts while protecting against proteolysis
and sintering, respectively [
]. In this work, we
evaluated their potential uses as intracellular biosensors
by studying the enzyme entrapment efficiency, loading
capacity, peroxide reactivity and selectivity, cellular
uptake, toxicity, and proliferation effects of HRP@HSNs.
Using dihydrorhodamine 123 (DHR123) as a substrate,
which has been commonly coupled with HRP to detect
intracellular hydrogen peroxide production, interactions
between HRP@HSNs and various types of ROS in
aqueous solutions were investigated by flow cytometry and
fluorescence microscopy. Furthermore, it was
demonstrated that utilization of HRP@HSNs with DHR123 can
simultaneously image and quantify physiological H2O2
levels in phorbol 12-myristate 13-acetate
(PMA)-stimulated RAW264.7 macrophages. Taken together, the
enzymatic nanoreactors of HRP@HSNs have the potential
for imaging ROS-associated inflammatory cells in vivo
and the encapsulated components may be extended to
multiple different enzymes , nanoparticles [
recognition molecules for synergistic applications.
Chemicals and Reagents
Decane, n-hexanol (98%), ammonium hydroxide (NH4OH,
35 wt%), tetraethyl orthosilicate (TEOS, 98%),
3aminopropyltrimethoxysilane (APTMS, 95%), and
fluorescein isothiocyanate (FITC) isomer were purchased from
ACROS. Polyoxyethylene (5) isooctylphenyl ether (Igepal
CA-520), HRP type VI-A (HRP),
3,3′5,5′-tetramethylbenzidine (TMB), citric acid, dimethyl sulfoxide (DMSO), and
rhodamine B isothiocyanate (RITC) were purchased from
2-(4-Iodophenyl)-3-(4-nitrophenyl)-5-(2,4disulfophenyl)-2H-tetrazolium was purchased from
Clontech. DHR123 and PMA were purchased from Cayman
Chemical. Hydrogen peroxide (H2O2, 35%) was purchased
from SHOWA Chemical Industry. Tert-butyl
hydroperoxide solution (70% in H2O) was purchased from Aldrich.
Iron (II) perchlorate (Fe(ClO4)2) was purchased from Alfa
Aesar. Ultrapure deionized (D.I.) water was generated by a
Millipore Milli-Q Plus system. All reagents were used
without further purification.
Synthesis of Hollow Silica Nanospheres (HSNs)
HSNs were synthesized by a reverse microemulsion
system accompanied by a selectively etching method as
described in our previous studies [
20 mL of decane as the oil phase, 1.63 mL of CA-520 as
a surfactant, and 550 μL of n-hexanol as a co-surfactant
were mixed as well as magnetically stirred with a 2-cm
PTFE-coated stir bar at 650 rpm. After that, 350 μL of
D.I. water were added to the mixture at room
temperature, generating a water-in-oil (w/o)
microemulsion system. Next, 25 μL of APTMS ethanolic solution
(200 μL of APTMS in 1.4 mL of absolute ethanol) and
100 μL of TEOS were added with stirring. After stirring
for 10 min, 250 μL of aqueous ammonia (35 wt%) was
introduced into the system with stirring at 20 °C. After
10 h, 95% ethanol was added to destabilize the
microemulsion system and the solid silica nanoparticles (SSNs)
were collected by centrifugation at 11,000 rpm for
20 min. To obtain HSNs, the SSNs were suspended in
D.I. water with stirring at 40 °C for 40 min. Next, the
HSNs were collected by centrifugation at 11,000 rpm for
20 min and washed with 95% ethanol several times.
Finally, the HSNs were suspended and kept in 99.5%
Synthesis of Horseradish Peroxidase-Encapsulated Hollow
Silica Nanospheres (HRP@HSNs)
HRP@HSNs were synthesized by a method based on our
previous studies [
]. Typically, the synthesis is similar
to the procedure above, except that 350 μL of D.I. water
was replaced by 350 μL of aqueous HRP (90 μL of 10 mg/
mL of an HRP solution in 350 μL of D.I. water). After
synthesis, HRP@HSNs were kept in D.I. water at 4 °C.
