Selective Inhibition of Human Brain Tumor Cells through

Transcription

Angewandte
Chemie
DOI: 10.1002/anie.200905126
Bionanotechnology
Selective Inhibition of Human Brain Tumor Cells through
Multifunctional Quantum-Dot-Based siRNA Delivery**
Jongjin Jung, Aniruddh Solanki, Kevin A. Memoli, Ken-ichiro Kamei, Hiyun Kim,
Michael A. Drahl, Lawrence J. Williams, Hsian-Rong Tseng, and KiBum Lee*
One of the most promising new chemotherapeutic strategies
is the RNA interference (RNAi)-based approach, wherein
small double-stranded RNA molecules can sequence-specifically inhibit the expression of targeted oncogenes.[1] In
principle, this method has high specificity and broad applicability for chemotherapy. For example, the strategy of small
interfering RNA (siRNA) enables manipulation of key
oncogenes that modulate signaling pathways and thereby
regulate the behavior of malignant tumor cells. To harness the
full potential of this approach, the prime requirements are to
deliver the siRNA molecules with high selectivity and
efficiency into tumor cells and to monitor both siRNA
delivery and the resulting knockdown effects at the single-cell
level. Although several approaches such as polymer- and
nanomaterial-based methods[2] have been attempted, limited
success has been achieved for delivering siRNA into the
target tumor cells. Moreover, these types of approaches
mainly focus on the enhancement of transfection efficiency,
knockdown of non-oncogenes (e.g. the gene coding for green
fluorescent protein (GFP)), and the use of different nanomaterials such as quantum dots (QDs), iron oxide nanoparticles, and gold nanoparticles.[3, 4] Therefore, to narrow the
gap between current nanomaterial-based siRNA delivery and
chemotherapies, there is a clear need to develop methods for
target-oriented delivery of siRNA,[5] for further monitoring
the effects of siRNA-mediated target-gene silencing by means
of molecular imaging probes,[4] and for investigating the
corresponding up/down-regulation of signaling cascades.[6]
Perhaps most importantly, to begin the development of the
necessary treatment modalities, the strategies for nanomate-
rial-based siRNA delivery must be demonstrated on oncogenes involved in cancer pathogenesis.
Herein, we describe the synthesis and target-specific
delivery of multifunctional siRNA–QD constructs for selectively inhibiting the expression of epidermal growth factor
receptor variant III (EGFRvIII) in target human U87 glioblastoma cells, and subsequently monitoring the resulting
down-regulated signaling pathway with high efficiency.[7]
Glioblastoma multiforme (GBM) is the most malignant,
invasive, and difficult-to-treat primary brain tumor. Successful treatment of GBM is rare with a mean survival of only 10–
12 months.[8] EGFRvIII, the key growth factor receptor
triggering cancer cell proliferation in many cancer diseases
such as brain tumors and breast cancer, is a constitutively
active mutant of EGFR which is expressed in only human
GBM and several other malignant cancers, but not in normal
healthy cells (Figure 1 A).[9] We targeted EGFRvIII, since it is
known that knockdown of this gene is one of the most
effective ways to down-regulate the PI3K/Akt signaling
pathway, a key signal cascade for cancer cell proliferation
and apoptosis.[6, 10] Hence by targeting EGFRvIII, our siRNA
delivery strategy based on multifunctional nanoparticles
[*] J. Jung, A. Solanki, K. A. Memoli, H. Kim, M. A. Drahl,
Prof. L. J. Williams, Prof. K.-B. Lee
Department of Chemistry and Chemical Biology, Rutgers University
Piscataway, NJ 08854 (USA)
Fax: (+ 1) 732-445-5312
E-mail: kblee@rutgers.edu
Homepage: http://rutchem.rutgers.edu/ ~ kbleeweb/
Dr. K. Kamei, Prof. H.-R. Tseng
Department of Molecular and Medical Pharmacology
University of California, Los Angeles
Los Angeles, CA 90095 (USA)
[**] We thank V. Starovoytov and Dr. Y. Horibe for helping us with TEM,
and the New Jersey Biomaterial Center (Prof. Kohn) for allowing us
to use the cell culture facilities. K.-B.L. acknowledges the NIH
Directors’ Innovator Award (1DP20D006462-01) and is also grateful
to the NJ commission on Spinal Cord grant (09-3085-SCR-E-0).
Supporting information for this article is available on the WWW
under http://dx.doi.org/10.1002/anie.200905126.
Angew. Chem. Int. Ed. 2010, 49, 103 –107
Figure 1. A) Quantum dots as a multifunctional nanoplatform to
deliver siRNA and to elucidate the EGFRvIII-knockdown effect of PI3K
signaling pathway in U87-EGFRvIII B) Detailed structural information
of multifunctional siRNA–QDs. C) Two different strategies for the
siRNA–QD conjugate. L1 shows the linker for attaching siRNA to QDs
through a disulfide linkage which is easily reduced within the cells to
release the siRNA. L2 shows the linker for covalently conjugating
siRNA to QDs which enables the tracking of siRNA–QDs within the
cells.
2010 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
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Communications
could potentially minimize the side effects caused by conventional chemotherapies, specifically immune suppression,
while significantly improving the efficacy of chemotherapy
against GBM.
We prepared two types of siRNA–QD conjugates, one for
siRNA delivery and the other for siRNA tracking (Figure 1 B,C). Core–shell CdSe/CdS/ZnS QDs with a diameter of
7 nm were synthesized[11] and coated with either trioctylphosphine oxide (TOPO) or hexadecylamine (HDA). In order to
make the QD constructs water-soluble and suitable for
conjugating with siRNA, we displaced these hydrophobic
ligands with a dihydrolipoic acid (DHLA) derivatized with an
amine-terminated poly(ethylene glycol) (PEG) spacer. The
expectation was that the dithiol moiety would provide strong
coordination to the QD surface and increase stability in
aqueous media, the PEG spacer would increase water
solubility and reduce nonspecific binding, and the amine
group would enable conjugation to the siRNA element.[12]
Two bifunctional linkers were synthesized and evaluated for
siRNA conjugation. The linker shown in L1, PTPPf [3-(2pyridyl)-dithiopropionic acid pentafluorophenyl ester], was
designed to release siRNA upon entering the cell by cleavage
of the disulfide linkage, through enzymatic reduction or
ligand exchange (e.g. glutathione).[13] The linker in L2, MPPF
(3-maleimidopropionic acid pentafluorophenyl ester), was
designed to be more robust, thereby enabling evaluation of
cellular uptake and localization of the siRNA construct within
the cellular compartments.[14] Details of the synthesis, characterization and conjugation protocols are given in the
Supporting Information.
The final design component was functionalizing the
construct for tumor-cell-selective transfection. For this purpose two functional peptides, thiol-modified RGD peptide
and thiol-modified HIV-Tat derived peptide, were attached to
the siRNA–QDs by the conjugation methods described
above. Brain tumor cells (U87 and U87-EGFRvIII) overexpress the integrin receptor protein avb3, which strongly
binds to the RGD binding domain.[15] RGD-functionalized
siRNA–QDs selectively accumulate in brain tumor cells in
vitro, and can be tracked by fluorescence microscopy.[16] In
addition, the HIV-Tat peptide enables efficient transfection of
siRNA–QDs in cells when it is directly attached to the QD
surface.[17] The density of siRNA on the QDs and the ratio
between siRNA strands and peptides were optimized for gene
knockdown. It was found that the density of 10 siRNAs per
nanoparticle and the ratio of 1:10 (siRNA for each peptide),
which was in close agreement with literature values,[4] was
optimal for knocking down the target genes (EGFP and
EGFRvIII) overexpressed in our U87 cell lines.
To optimize gene silencing with our siRNA–QD constructs and to assess the transfection efficiency and RNA
interference (RNAi) activity, we examined the suppression of
EGFP expressed in U87 cell lines that were genetically
modified to express EGFP. The cytotoxicity of the constructs
was determined by serial dilution studies. The range of
concentration causing minimal or negligible cytotoxicity was
identified, and subsequent experiments employed the concentrations within this range (see Figure S1 in the Supporting
Information).[18] Importantly, the EGFP cell line has been
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widely used to investigate siRNA-based silencing of EGFP,
since the suppression of EGFP expression does not compromise cell viability. The transfection efficiency of three different kinds of constructs were evaluated; constructs modified
with the RGD peptide only, those modified with the HIV-Tat
peptide only, and those with both HIV-Tat and RGD peptide.
Although the siRNA–QDs modified with only RGD showed
considerable selective internalization within U87-EGFP cells,
siRNA–QDs modified with a combination of RGD and HIVTat peptides (the ratio of siRNA/RGD/HIV-Tat being 1:10:10
per QD) showed maximum internalization within U87-EGFP
cells, in close agreement with previous studies.[4] This optimal
condition was used for subsequent siRNA–QD experiments.
The U87-EGFP cell line was then treated with siRNA–
QDs (siRNA/QDs = 0.12 mm :0.011 mm), modified with HIVTat [ 1.2 mm] and RGD [ 1.2 mm], and simultaneously
imaged using fluorescence microscopy (Figure 2). Cationic
lipids (X-tremeGENE, Roche) were used to further enhance
cellular uptake and prevent degradation of the siRNA within
the endosomal compartment of the cells. The siRNA–QDs
showed significant internalization into the cells. Knockdown
of the EGFP signal was observed after 48–72 h (Figure 2 B).
Fluorescence intensity was influenced by other factors such as
exposure time, media conditions, and cell shrinkage. To
minimize the influence from these external factors, the
Figure 2. Knockdown of EGFP in U87 cells using siRNA–QDs modified
with RGD and HIV-Tat peptides. (Note that yellow arrows mark U87EGFP cells transfected with the siRNA–QDs and the blue arrows
indicate PC-12 cells.) A) Control U87-EGFP cells without siRNA–QDs;
phase-contrast image (A1) and the corresponding fluorescence image
(A2). B) EGFP knockdown using multifunctional siRNA–QDs;
B1) Phase-contrast image shows that the morphology of U87-EGFP
cells has not changed relative to the control cells in (A). B2) Fluorescence image clearly shows the knockdown of EGFP in cells (marked by
yellow arrows) which have internalized the siRNA–QDs (red) after
48 h. C) U87-EGFP control cells (without siRNA–QDs) and U87-EGFP
cells transfected with siRNA–QDs were cocultured so as to investigate
them under the same conditions; C1) Phase-contrast image clearly
shows no difference in the morphology of the U87-EGFP control cells
and the siRNA–QDs transfected cells. C2) Fluorescence image clearly
shows the decrease in the EGFP signal in the U87-EGFP cells transfected with siRNA–QDs as compared to the surrounding U87-EGFP
control cells. D) Phase-contrast image showing the target-oriented
delivery of siRNA–QDs in cocultures of the malignant U87-EGFP cells,
overexpressing the avb3 integrin receptors, and the less tumorigenic
PC-12 cells (blue arrows) incubated with the siRNA–QDs. It can be
clearly seen that most of the siRNA–QDs, owing to the presence of
RGD and HIV-Tat peptides, were taken up by the U87-EGFP cells and
not by the PC-12 cells. Scale bars: 100 mm.
Angew. Chem. Int. Ed. 2010, 49, 103 –107
Angewandte
Chemie
control U87-EGFP cells (without siRNA) were trypsinized
and co-cultured with U87-EGFP cells transfected with
siRNA–QDs in the same well. The U87 cells containing
siRNA–QDs were easily distinguishable from the control
cells owing to the bright fluorescence of the QDs (Figure 2 C2). Cells with internalized siRNA–QDs showed considerable knockdown of the EGFP protein relative to the
surrounding control U87-EGFP cells (Figure 2 C).
To further demonstrate the target-specific delivery of the
siRNA–QDs, we incubated the siRNA–QDs modified with
Tat and RGD against EGFP in co-cultures of the U87-EGFP
cell line with other less-tumorigenic cell lines, such as PC-12
and SK-N-BE(2)C (see Figure S2 in the Supporting Information), which have a considerably small number of integrin
receptors.[19] The presence of RGD tripeptide molecules on
the surface of the siRNA–QDs led to specific binding with
integrin receptors overexpressed in the U87 cells, resulting in
higher cellular uptake by the malignant U87 cells than by the
less tumorigenic PC-12 cells as seen by the selective
accumulation of the QDs within the U87-EGFP cells
(Figure 2 D). These results confirmed our hypothesis that
the target-specific delivery of the siRNA–QDs into brain
cancer cells can be significantly enhanced by functionalizing
the QDs with targeting moieties like RGD tripeptide.
The intracellular delivery of the siRNA–QDs within the
U87-EGFP cells was also confirmed by transmission electron
microscopy (TEM), which clearly shows the presence of QDs
in the cytoplasm of the cells (Figure 3 A). The knockdown
efficiency of the siRNA–QDs was similar to or slightly better
than that of the positive control consisting of U87-EGFP cells
transfected with only siRNA using X-tremeGENE (see
Figure S3 in the Supporting Information). This high transfection efficiency appears to result from synergistic effects of
the two transfection peptides. Decrease in fluorescence
intensities (EGFP signal, green fluorescence) within cells
treated with the above-mentioned systems were then compared with the intensity of U87-EGFP without siRNA. As
shown in (Figure 3 B), the decrease in fluorescence intensity
of U87-EGFP incubated with siRNA–QDs and siRNA alone
was comparable, but drastically lower than that observed for
the control without siRNA. Cells containing siRNA–QDs
show a weaker green fluorescence (EGFP signal) than the
control. This data strongly suggests that siRNA–QDs can be
used simultaneously as delivery and imaging probes.
Having demonstrated the selective manipulation of the
U87-EGFP cell line, we then focused on the knockdown of
EGFRvIII with our siRNA–QD constructs. U87-EGFRvIII
cells were genetically modified to overexpress EGFRvIII, a
mutant-type epidermal growth factor receptor (EGFR) only
expressed within cancer cells.[20] This cell type was incubated
with our siRNA–QDs modified with Tat and RGD peptides
and armed with EGFRvIII-targeting siRNA. The cells were
simultaneously imaged for the internalization of siRNA–QDs
using fluorescence microscopy. Significant cell death was
observed in the wells loaded with siRNA–QDs against
EGFRvIII after 48 h (Figure 4 A). Quantitative analysis
revealed that the number of viable U87-EGFRvIII cells, as
observed by fluorescence microscopy, decreased with increasing incubation time. Relative to the control (U87-EGFRvIII
Angew. Chem. Int. Ed. 2010, 49, 103 –107
Figure 3. Knockdown efficiency of EGFP within U87-EGFP cells and
internalization of multifunctional siRNA–QDs. A) TEM analysis of the
internalization of the multifunctional siRNA–QDs into the U87-EGFP
cells; A1) Presence of multifunctional siRNA–QDs (yellow arrows)
within the cytoplasm and the endosome (scale bar: 5 mm).
A2) Enlarged image showing individual siRNA–QDs within the cytoplasm (scale bar: 2.5 mm). B) The bar graph represents the knockdown
of EGFP over 24 h, 48 h, and 96 h in U87-EGFP cells treated with
siRNA [0.12 mm] only (dark gray), and siRNA–QD [siRNA:QD =
0.12 mm:0.011 mm] (light gray). The EGFP knockdown data was normalized with the expression levels of EGFP in the control U87-EGFP
cells (black).
without siRNA–QDs), there was a significant decrease in the
number of viable cells, thus demonstrating the effectiveness of
our nanoparticle-based siRNA delivery to knockdown the
oncogene. The result was confirmed using the MTT assay
which showed a decrease in the number of viable cells in the
well incubated with siRNA–QDs against EGFRvIII (Figure 4 B). This assay further confirmed that the QDs themselves were noncytotoxic when used alone as they did not
result in any appreciable cell death (see Figure S1 in the
Supporting Information). The knockdown of EGFRvIII and
the inhibition of the downstream proteins in the PI3K
signaling pathway were confirmed using Western immunoblotting. The results (Figure 4 C) confirm a considerable
decrease in the expression of EGFRvIII, and down-regulation
of phospho-Akt and phospho-S6 relative to the control. Thus,
these results demonstrate the specificity of the siRNA against
EGFRvIII, the inherent noncytotoxicity of the QDs, and the
facile evaluation and manipulation of cancer cell proliferation
with multifunctional QD constructs.