Synthesis of FITC-HSNs and HRP@FITC-HSNs
HSNs and HRP@HSNs with incorporated green-emitting
fluorescein dye (designated FITC-HSNs and
HRP@FITCHSNs) were synthesized similar to the procedure above,
except that ethanolic APTMS solution was replaced by an
FITC-APTMS one. An ethanolic FITC-APTMS solution
was prepared by mixing 10 mg of FITC and 200 μL of
APTMS with 1.4 mL of absolute ethanol under dark
conditions for 18 h at room temperature.
HRP Entrapment Efficiency and Loading Capacity of
First, a mixture comprised of HRP (6 mg in 500 μL of
D.I. water) and RITC (3 mg in 350 μL of DMSO) was
stirred under dark conditions for 24 h at 4 °C. After that,
the mixture was transferred to a dialysis membrane
composed of regenerated cellulose with a molecular
weight cutoff of 12~14 kDa. Then, to remove the
unreacted RITC, the dialysis bag was dialyzed against
1 L of D.I. water and gently stirred for 3 days. Finally,
the RITC-labeled HRP (designated RITC-HRP) was used
to synthesize RITC-HRP@HSNs.
To determine the HRP loading capacity,
RITCHRP@HSNs were dissolved in 1 mL of NaOH (1 M) for
1 h, and the amount of the entrapped RITC-HRP was
calculated from a calibration curve established by plotting
the fluorescence intensity versus the concentration of
RITC-HRP. The fluorescence was measured with a Hitachi
F-4500 Instrument at an excitation wavelength of 543 nm
and an emission wavelength of 550~650 nm. The HRP
entrapment efficiency and loading capacity of HRP@HSNs
were defined as follows: entrapment efficiency (%) = mass
of RITC-HRP in RITC-HRP@HSNs/initial mass of
RITCHRP; and loading capacity = mass of RITC-HRP in
HRPRITC@HSNs/mass of RITC-HRP@HSNs.
HRP Activity Assay
To detect the activity of the peroxidase enzyme, a
chromogenic substrate of TMB was used. TMB can be
converted into a colored product when oxidized by HRP
using hydrogen peroxide as the oxidizing agent. First,
various concentrations of native HRP and HRP@HSN
were prepared in phosphate and citrate buffer (pH 5.2).
Then, each solution was supplemented with 50 μL of the
TMB solution (20 μM in DMSO) and 50 μL of H2O2
(20 μM in D.I. water). The reaction was monitored by
measuring the absorbance at 655 nm using a microplate
reader (BioTek Synergy Hybrid Reader). The activity of
HRP encapsulated in HSNs was calculated from the
calibration curve of native HRP.
Reactivity Assay of HRP@HSNs to Various ROS
DHR123 (20 μM) alone or mixed with HRP@HSNs
(50 μg/mL) was incubated with various types of ROS
(100 μM) in 100 μL of DMEM solution (pH 7.4). The
fluorescence emission at 530 nm (λex = 488 nm) was
monitored every 5 min for the first 120 min. The ROS
investigated were obtained as follows: hydrogen peroxide
(H2O2) and tert-butyl hydroperoxide (TBHP) were
prepared from commercially available 32 and 70%
aqueous solutions, respectively. Superoxide (O2•−) was
generated from 10 mM stock of potassium superoxide
(KO2) in DMEM. Hydroxyl radicals (•OH) and
tertbutoxy radicals (•OtBu) were produced by the reaction
of 1 mM Fe(ClO4)2 with 100 μM H2O2 or 100 μM
Cell Culture and Viability Assay
The RAW264.7 mouse macrophage cell line was
obtained from ATCC. RAW264.7 cells were maintained in
DMEM with 10% FBS, 100 U/mL penicillin, and 100 μg/
mL streptomycin (Gibco) at 37 °C in 5% CO2
atmosphere. Typically, 2 × 105 RAW264.7 cells per well were
seeded in 24-well plates for the viability assays. After
24 h, cells were washed twice with PBS and incubated
with different amounts (0, 50, 100, and 200 μg/mL) of a
nanoparticle suspension in serum-free DMEM for 2 h.
For the cytotoxicity assay, nanoparticle-treated cells
were washed twice with culture medium followed by
incubation with WST-1 reagent (Clontech) at 37 °C for
2 h. For the proliferation assay, cells after treatment with
nanoparticles for 2 h were allowed to grow in regular
growth medium for 24 h followed by incubation with
the WST-1 reagent. Cell viability was determined by the
formazan dye generated by live cells, and the absorbance
at 450 nm was measured, with a reference wavelength of
650 nm, using a microplate reader (Bio-Rad, model 680).
Cell Uptake Analysis
RAW264.7 cells at 1 × 106 per well were seeded in
sixwell plates overnight. Then, RAW264.7 macrophages
were treated with different amounts (0, 50, 100, and
200 μg/mL) of a nanoparticle suspension in serum-free
DMEM media for 2 h. After that, cells were washed
three times with PBS and detached by a trypsin-EDTA
solution. The uptake of nanoparticles by RAW264.7
macrophages was examined by flow cytometry. Trypan
blue was utilized to quench the fluorescence of
nanoparticles adsorbed onto the exterior membrane of cells.
Flow Cytometry Analysis of ROS Production in
PMA-Stimulated RAW264.7 Macrophages
Typically, after 2 h of treatment of RAW264.7
macrophages with nanoparticles, cells were washed three times
with PBS followed by incubation with 20 μM DHR123
in serum-free DMEM for 30 min. Then, RAW264.7 cells
were washed with PBS and incubated with culture
medium containing PMA at different concentrations for
1 h. After washing, RAW264.7 macrophages were
harvested and analyzed by a FACS Canto II flow cytometer.
RAW264.7 cells at 3 × 104 per well were seeded in 96-well
plates for semi-quantitative assays. After incubation with
50 μL of 100 μg/mL of a nanoparticle suspension in
serum-free DMEM for 2 h, nanoparticle-treated cells were
treated with 50 μL of serum-free DMEM containing
different concentrations of PMA, and 20 μM DHR123 for an
additional 1 h at 37 °C. At the same time, external
standards of H2O2 mixed with 50 μg/mL HRP@HSNs were
used to develop a calibration curve by plotting the
fluorescence intensity versus the concentration of H2O2. The
fluorescence intensity was measured with a microplate
reader (BioTek Synergy Hybrid Reader) with excitation at
488 nm and emission at 530 nm. Using the established
calibration curve, amounts of H2O2 in RAW264.7 cells
stimulated with various amounts of PMA were calculated.
Transmission electron microscopic (TEM) images were
taken on a JEOL JEM-1200 EX II operating at 100 kV.
Images were recorded with a GatanOrius CCD camera.
Samples were dispersed in 95% ethanol and dropped
onto a carbon-coated copper grid, and then air-dried
and examined. To verify HRP in the hollow spheres, a
negative staining sample was stirred in 1% aqueous
uranyl acetate (UA) for 1 h, and then centrifuged to
remove the remaining UA. Finally, the sample was
dispersed in ethanol and dropped onto the copper grid
for imaging. Dynamic light scattering (DLS) and zeta
potential measurements were performed on a Zetasizer
Nano ZS (Malvern, UK). Optical images of RAW264.7
cells were obtained with a Zeiss Axio Observer Z1
Results and Discussion
Design and Synthesis of HSNs and HRP@HSNs
Typically, HSNs and HRP@HSNs were synthesized via an
ammonia-catalyzed sol-gel process combined with a
water-in-oil (w/o) microemulsion system according to our
previous method [
]. Scheme 1 illustrates the
synthesis of HRP@HSNs. According to TEM images (Fig. 1),
HSNs with and without encapsulated HRP showed an
average diameter of 45 nm (Additional file 1: Figure S1).