In summary, we have demonstrated an application of
multifunctional siRNA–QDs focusing on targeted delivery,
high transfection efficiency, and multimodal imaging/track-
www.angewandte.org
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Communications
egies but also for dissecting signaling cascades triggered by
inhibiting specific proteins. Collectively, our strategy for
siRNA delivery based on multifunctional QDs has significant
potential for simultaneous prognosis, diagnosis, and therapy.
Received: September 13, 2009
Published online: November 30, 2009
.
Keywords: antitumor agents · gene knockdown · nanoparticles ·
nonviral siRNA delivery · target-specific delivery
Figure 4. Knockdown of EGFRvIII in U87-EGFRvIII using multifunctional siRNA–QDs. A) Phase-contrast images showing the internalization of siRNA–QDs into the U87-EGFRvIII cells. A1) Morphology of
U87-EGFRvIII cells before incubation with siRNA–QDs on Day 0.
A2) U87-EGFRvIII cells after incubation with siRNA–QDs (red) on
Day 0. A3) Morphology of U87-EGFRvIII cells 48 h after incubation
with siRNA–QDs. Note that effect of the EGFRvIII knockdown by the
siRNA–QDs can be clearly seen as the cells have clearly shrunk (yellow
arrows) and appear to have collapsed (cf. Day 0), marking the onset of
apoptosis; scale bar: 100 mm. B) Cell viability assay using MTT assay.
B1) Optical image of cell viability (MTT) assay in a well plate. Dark
blue color indicates a high number of viable cells and pale blue
indicates a low population of viable cells. B2) MTT-assayed wells were
quantified with UV absorbance and the data was converted to cell
viability data. Untreated control C1 and C2 represent control cell
population and viable cell population, respectively, in the presence of a
cationic lipid based transfection reagent. siRNA–QD-transfected cells
in experiment E1 and siRNA treated cells in E2 show low numbers of
the viable cells due to knockdown of EGFRvIII gene. C) Western
immunoblotting shows the silencing effect of the EGFRvIII gene.
Protein expression level of EGFRvIII is dramatically decreased, and
phosphorylation levels of key proteins in PI3K signaling pathway are
reduced significantly. The upstream protein (AKT) and the downstream
protein (S6), which play an important role in cell proliferation, are
selected to investigate the gene-knockdown effect on the PI3K signaling pathway.
ing. Our siRNA–QDs could be used for the development of
novel chemotherapies and diagnostics relevant to brain
cancer research. These novel methods and applications
complement recent advances in nanomaterial-based siRNA
delivery, nanomaterial-based molecular imaging, and siRNAbased chemotherapeutic strategies reported recently. While
the ability to functionalize as well as control the surface of
quantum dots with specific linkers and multifunctional
molecules (siRNA and peptides) is critical for nanoparticlebased drug delivery, this method could also provide highly
useful information regarding biological surface chemistry of
nanomaterials. In addition, the application of multifunctional
siRNA–QDs to modulate the key cancer signaling pathways
is important not only for selective chemotherapeutic strat-
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PAPER
www.rsc.org/loc | Lab on a Chip
An integrated microfluidic culture device for quantitative
analysis of human embryonic stem cells†
Ken-ichiro Kamei,‡ab Shuling Guo,‡cd Zeta Tak For Yu,‡abe Hiroko Takahashi,ab Eric Gschweng,cd
Carol Suh,ab Xiaopu Wang,ab Jinghua Tang,d Jami McLaughlin,c Owen N. Witte,*acdg
Ki-Bum Lee*abf and Hsian-Rong Tseng*ab
Received 29th May 2008, Accepted 14th October 2008
First published as an Advance Article on the web 20th November 2008
DOI: 10.1039/b809105f
We have successfully designed and fabricated an integrated microfluidic platform, the hESC-mChip,
which is capable of reproducible and quantitative culture and analysis of individual hESC colonies in
a semi-automated fashion. In this device, a serpentine microchannel allows pre-screening of dissociated
hESC clusters, and six individually addressable cell culture chambers enable parallel hESC culture, as
well as multiparameter analyses in sequence. In order to quantitatively monitor hESC proliferation and
pluripotency status in real time, knock-in hESC lines with EGFP driven by the endogenous OCT4
promoter were constructed. On-chip immunoassays of several pluripotency markers were carried out to
confirm that the hESC colonies maintained their pluripotency. For the first time, our studies
demonstrated well characterized hESC culture and analysis in a microfluidic setting, as well as a proofof-concept demonstration of parallel/multiparameter/real-time/automated examination of self-renewal
and differentiation in the same device.
Introduction
Human embryonic stem cells (hESCs),1–3 derived from the inner
cell mass of blastocyst-stage embryos, hold great potential for the
treatment of many devastating diseases and injuries. This is
mainly due to two distinct properties: (i) they can self-renew
indefinitely and (ii) they can potentially generate all cell types in
the human body. Intrinsic regulators (e.g., growth factors and
signaling molecules) and cellular microenvironments (e.g.,
extracellular matrices, ECMs) play critical roles in the regulation
of self-renewal and differentiation of hESCs. Conventionally,
hESCs are passaged in clusters (containing approximately
a
Department of Molecular & Medical Pharmacology, University
of California, Los Angeles, CA 90095, USA. E-mail: hrtseng@mednet.
ucla.edu
b
Crump Institute for Molecular Imaging, David Geffen School of Medicine,
University of California, Los Angeles, CA 90095, USA
c
Department of Microbiology, Immunology and Molecular Genetics,
d
The Eli and Edythe Broad Center of Regenerative Medicine and Stem Cell
Research, University of California, Los Angeles, CA 90095, USA. E-mail:
owenw@microbio.ucla.edu
e
Department of Mechanical and Aerospace Engineering, University of
California, Los Angeles, CA 90095, USA
f
Department of Chemistry & Chemical Biology, Institute for Advanced
Materials, Devices and Nanotechnology, The Rutgers Stem Cell
Research Center, Rutgers, The State University of New Jersey,
Piscataway, NJ 08854, USA. E-mail: kblee@rci.rutgers.edu
g
The Howard Hughes Medical Institute, David Geffen School of Medicine,
† Electronic supplementary information (ESI) available: Fabrication of
hESC-mChips, generation of genetically modified hESCs (i.e.,
HSF1-LG, HSF1-OCT4-EGFP and H1-OCT4-EGFP), microscopy
settings and induction of hESC differentiation. See DOI:
10.1039/b809105f
‡ These three authors contributed equally to this work.
This journal is ª The Royal Society of Chemistry 2009
20–200 cells) using well plates or culture dishes. Growth-arrested
mouse embryonic fibroblast (mEF) feeder layers are co-cultured
in serum replacement-containing medium to supply the essential
intrinsic regulators and environmental cues. However, there have
been concerns associated with xenogenic contamination that
would restrict potential therapeutic applications of hESCs in
clinical settings.4,5 In order to harness the unique potential of
hESCs and to improve self-renewal6–12 and controlled differentiation of hESCs,13–16 systematic approaches have been adopted
to screen a broad range of serum- and feeder-free culture
conditions to obtain a better understanding of the roles of
intrinsic regulators and cellular microenvironments. The cost to
perform these screening experiments is high, since they consume
a considerable quantity of hESCs, ECM materials and culture
media containing expensive growth factors. There is a clear need
for developing a miniaturized platform on which to carry out
large-scale screening in a cost-efficient fashion.