UA staining clearly displayed an enhanced electron
density inside the HRP@HSNs, but no staining was observed
outside the HRP@HSNs (Fig. 1b), indicating that the HRP
enzymes were successfully entrapped within the interior
cavity of HRP@HSNs.
The DLS measurements and zeta potential analyses
performed at room temperature are shown in Table 1. DLS
data showed that both HSNs and HRP@HSNs gave
positive zeta potentials in water (pH ~ 6.5) with hydrodynamic
diameters of 188 ± 4 and 184 ± 6 nm in water, respectively.
However, when the nanoparticles were dispersed in
serum-free DMEM, the hydrodynamic diameters
increased to 1767 ± 94 nm for HSNs and 1598 ± 127 nm for
HRP@HSNs. These indicate a small degree of aggregation
of HSNs, but they were still well suspended in media.
Meanwhile, the negative zeta potentials of both
nanoparticles measured in media implied that some of ions and
biomolecules from the biological milieu may have been
adsorbed onto the nanoparticle surfaces [
this condition, the positively charged surfaces of
nanoparticles were covered by negatively charged substances,
which rapidly caused aggregation of the nanoparticles
through electrostatic interactions. To reduce non-specific
aggregation and promote the colloidal stability of
nanoparticles, bovine serum albumin (BSA) was introduced to
the biological media . Subsequently, the hydrodynamic
diameters of HSNs and HRP@HSNs showed considerable
decreased hydrodynamic diameters to 197 ± 43 and 195 ±
19 nm respectively.
HRP Entrapment Efficiency and Loading Capacity of
To investigate the efficiency and loading capacity of
HRP entrapment, fluorescent dye (RITC)-labeled HRP
was prepared (designated RITC-HRP). The fluorescence
intensity of the RITC-HRP@HSNs was measured by
suspending the nanoparticles in 1 M NaOH, and the
amount of encapsulated RITC-HRP was determined
according to a calibration curve established by plotting the
fluorescence intensity versus the concentration of native
RITC-HRP under the same conditions (Additional file 1:
Figure S2). To study the effects of the enzyme
concentration on the entrapment efficiency and loading
capacity, three different amounts of HRP (11.1, 22.2, and 33.
3 nmol) were introduced to the synthesis. It is worth
noting that in this range of concentration, regardless of
how much enzyme was introduced, the entrapment
efficiency of enzymes for each of the three cases was about
6%. This low efficiency may be due to the fact that only
a fraction of the microemulsion droplets nucleated and
grew to HSN; most of the microemulsion droplets were
not nucleated and remained in its small size of ~ 8 nm
]. Future work may be needed for increasing the
loading efficiency. However, the HRP loading capacity of
Scheme 1 Flow chart of the synthesis of horseradish peroxidase-encapsulated hollow silica nanospheres (HRP@HSNs). APTMS,
3aminopropyltrimethoxysilane; TEOS, tetraethyl orthosilicate; SSN, solid silica nanoparticle
HRP@HSNs gradually increased to 12.5 ± 1.2 μg HRP/
mg HSNs when 33.3 nmol of HRP was used
(Additional file 1: Table S1). This indicates that the HRP
loading capacity can be controlled by the amount of
enzyme present in the reaction.
Cytotoxicity and Cellular Uptake of HSNs and HRP@HSNs
To evaluate the in vitro cytotoxicity of HSNs and
HRP@HSNs, cell viability was examined by WST-1
assays. As shown in Additional file 1: Figure S3, no
significant change in RAW264.7 cell proliferation was
observed after treatments of nanoparticles for either 2 h,
or 2 h followed by an additional 24 h of culture. No
obvious effect on the cellular mitochondrial function
caused by the silica nanoparticles was found at the
indicated time points, regardless of the presence or absence
of HRP inside HSNs.
Next, FITC-conjugated HSNs and HRP@HSNs were
respectively prepared to investigate the concentration
effect of nanoparticles on RAW264.7 labeling. Flow
cytometric results (Additional file 1: Figure S4) showed that
RAW264.7 cells were successfully labeled with
FITCHSNs and HRP@FITC-HSNs at different concentrations
for 2 h in serum-free media. In both cases,
dosedependent increases in labeling efficiency were found,
and more than 80% of RAW264.7 cells were labeled by
HSN hollow silica nanospheres, HRP@HSNs horseradish peroxidase-encapsulated HSNs
aAll particles were measured at a concentration of 0.3 mg/mL. Each measurement was repeated at least three times
exposure to nanoparticles at a concentration of > 50 μg/
mL for 2 h. Properties such as high-efficiency
intracellular labeling with a short incubation time, a relatively low
dose of nanoparticles, and non-cytotoxicity make
HRP@HSNs suitable for intracellular detection of ROS.
Reactivity of HRP@HSNs to Various ROS
According to HRP enzyme activity assay using TMB as
substrate, about 40% of the initial enzyme activity
remained upon subsequent encapsulation of HRP into
HSNs. This decrease in the observed specific activity of
the encapsulated enzyme (moles of substrate converted
per unit enzyme per unit time) could have resulted from
mass transfer limitations, that occur when substrates cross
the silica shell toward the HRP [
]. Nevertheless, the
encapsulation strategy provides additional features, for
example, the porous silica shell can protect HRP against
proteolysis while allowing transport of small molecules of
reactants and products [
]. Taken together, the
observed reactivity of HRP@HSNs to ROS, evaluated by
incorporating a fluorescent probe (DHR123), could be
resulted from a combination of the affinity of the
nanoparticles as well as intrinsic property of HRP for ROS.
Cell-free systems were used to generate a variety of
biologically relevant ROS, including hydrogen peroxide
(H2O2), TBHP, hydroxyl radicals (•OH), tert-butoxy
radicals (•OtBu), and superoxide (O2•−). First, DHR123 was
incubated with a panel of ROS in the absence and
presence of HRP or HRP@HSNs followed by measuring
the fluorescence intensity of the product rhodamine 123
(R123). As shown in Fig. 2, regardless of which type of
ROS was employed, the fluorescence intensity was
measured in a time-dependent manner (30, 60, 90, and
120 min). However, apparent differences in intensity
among various ROS depend on the intrinsic properties
of DHR123. On the one hand, in agreement with a
previous study [
], Fig. 2a shows that neither H2O2 nor
O2•− could oxidize DHR123 to R123. Additionally,
DHR123 exhibited higher reactivity for •OtBu and •OH
radicals over other ROS. With the catalytic activity of
HRP, remarkable increases in the fluorescence intensity
were observed in the presence of native HRP and
HRP@HSNs as shown in Fig. 2b, c. It was noted that the
higher fluorescence intensity found in the case of native
HRP compared to HRP@HSNs at the same reaction
time was positively correlated with their observed
To allow a direct comparison between various ROS,
data at a time interval of 60 min were selected and
reported as relative fluorescence intensity normalized to
the control (Additional file 1: Figure S5). Subsequent
analysis of the enhanced intensity ratio was shown by
dividing the relative fluorescence intensity of DHR123 +
HRP or DHR123 + HRP@HSNs by DHR123 (Fig. 2d). In
both HRP-containing cases, a similar trend of the
enhanced intensity ratio over various ROS as well as
significant increases in the reactivity of DHR123 to
H2O2 and O2•− were observed, demonstrating that the
encapsulated HRP gave a high degree of intrinsic
enzyme activity, and the silica shells of HRP@HSNs
allowed transport of small molecules to carry out
Intracellular ROS Detection with HRP@HSNs
To assess the ROS detection functionality of HRP@HSNs
inside cells, RAW264.7 macrophages were incubated with
HRP@HSNs for 2 h followed by washing and then
incubating with DHR123 (20 μM) for 30 min. Subsequently,
cells were washed and treated with PMA (1 μg/mL) for an
additional 1 h. It is known that stimulating macrophages
with PMA results in production of superoxide, which is
dismuted to hydrogen peroxide by superoxide dismutase
or by spontaneous dismutation [
]. Thus, PMA can
function as a stimulant to generate H2O2 in RAW264.7
macrophages to evaluate the intracellular H2O2-sensing
capability of HRP@HSNs. As shown in Fig. 3a, both
cases of RAW264.7 macrophages cultured alone and
cultured with HSNs showed weak fluorescence in the
flow cytometric analysis, indicating that non-stimulated
cells produced a weak basal level of ROS, where no
significant ROS were induced in the presence of HSNs.