There is a growing interest to develop microfluidics-based
technologies17 for performing cell culture and analysis. Microfluidic systems offer intrinsic advantages over conventional
macroscopic culture such as reduced sample/reagent consumption and precise control over the delivery of culture fluids
and soluble factors. A continuous-flow microfluidic system
composed of the simplest device configuration (i.e., individual
microchannels and the respective inlets/outlets) has been utilized
to implement miniaturized cell culture and analysis.18 In this
case, bovine adrenal capillary endothelial cells were seeded in
protein-coated microchannels, where culture media and assay
reagents were introduced and withdrawn from the microchannels
through inlets and outlets, respectively. Several challenges
remain, however, to explore these continuous-flow cell culture/
assay chips for systematic screenings where combinations of
multiple parameters should be tested to obtain the desired
Lab Chip, 2009, 9, 555–563 | 555
outcomes. For example, when many cell culture conditions are
screened in a microchannel network, it is inevitable that the
individual conditions would be cross-contaminated through
diffusion. Moreover, this multiparameter screening necessitates
a delicate microfluidic delivery/mixing system for handling small
amounts of culture components coordinated by an automated
operation system. To overcome these challenges, different
miniaturized functional modules, including isolation valves
and mechanical pumps, have been developed to prevent cross
contamination and to attain precise fluidic delivery and mixing.
Most importantly, these miniaturized valves and pumps can
be digitally controlled, thus allowing automated cell culture in
a microfluidic chip.19
Among the exciting automated microfluidic systems, the
poly(dimethylsiloxane) (PDMS)-based integrated microfluidic
system represents a large-scale architecture of microchannel
networks that enables the execution of sequential and parallel
processes in individual devices.20 Particularly, the biocompatible
and gas-permeable properties of PDMS matrices help to retain
proper physiological conditions for a wide range of mammalian
cells suitable for different screening applications. The cooperation of integrated hydraulic valves confines distinct regions for
testing specific screening combinations/conditions without the
concern of cross contamination,21,22 and a peristaltic pump
(composed of three consecutive isolation valves) is capable of
delivering, metering, and mixing of nanoliter (nL)-level fluids
with great precision.22 Over the past seven years, different
PDMS-based integrated microfluidic devices have been developed for complicated chemical23,24 and biological operations,25–27
including recent demonstrations on culturing human mesenchymal stem cells.28 Obviously, the characteristics of the
PDMS-based integrated microfluidic system meet the needs of
conducting systematic screenings of optimal hESC culture
conditions. Although there are examples of the culture of human
neural stem cells29,30 and mouse embryonic stem cells31 in
different microfluidic systems, there are few reports to demonstrate the culture and manipulation of hESCs in a microfluidic
platform.32–35
Here, we demonstrate an integrated microfluidic platform
(hESC-microChip, hESC-mChip), which allows reproducible and
quantitative culture and analysis of individual hESC colonies in
a semi-automated fashion. Initially, several challenges were
envisioned to conduct this study in a hESC-mChip. For example,
hESCs are extremely sensitive to changes of intrinsic regulators,
cellular microenvironments and ambient pressure/temperature.
The effects of culturing hESCs into a hESC-mChip on hESCs
should be addressed in these contexts. Further, hESCs have to be
passaged in clusters and co-cultured in the presence of growtharrested mEF feeder layers. Experience in handling hESC clusters in the chip and co-culturing of hESC clusters with the
adherent mEF cells should be acquired. Moreover, to confirm
that the chip-cultured hESC colonies maintain their pluripotency
over a certain culture period, immunoassays for a number of
pluripotency markers have to be carried out in sequence. Each
immunoassay for chip-based operation will be optimized and
some of them will be compiled in sequences. The goal of our
study was not meant to unveil novel insights in hESC biology or
develop a new type of microfluidic technology, but to acquire
solid experience and practical knowledge of performing
556 | Lab Chip, 2009, 9, 555–563
microfluidic hESC culture, which will constitute a useful
foundation for exploring further application of microfluidic
platforms in hESC research.
Experimental
hESC culture in a hESC-mChip
All hESC research described here was approved by the UCLA
Embryonic Stem Cell Research Oversight Committee. A newly
fabricated hESC-mChip was sterilized under UV light for 15 min
prior to on-chip hESC culture. Based on a two-layer coating
approach, a bovine fibronectin solution (FN, 1 mg mL1 in PBS,
Sigma) and a gelatin solution (0.2% in PBS) were sequentially
introduced into the hESC-mChip from ‘‘Inlet 2’’ using Teflon
tubing (Fig. S1†). g-Irradiated mEFs (1 107 cells mL1) were
loaded into the cell culture chambers from ‘‘Inlet 3’’. mEFs were
cultured for 12 hr in a humidified incubator (37 C, 5% CO2,
Thermo Fisher Scientific) before loading cells. hESCs cultured in
a 6-well plate were passaged with 1 mg mL1 of collagenase IV in
DMEM/F12 (see the ESI†). The freshly dissociated hESC clusters were introduced into the cell culture chambers through ‘‘Inlet
1’’ connected to a serpentine microchannel, where every hESC
colony was visually inspected (Fig. 1c). Gravity flow36 was
adopted in order to introduce hESC clusters into each cell culture
chamber. To ensure the quality and uniformity of hESC colonies
in our studies, only hESC clusters with desired sizes (100 20 mm) and disc-shaped morphologies were selected for seeding.
In general, four to six hESC colonies were accommodated in each
cell culture chamber. The locations of individual hESC colonies
were registered according to the ruler, allowing continuous fate
mapping by an inverted microscope. The hESC-mChip-based
hESC culture was carried out in a humidified incubator (37 C,
5% CO2). By programming the cooperation of isolation valves
and peristaltic pumps, media stored in Teflon tubing was
introduced into each cell culture chamber every 12 hr.
Immunocytochemistry and histology
hESC colonies were fixed by introducing paraformaldehyde (4%,
Electron Microscope Science) into the cell culture chambers
in the hESC-mChip. After permeabilization with Triton
X-100 (0.5%, Fluka) in PBS for 30 min, a blocking solution
containing normal goat serum (5%, Vector Laboratory), normal
donkey serum (5%, Jackson Laboratory), bovine serum albumin
(3%, Fraction V, Sigma) and N-dodecyl-b-D-maltoside (0.1%,
Pierce)37 was loaded into the device from ‘‘Inlet 1’’, and the device
was incubated at room temperature for 1 hr. After rinsing with
PBS containing 0.1% Tween 20 (PBS-T), the hESC colonies were
incubated with human specific antibodies for OCT4 (2 mg mL1,
mouse monoclonal IgG, Santa Cruz Biotechnology), NANOG
(2 mg mL1, rabbit polyclonal IgG, Abcam), SSEA1 (2 mg mL1,
mouse monoclonal IgM, Santa Cruz Biotechnology), SSEA4
(2 mg mL1, mouse monoclonal IgG, Santa Cruz Biotechnology),
TRA-1-60 (2 mg mL1, mouse monoclonal IgM, Santa Cruz
Biotechnology) or TRA-1-81 (2 mg mL1, mouse monoclonal
IgM, Santa Cruz Biotechnology) for 24 hr at 4 C. After rinsing
the cell culture chambers with a blocking solution, the respective
secondary antibody: Alexa Fluor 514-conjugated goat antimouse IgG (H + L) (10 mg mL1, Invitrogen), R-Phycoerythrin
(R-PE)-conjugated goat anti-mouse IgM (10 mg mL1, BD
Pharmigen), Cy5-conjugated goat anti-rabbit IgG (H + L)
(7.5 mg mL1, Jackson ImmunoResearch), or Alexa Fluor 750conjugated goat anti-mouse IgG (H + L) (20 mg mL1, Invitrogen) was loaded into the cell culture chambers to detect the
bound primary antibodies. After incubating at room temperature for 1 hr, the chambers were rinsed with PBS-T. Finally,
10 mg mL1 of DAPI solution was loaded for nuclear staining.
For alkaline phosphatase (AP) staining, the hESC colonies were
fixed with paraformaldehyde (4%) for 30 min at room temperature. After fixation, a freshly prepared AP staining solution (1 mg
mL1 Fast Red TR salt in water with 0.01% AS-MX alkaline
phosphate solution, Sigma) was loaded into the cell culture
chambers and incubated for 30 min in the dark. Fluorescence and
phase contrast images were taken with an inverted microscope
(TE2000S, Nikon), and quantitatively analyzed with MetaMorph
software (version 7.1.3.0; Molecular Devices) (Fig. S2†).
Results
Design and operation of hESC-mChips
Fig. 1 Design of the hESC-mChip. (a) Schematic illustration of a hESCmChip capable of semi-automated operation for hESC culture and analysis. The functions of different hydraulic valves are illustrated by their
colors: Red for pneumatic valve operation and yellow for fluidic delivery
and metering. The 6 1 array of cell culture chambers (with dimensions of
3000 mm (l) 500 mm (w) 100 mm (h) and total volume of 150 nl) are
numbered 1 to 6. Each cell culture chamber is separated by hydraulic
valves to achieve individual addressability. There are four inlets and two
outlet channels in each device, providing accesses to hESC colonies,
culture media and immunostaining reagents. (b) Optical micrograph of
the actual device. Food dyes were introduced into the various microchannels to help visualize the functional components of the hESC-mChip:
Red and yellow as illustrated in (a); blue indicates the fluidic channels. A
ruler was fabricated alongside of each cell culture chamber to serve as
a landmark that directs continuous fate mapping of individual hESC
colonies by an optical microscope. For hESC culture in the hESC-mChip,
freshly prepared hESC clusters were introduced into cell culture chambers
through the inlet connected to a serpentine microchannel as shown in (c),
where every hESC cluster was visually inspected (i,iv). To ensure the
uniformity of hESC clusters used in our studies, only hESC clusters with
the desired size and morphology were introduced into cell culture chambers (i–iii). Undesirable hESC clusters were removed as waste (iv–vi).
A typical hESC-mChip (Fig. 1a and b) is composed of a 6 1
array of cell culture chambers (with dimensions of 3000 mm (l) 500 mm (w) 100 mm (h) and total volume of 150 nL) for
accommodating individual hESC colonies. A ruler was fabricated alongside each cell culture chamber as a landmark, so that
individual hESC colonies were registered for continuous fate
mapping using an inverted microscope. There are four inlets and
two outlet channels in each device, providing accesses to culture
media and immunostaining reagents. For hESC culture in the
hESC-mChip, freshly dissociated hESC clusters (obtained by
digesting conventionally cultured hESC colonies with collagenase IV) were introduced into cell culture chambers through the
inlet via a serpentine microchannel where every hESC cluster
was visually inspected (Fig. 1c). To ensure the uniformity of
hESC clusters in our studies, only disc-shaped clusters with
diameters within 100 20 mm were introduced to the cell culture
chamber. In general, four to six hESC clusters were selected and
seeded per chamber. To allow parallel examination of multiple
variables over time, six pairs of hydraulic valves (Fig. 1a and b)
conferred individual addressability to the six cell culture
chambers in the device. A laptop computer was utilized to
control the valves and pumps to achieve automated operation of
the hESC-mChip.
To ensure general applicability of the hESC-mChips, we conducted our studies using a collection of hESC lines, including two
parental hESC lines (i.e., HSF1 and H1) and three genetically
modified hESC lines–(i) HSF1-LG which expresses firefly luciferase and enhanced green fluorescent protein (EGFP) as a fusion
protein driven by the ubiquitin promoter, (ii) HSF1-OCT4EGFP and (iii) H1-OCT4-EGFP which express EGFP under the
endogenous OCT4 promoter. In our proof-of-concept studies,
hESC-mChip-based culture experiments were carried out in the
presence of g-irradiated mEFs, using serum replacement-containing media with either 10 or 100 ng mL1 of bFGF. The
g-irradiated mEFs were seeded in the protein-coated cell culture
chambers for 12 hr prior to the introduction of the dissociated
hESC clusters. Throughout the experiment, hESC-mChips
Lab Chip, 2009, 9, 555–563 | 557
were stored in a humidified incubator (5% CO2, 37 C). The
gas-permeability of PDMS allowed rapid gas exchange between
the atmosphere around the hESC-mChips and the media in the
cell culture chambers. The results revealed that medium with
a concentration of 100 ng mL1 bFGF gave better reproducibility of hESC self renewal in the device. Due to the higher
surface area-to-volume ratio of the microfluidic environment,
a significant amount of bFGF was absorbed on the microchannels surfaces. The use of a higher concentration of bFGF
was sufficient to maintain the chip-cultured hESC colonies. Since
the hESC-mChip consumes only 150 nL of medium in each
culture chamber, the use of 100 ng mL1 bFGF has very limited
impact on experimental cost.
Optimization of hESC culture conditions
Since this digitally controlled hESC-mChip is capable of smallscale screening, we were able to utilize these devices to progressively define an optimal surface coating protocol and a cell
feeding schedule which are optimized for the hESC colonies.
Initially, several protein coating combinations and approaches
were examined in the device in search of a recipe (Fig. S3†) which
led to efficient plating of the g-irradiated mEF layer and
reproducible self-renewal of hESC colonies. We identified
a layer-by-layer coating method: a layer of fibronectin (FN) was
first coated onto the PDMS surfaces (by introducing 1 mg mL1
FN solution into the cell culture chambers and incubated at 37
C for 30 min), followed by sequential deposition of a gelatin
layer (0.2% gelatin solution at 37 C for 15 hr). This coating
method resulted in a uniform and long-lasting FN/gelatin layer
on the PDMS surface for maintaining hESC colonies. Using
a hESC-mChip with six FN/gelatin-coated cell culture chambers,
we then carried out a parallel examination of different cell
feeding schedules. By programming the cooperation of hydraulic
valves and peristaltic pumps, the medium stored in Teflon tubing
was periodically introduced into each cell culture chamber at
different feeding intervals (i.e., 3, 6, 12, 18, 24 and 36 hr). As
a result of monitoring morphology and survival rate of hESC
colonies, we identified a 12-hr feeding cycle which allowed the
reproducible self renewal of hESC colonies in our hESC-mChip
for 6 days. By using the optimized hESC culture condition
(i.e., in the presence of serum replacement-containing medium,
g-irradiated mEFs and FN/gelatin coated cell culture chambers,
as well as using a cell feeding cycle of 12 hr), we were able to
culture HSF1, H1, HSF1-LG, HSF1-OCT4-EGFP and H1OCT4-EGFP in the hESC-mChips for 6 days (Fig. S4†). In
addition, HSF1 cells could be cultured in our mChips up to
12 days for the longest culturing periods (Fig. S5†). By chance,
a single hESC (HSF1) colony was cultured in a cell culture
chamber (Fig. S6†). There was no significant difference observed
in contrast with the multi-colonies culture.
Chip-based immunocytochemistry to confirm hESC pluripotency
To confirm pluripotency of hESC-mChip-cultured hESCs,
immunocytochemistry for a number of pluripotency markers,
including alkaline phosphatase (AP), stage-specific embryonic
antigen 4 (SSEA4), OCT4 (also known as POU5F1), NANOG,
tumor-related antigen (TRA)-1-60 and TRA-1-81, was carried
558 | Lab Chip, 2009, 9, 555–563
out in the same device. The digitally controlled interface allowed
automated execution (Supplementary Methods) of the immunostaining processes, where multiple reagents, including paraformaldehyde (4%) for fixation, Triton X-100 (0.5%) in PBS for
permeabilization of the cell membrane, and antibodies for fluorescent immunocytochemical analyses, were introduced into the
cell culture chambers in sequence. It is noteworthy that mixtures
of four different pluripotency markers could be introduced in
individual culture chambers, allowing four fluorescence immunocytochemical analyses at the same time. Finally, the resulting
hESC-mChip was mounted on either a fluorescence microscope
or a confocal microscope to collect immunofluorescence micrographs. Fig. 2a and b show immunofluorescence images of
hESC-mChip-cultured HSF1 and H1 colonies, respectively.