In addition, cells treated with HRP@HSNs showed
significant intensity increase (Fig. 3a), suggesting that
the delivered HRP@HSNs gave extra catalytic activity
For stimulation experiments, PMA-treated cells
typically generated more than twofold higher levels of
R123 fluorescence compared to unstimulated cells. In
addition, cells treated with HRP@HSNs had the highest
level of fluorescence, followed by HSNs and then cells
alone. It was noted that treating the stimulated
RAW264.7 macrophages with HSNs resulted in a small
increase in fluorescence intensity compared to that of
the control. This result suggested that cellular stress
responses are triggered extremely rapidly, and sensitive to
external stimuli, including exposure to nanoparticles
]. In addition, both PMA (0.1, 0.25, 0.5, 1, and 2 μg/
mL) and HRP@HSNs (50, 100, and 200 μg/mL) induced
expression of R123 in a dose-dependent manner, as
evident in Fig. 3b, c.
In accordance with the flow cytometric analysis, Fig.
3d displays the representative fluorescence images of
RAW264.7 macrophages stimulated with and without
PMA in the presence and absence of nanoparticles. The
system was capable of visualizing endogenous H2O2
generation in RAW264.7 cells, and the weakest fluorescence
intensity was observed in cells treated with HRP@HSNs
followed by PMA stimulation. As shown in Fig. 4a, the
cell viability of RAW264.7 macrophages in the presence
of the stimulant PMA or exogenous H2O2 was examined
by WST-1 assays. Whereas ROS have been implicated in
], only a small effect on cell viability was
found at the indicated time point, making the following
semi-quantitative analysis practical and meaningful.
Application of HRP@HSNs In Vitro for Quantitative
Analysis of H2O2
To evaluate the capacity of HRP@HSNs for quantifying
endogenous hydrogen peroxide produced in
PMAstimulated RAW264.7 cells, a calibration curve from the
exogenous H2O2 experiment, with a detection range of
0.625~15 μM, was established by microplate
measurements (Fig. 4b, inset). The standard calibration curve
appears to be linear as expected. Then, RAW264.7 cells
were treated with 100 μg/mL of HRP@HSNs for 2 h,
followed by co-incubation with various concentrations
of PMA and 20 μM of DHR123 at 37 °C for 1 h. After
that, the concentration of H2O2 endogenously produced
by PMA-stimulated RAW264.7 cells was determined by
measuring the fluorescence intensity, followed by
conversion using the established calibration curve. Notably,
because most of the HRP@HSNs were uptaken within
the cells, the H2O2-triggered fluorescence of R123 could
be attributed to intracellular enzyme-catalyzed reactions
rather than the extracellular contribution. Although
H2O2 is able to diffuse across biomembranes, due to its
limited diffusion and rapid enzymatic consumption
inside cells, concentration gradients of H2O2 are formed
across membranes [
]. Typically, under normal
physiological conditions, H2O2 has an extracellular
concentration estimated at 10− 7~10− 6 M, which is about
10-fold higher than that observed in intracellular fluid [
]. In pathological conditions, extracellular
concentrations of H2O2 are in the range of 10~50 μM and are
additionally elevated to as high as 10− 4 M in apoptosis . As
shown in Fig. 4b and Additional file 1: Table S2,
endogenous hydrogen peroxide caused by PMA-stimulated
RAW264.7 cells was created in a dose-dependent manner
and produced at levels of about 10 μM when the
concentration of PMA used exceeded 0.25 μg/mL. Taken together,
these results indicate that HRP@HSNs were capable of
detecting semi-quantitatively endogenous the concentration
of hydrogen peroxide of RAW264.7 macrophages under
oxidative stress conditions.