These cells retained characteristic hESC morphology, and strong
fluorescence signals of pluripotency markers, indicating that they
maintained their stemness over the six-day culture period. Threedimensional (3D) confocal micrographs of hESC-mChipcultured hESCs (Fig. 2c–e, and the visualization of its 3-D
structure in a movie clip in Supplemental Information) revealed
3D structures of the hESC colonies, and merged 3D confocal
micrographs indicate the co-localization of different pluripotency markers.
Quantification of hESC growth in the hESC-mChip
To monitor hESCs in vitro and in vivo, HSF1-LG cells were
generated by infecting HSF1 cells with lentivirus containing
a mutated thermostable firefly luciferase (mtfl)38 and EGFP as
a fusion protein (LG) driven by a ubiquitin promoter (Fig. S7a–
h†). The EGFP signal allows the quantification of cell growth in
real time.39 To test this idea, freshly dissociated HSF1-LG clusters were cultured in the hESC-mChip for 6 days,40 and their
EGFP signals were measured every other day (Fig. 3a). In
parallel, these dissociated clusters were cultured in conventional
culture dishes under similar conditions. The growth rates of
hESCs were quantified by measuring the increased surface area
or integrated EGFP intensities of individual hESC colonies at
different time points. As shown in Fig. 3b and c, both quantification approaches gave similar results. Although inhibition of
cell proliferation has been reported in other microfluidic cell
culture settings,41 possibly due to the constrained accumulation
of soluble factors in the diffusion dominant microfluidic environment, the growth rates of hESC-mChip cultured colonies were
not significantly different from those observed for hESCs in
conventional dishes (p ¼ 0.21).
Multiparameter monitoring of hESC pluripotency status
In order to monitor the pluripotent status of hESCs in realtime, we constructed OCT4-EGFP knock-in reporter lines in
HSF1 and H1 cells (i.e., HSF1-OCT4-EGFP and H1-OCT4EGFP) (Fig. S8a–d†). In both cases, the linearized OCT4EGFP knock-in construct42 was introduced into hESCs via
Nucleofector (Amaxa Biosystems). These genetically modified
hESCs could be passaged as their parental HSF1 or H1 cells.
To ensure that EGFP expression in these hESCs faithfully
represents pluripotency, both HSF1-OCT4-EGFP and H1OCT4-EGFP were induced to differentiate in the presence of
Fig. 2 On-chip immunocytochemistry to confirm hESC pluripotency. Bright-field and fluorescence micrographs of hESC-mChip-cultured hESC
colonies stained with a collection of pluripotency markers: (a) Three HSF1 colonies were stained by DAPI and alkaline phosphatase (AP), as well as
immunostained for OCT4, NANOG, TRA-1-60 and TRA-1-81. (b) Two H1 colonies were stained with DAPI, SSEA4, NANOG, TRA-1-60 and
TRA-1-81. The characteristic morphologies and strong fluorescent signals of pluripotency markers indicate that the hESCs cultured in hESC-mChips
retained their pluripotency over the six-day culture period. (c–f) Three-dimensional (3D) confocal micrographs of a genetically modified hESC colony
(HSF1-LG). (c) DAPI nuclear staining, (d) EGFP expression, (e) OCT4 immunostaining and (f) the merged image. These images revealed information
on the 3D structure of the hESC colonies.
fetal bovine serum (FBS, 15%) and the absence of mEFs. After
about 10 days in culture, over 90% of the cells lost EGFP
expression, correlating with their differentiated morphology
(Fig. S8e†). Additionally, if the EGFP signal truly correlates
with the endogenous OCT4 expression, this marker could be
used to rescue pluripotent cells from a differentiated population. To show this, OCT-EGFP-knock-in cells were differentiated as embryoid bodies in serum containing medium. After
21 days, the EGFP positive population (approximately 3%) was
sorted from the non-expressing cells (Fig. S8g†) and re-plated
into conventional culture conditions. Indeed, these cells re-grew
into typical ES colonies and maintained pluripotency markers
(Fig. S8h–j†).
Either HSF1-OCT4-EGFP or H1-OCT4-EGFP hESCs were
utilized for the demonstration of parallel examinations of
controlled differentiation and proliferation in individual hESCmChips. In a given study, differentiation of hESCs was carried
out in cell culture chambers No. 1, 3 and 5, where only a layer of
Lab Chip, 2009, 9, 555–563 | 559
Fig. 3 Real-time quantitative monitoring of growth of hESC-mChip-cultured hESC colonies. (a) Fate mapping of hESCs cultured in a hESC-mChip
with bright-field microscopy. As we show in Fig. 2, the hESC colonies still had pluripotency, even in hESC colonies attached onto the channel. And,
since there are PDMS walls in the EGFP images of HSF1-LG at day 6, there is no EGFP signal from those areas. In addition, since EGFP expression in
HSF1-LG is under the regulation of a ubiquitin promoter which constitutively active in any kinds of cells, EGFP intensity doesn’t reflect their pluripotency. (b) Quantitative comparison of growth rate of the size of hESC colonies in conventional culture dishes and hESC-mChips. (c) Quantitative
comparison of growth rate of EGFP intensity of hESC colonies in conventional culture dishes and hESC-mChips. Each bar represents the standard
deviation (n > 7).
FN was coated and no feeder cells were applied. In parallel,
proliferation of hESCs was carried out in culture chambers No.
2, 4 and 6, where FN/gelatin coating was applied and g-irradiated mEFs were cultured. After 24 hr, the genetically modified
hESCs clusters were introduced into the 6 cell culture chambers.
After 3 hr, differentiation medium (containing 5 mM retinoic acid
(RA) and 15% FBS) and self-renewal medium were separately
introduced into the respective sets of chambers with a 12-hr
feeding schedule. The EGFP signals in the differentiating or selfrenewing cells were monitored every other day to record the
status of their pluripotency. hESC colonies in differentiation
medium gradually lost their compact morphologies and spread
560 | Lab Chip, 2009, 9, 555–563
out. Concurrently, the EGFP signals started to diminish after
2 days, whereas the hESC colonies in the self-renewal medium
grew larger accompanied by increased EGFP signal (Fig. 4a).
After 4 days of culture in a hESC-mChip, immunocytochemistry
for SSEA1 was performed to confirm differentiated or pluripotent status. In general, SSEA1 is the marker for pluripotency for
murine ESCs, whereas only differentiated hESCs show expression.42 As shown in Fig. 4b and 4c, hESCs in differentiation
medium showed strong staining for SSEA1, correlating with the
loss of EGFP signal. In contrast, hESCs in the self-renewal
medium maintained strong expression of EGFP, but no expression of SSEA1 was detected. This demonstrated that a single
Fig. 4 A single hESC-mChip serves as a platform for parallel examination of controlled differentiation and self-renewal for hESCs. Either differentiation medium (containing 5 mM retinoic acid and 15% FBS) or self-renewal medium (100 ng mL1 bFGF) was introduced into cell culture chambers
No. 1/3/5 or No. 2/4/6, respectively. (a) Fate mapping of HSF1-OCT4-EGFP colonies under the differentiation and self-renewal conditions using an
inverted fluorescent microscope. After 2 days, hESC colonies under the differentiation condition gradually lost their hESC morphology and EGFP
signal, whereas the hESC colonies under the self-renewal condition grew larger accompanied by an increased EGFP signal. (b) HSF1-OCT4-EGFP cells
were immunostained for SSEA1 (a differentiation marker) at Day 4. (c) Quantitative comparison of EGFP intensity of hESC colonies in differentiation
medium and self-renewal medium. Each bar represents the standard deviation (n > 7).
hESC-mChip could carry out controlled self-renewal and differentiation in parallel without cross contamination.