In summary, we have demonstrated that hollow silica
nanospheres encapsulating HRP can be synthesized via a
microemulsion-templating system and act as
intracellular fluorescent ROS sensors. The shells of HRP@HSNs
are permeable to small molecules, such as the enzyme
substrates, which allows them to react with large
enzyme payloads in the hollow cavity. Both the effective
intracellular delivery and satisfactory catalytic activity
of HRP@HSNs significantly enhance
reductiontriggered fluorescence and constitute the ability of
semi-quantitative measurements of endogenous H2O2
in RAW264.7 macrophages under oxidative stress
Because the concentration and location of H2O2 in
eukaryotic cells strongly rely on the types of cells, and
cellular compartments [
], specific targeting of tumor cells
or organelles could further be achieved by surface
modification of HRP@HSNs with monoclonal antibodies or
peptides. Also, non-enzymatic H2O2 detection could be
realized by replacing the interior nanoreactors of HRP
with nanoparticles [
] or boronate-based fluorescent
]. Future efforts should be devoted to
maximizing the sensitivity and specificity for H2O2 as well as
enabling more informative designs of next-generation
nanomaterials. Such hollow capsules could be a promising
platform for modern nanomedicines that aims to
simultaneously image, sensing, and deliver therapeutic
molecules specifically to defective cells.
Additional file 1: Size distribution histograms of HSNs and HRP@HSNs.
Calibration curve of fluorescence intensity versus RITC-HRP. Flow cytometry,
cytotoxicity, and cell proliferation assays. Entrapment efficiency and loading
capacity of HRP@HSNs. (DOC 450 kb)
APTMS: 3-Aminopropyltrimethoxysilane; BSA: Bovine serum albumin;
DHR123: Dihydrorhodamine 123; DLS: Dynamic light scattering;
FITC: Fluorescein isothiocyanate; HRP: Horseradish peroxidase; HSNs: Hollow
silica nanospheres; Igepal CA-520: Polyoxyethylene (5) isooctylphenyl ether;
mETC: Mitochondrial electron transport chain; PET: Positron emission
tomography; PMA: Phorbol 12-myristate 13-acetate; R123: Rhodamine 123;
RITC: Rhodamine B isothiocyanate; ROS: Reactive oxygen species; SSN: Solid
silica nanoparticles; TBHP: Tert-butyl hydroperoxide; TEM: Transmission
electron microscopic; TEOS: Tetraethyl orthosilicate; TMB:
The authors would like to thank Dr. Yann Hung for her encouragement and
many useful comments.
This work was supported by the National Nanotechnology Project NSC
1002120-M-002-001 under Ministry of Science and Technology (MOST), Taiwan,
and in part by grants from Taipei Medical University (TMU 105-AE1-B44 and
HYC performed the experiments, analyzed the results, and wrote the manuscript.
CTC directed this work and contributed to analyzing the results. YPC, FPC, and
FCC participated in the sample fabrication and characterizations. Both SHW and
CYM contributed to the data interpretation, manuscript writing, and supervised
the research. All authors read and approved the final version of the manuscript.
The authors declare that they have no competing interests.
Springer Nature remains neutral with regard to jurisdictional claims in published
maps and institutional affiliations.
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