Discussion
We have successfully demonstrated reproducible and quantitative culture and analysis of individual hESC colonies in an
integrated microfluidic platform, the hESC-mChip. The six
individually addressable cell culture chambers in the hESCThis journal is ª The Royal Society of Chemistry 2009
mChip allowed parallel examination of combinations of variables
over time to obtain optimal culture conditions for self-renewal
and controlled differentiation of hESCs. In addition to the
intrinsic advantages of microfluidic systems, the hESC-mChip
provides an opportunity to culture hESCs in different conditions
in parallel as well as to run sequential phenotypical and functional analyses. Several small-scale screenings were performed to
identify the optimal chip-based culture conditions that are widely
applicable for a collection of hESCs, including two parental
Lab Chip, 2009, 9, 555–563 | 561
hESC lines and three genetically modified hESC lines. Semiautomated immunoassays for a number of pluripotency markers
were carried out in sequence to confirm that the chip-cultured
hESC colonies maintained their pluripotency over a culture
period of at least 6 days.43 Two more hESC lines, HSF6 and H9,
could also be cultured in the hESC-mChip, and maintained their
pluripotency (data not shown). Finally, we were able to
demonstrate parallel examination of proliferation or controlled
differentiation in a single hESC-mChip. Three genetically modified hESC lines allowed quantitative monitoring of hESC
proliferation and pluripotency of the hESC-mChip-cultured
hESC colonies in a real-time manner.
Conventional hESC research is conducted in a collective
fashion which overlooks a great deal of information on individual hESC colonies and their microenvironments over time.
Lack of precise control of experimental and analytical conditions
makes it difficult to interpret the results obtained from different
experiments. In the hESC-mChip, there are six identical cell
culture chambers providing a closely related microenvironment
for multiparameter analysis. In each cell culture chamber, there is
a built-in landmark to register individual hESC colonies for
continuous fate mapping. The hESC-mChip is controlled by
a laptop PC, allowing reproducible culture and analysis of
individual hESC colonies in a semi-automated fashion. Although
there were microfluidic devices developed for the culture of
hESCs,33,34,44 no quantitative and integrated culture and analysis
has been reported. In conjunction with a fluorescent microscope
and three genetically modified hESC lines, we demonstrated, for
the first time, that the hESC-mChip is capable of integrated and
quantitative culture and analysis of hESCs.
We also realized that we have a limited number of samples
per chip in the hESC-mChip, and it cannot be operated in a fully
automated fashion. Currently, a new generation of fully automated hESC-mChip incorporating hundreds of individual cell
culture chambers is under development. We envision that the
new generation hESC-mChip will be applied for high-throughput
screening of feeder-free and chemically defined conditions
which better regulate self-renewal and differentiation of hESCs.
Furthermore, by using HSF1-LG and OCT4-EGFP knock-in
cell lines, the integrated microfluidic hESC culture platform can
provide a new screening system of the condition for single hESC
expansion and fate mapping for individual hESCs.
Acknowledgements
This work was partially supported by the Eli and Edythe Broad
Center of Regenerative Medicine and Stem Cell Research at the
Institute of Molecular Medicine at University of California, Los
Angeles and the DOE-UCLA Institute of Molecular Medicine.
O.N.W. is an investigator of the Howard Hughes Medical Institute. We thank Dr. Michael Teitell at UCLA for his expert opinions
on teratoma histology. We thank Dr. James Thomson at WiCell
for kindly providing the OCT4-EGFP knock-in DNA construct.
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Lab Chip, 2009, 9, 555–563 | 563
Cell Stem Cell
Resource
Phosphoproteomic Analysis
of Human Embryonic Stem Cells
Laurence M. Brill,1,2,5,* Wen Xiong,3,5 Ki-Bum Lee,3,5,6 Scott B. Ficarro,1,7 Andrew Crain,2,4 Yue Xu,3 Alexey Terskikh,2
Evan Y. Snyder,2,* and Sheng Ding3,*
1Genomics
Institute of the Novartis Research Foundation, 10675 John Jay Hopkins Drive, San Diego, CA 92109, USA
Institute for Medical Research, 10901 North Torrey Pines Road, La Jolla, CA 92037, USA
3Department of Chemistry, The Scripps Research Institute, 10550 North Torrey Pines Road, La Jolla, CA 92037, USA
4Biomedical Sciences Graduate Program, University of California, San Diego, 9500 Gilman Drive, La Jolla, CA 92093, USA
5These authors contributed equally to this work
6Present address: Department of Chemistry and Chemical Biology, Institute for Advanced Materials, Devices, and Nanotechnology,
The Rutgers Stem Cell Research Center, Rutgers University, 610 Taylor Road, Piscataway, NJ 08854, USA
7Present address: Dana-Farber Cancer Institute, 44 Binney Street, Boston, MA 02115, USA
*Correspondence: lbrill@burnham.org (L.M.B.), esnyder@burnham.org (E.Y.S.), sding@scripps.edu (S.D.)
DOI 10.1016/j.stem.2009.06.002
2Burnham
SUMMARY
Protein phosphorylation, while critical to cellular
behavior, has been undercharacterized in pluripotent
cells. Therefore, we performed phosphoproteomic
analyses of human embryonic stem cells (hESCs)
and their differentiated derivatives. A total of 2546
phosphorylation sites were identified on 1602 phosphoproteins; 389 proteins contained more phosphorylation site identifications in undifferentiated
hESCs, whereas 540 contained more such identifications in differentiated derivatives. Phosphoproteins
in receptor tyrosine kinase (RTK) signaling pathways
were numerous in undifferentiated hESCs. Cellular
assays corroborated this observation by showing
that multiple RTKs cooperatively supported undifferentiated hESCs. In addition to bFGF, EGFR, VEGFR,
and PDGFR activation was critical to the undifferentiated state of hESCs. PDGF-AA complemented a
subthreshold bFGF concentration to maintain undifferentiated hESCs. Also consistent with phosphoproteomics, JNK activity participated in maintenance
of undifferentiated hESCs. These results support the
utility of phosphoproteomic data, provide guidance
for investigating protein function in hESCs, and
complement transcriptomics/epigenetics for broadening our understanding of hESC fate determination.
INTRODUCTION
Human embryonic stem cells (hESCs) are a model developmental
system that may have potential clinical value for mitigating
diseases. Mechanisms of hESC fate determination are not well
defined, although there has been progress in elucidating molecular circuitry of self-renewing ESCs. Transcriptional profiles of
hESCs (Brandenberger et al., 2004; Sato et al., 2003; Sperger
et al., 2003) and more limited ChIP-on-chip (Boyer et al., 2005)
and proteomic (Bendall et al., 2007; Van Hoof et al., 2006) analyses
204 Cell Stem Cell 5, 204–213, August 7, 2009 ª2009 Elsevier Inc.
suggest mechanisms underlying hESC self-renewal and differentiation. In addition to transcriptional and translational regulation,
cell-fate determination is controlled by protein phosphorylation,
a critical determinant of cell signaling (Mann et al., 2002; Schlessinger, 2000). Recent phosphoproteomic analyses of human
mesenchymal stem cells identified 716 and 703 protein phosphorylation sites (Thingholm et al., 2008a, 2008b). However, protein
phosphorylation has not been well characterized in pluripotent
cells. Therefore, we performed a large-scale multidimensional
liquid chromatography (MDLC)- tandem mass spectrometry
(MS/MS)-based phosphoproteomic analysis of undifferentiated
hESCs and their differentiated derivatives for identification of
protein phosphorylation sites in these cells.
Undifferentiated hESCs were cultured under feeder-free conditions with bFGF. Comparable differentiated derivatives were obtained by removal of bFGF and treatment with retinoic acid (RA),
which induces nearly complete, albeit nonspecific, differentiation
to a heterogeneous population of cells. Removal of bFGF alone
does not result in complete differentiation, whereas concurrent
RA treatment causes virtually complete loss of the undifferentiated population in 4 days (required for this type of analysis). Our
data provide a freely available resource of protein phosphorylation sites in hESCs and differentiated derivatives (http://www.
ebi.ac.uk/pride/). These data have begun to prove informative
and predictive. For example, as proof of concept, pathway analyses of the phosphoproteins suggested potential responses of
hESCs to perturbations of receptor tyrosine kinase (RTK)
signaling pathways. To test some RTK pathways for a role in
the maintenance of undifferentiated hESCs, we treated hESC
cultures with selected agonists or antagonists of these pathways.
Their effects were consistent with predictions of the phosphoproteomic analyses. Furthermore, the data suggested previously
unidentified protein roles in hESC self-renewal or differentiation,
thus providing extensive guidance for future research.
RESULTS
Phosphoproteomic Analysis of hESCs
Because phosphoproteomic analysis is challenging (Mann et al.,
2002) and has not been reported in hESCs, we chose to analyze
Cell Stem Cell
Human ESC Phosphoproteomics
Figure 1. Undifferentiated hESCs Expressed Markers of Pluripotency, whereas
the Markers Were Downregulated upon
Differentiation
Cells were cultured to yield undifferentiated
hESCs (hESCs), or differentiated hESC derivatives
(derivs) under feeder-free conditions by withdrawing bFGF and including 5 mM RA in the media
for the final 4 days of culture. Nuclei were stained
with DAPI (A and B; left column).
(A) Cells were stained with antibodies detecting
OCT4 (center column), and OCT4 and DAPI
images were merged (right column).
(B) Cells were stained with antibodies detecting
SSEA-4 (center column), and SSEA-4 and DAPI
images were merged (right column). All photomicrographs were at the same magnification. The
scale bar represents 50 mM.
the well-characterized hESC line H1 (WiCell; WA01) (Thomson
et al., 1998), which has been used in molecular studies of hESCs
(Bendall et al., 2007; Brandenberger et al., 2004; Wang et al.,
2007). Fifty-nine hESC lines, including H1, showed remarkable
conservation of hESC markers (Adewumi et al., 2007), which
provided confidence that our findings would be representative.
Before analyzing protein phosphorylation, the undifferentiated
hESC markers OCT4 (Thomson et al., 1998) and SSEA-4 (Reubinoff et al., 2000) were examined to assess whether the hESCs were
truly undifferentiated under our culture conditions and whether
differentiation was complete. Undifferentiated hESCs were
cultured on Matrigel-coated plates in feeder-free cultures using
conditioned media (CM) that contained 8 ng/ml of added bFGF.
A heterogeneous population of differentiated derivatives of the
hESCs was obtained by removal of bFGF and treatment with
5 mM RA for 4 days. OCT4 was detected in 97% of the hESCs
under the feeder-free conditions, whereas it was nearly undetectable in differentiated derivatives (Figure 1). Similarly, SSEA-4
was positive in the undifferentiated hESCs and nearly absent in
differentiated derivatives. Moreover, the nucleus-to-cytoplasm
ratio, also monitored as an indicator of whether hESCs are undifferentiated or differentiated, was consistent with OCT4 and
SSEA-4 expression (Figure 1). These observations suggested
that our cells represented two distinct populations—‘‘undifferen-
tiated’’ or ‘‘differentiated’’ hESC derivatives—that might then be reliably subjected to phosphoproteomic analysis,
using MDLC-MS/MS technology, that
can result in unbiased discovery of protein
phosphorylation sites (Kruger et al., 2008).
Phosphoproteomic analyses of hESCs
and their differentiated derivatives were
performed using automated MDLC,
a linear ion trap mass spectrometer, and
readily available bioinformatics algorithms. Phosphorylated peptides from
total proteins from undifferentiated
hESCs or their differentiated derivatives
were separated, enriched, and analyzed
using MDLC comprised of strong cation
exchange chromatography (SCX), reversed-phase (RP) desaltFe3+-immobilized metal affinity chromatography (desalt-IMAC),
and RP HPLC coupled to nano-electrospray ionization-tandem
mass spectrometry (ESI-MS/MS; see a schematic diagram in
Figure S1, available online). IMAC, for phosphopeptide enrichment, coupled to RP HPLC-ESI-MS/MS is a robust technique
for phosphoproteomic analyses (Bodenmiller et al., 2007; Brill
et al., 2004; Gruhler et al., 2005), and automation improves
reliability and reproducibility (Ficarro et al., 2005). Because phosphorylated proteins are frequently at low abundance, substoichiometrically phosphorylated, and difficult to identify (Mann
et al., 2002), replicate analyses were performed to increase
phosphoproteome coverage. Replicates increase proteome
coverage, especially of lower abundance proteins (Liu et al.,
2004), and the impact of experimental variation in LC-MS/MS
can be minimized by replicates (Washburn et al., 2003). Phosphopeptides were identified with high confidence (see Supplemental Experimental Procedures). Examples of typical MS/MS
spectra used to identify phosphopeptides are in Figure S2.
To complement identification, extracted ion chromatograms
(XICs) were used to quantify the relative abundance of phosphopeptides. The normalized abundance of randomly selected
phosphopeptides identified in all four phosphoproteomic analyses (two biological replicates, i.e., phosphopeptides from two
Cell Stem Cell 5, 204–213, August 7, 2009 ª2009 Elsevier Inc. 205
Cell Stem Cell
pairs of independent cultures of undifferentiated hESCs or their
differentiated derivatives) demonstrated relatively low variability
(Table S1). This degree of consistency agrees with previous findings in which proteomic data can be reliably compared among
experiments (Washburn et al., 2003).
In contrast, differential phosphopeptide identification implies
differential phosphopeptide abundance. We used data-dependent MS/MS, and peptide abundance and identification correlate
in data-dependent MS/MS (Liu et al., 2004). Selected phosphopeptides identified in undifferentiated hESC or differentiated
derivative cell populations were also quantified using XICs.
Furthermore, signal from each of the selected phosphopeptides
was manually sought in the MS/MS data from analyses in which it
had not been identified by SEQUEST searches, in order to test
whether the phosphopeptide was detectable and, if so, its relative abundance among the phosphoproteomic analyses. Only
a fraction of the phosphopeptides not identified in SEQUEST
searches were detectable (via a poor quality MS/MS spectrum)
when searching the raw data (Table S2). However, every phosphopeptide that was examined demonstrated a higher normalized abundance in analyses in which it was identified than in
analyses in which it was not identified by SEQUEST searches.
Although lack of identification of a phosphopeptide is not
evidence for its absence, identification versus lack of identification implies that the phosphopeptide is likely to be more abundant in the cell population in which it was identified, consistent
with our results (Table S2) and those of others (Liu et al., 2004).
Western blots were performed on proteins from undifferentiated hESCs and differentiated derivatives, using antibodies
recognizing phosphorylation sites previously identified by
MDLC-MS/MS. All nine antibodies that were used recognized
bands with the expected mobility on western blots, providing
confidence in phosphorylation site identifications.
Representative western blots, including normalized integrated
intensities of phosphoprotein bands, are shown in Figure S3.
Phosphorylation of mTOR on Ser2448 was apparently more
abundant in undifferentiated than differentiated cells (Figure S3A),
and mTOR Ser2448 phosphorylation was identified in undifferentiated, but not differentiated cells, using MDLC-MS/MS (Table
S3A). PAK1 phosphorylation on Ser144 was identified twice
in undifferentiated cells and once in differentiated cells by
MDLC-MS/MS (Table S5A), and western blots suggested that
PAK1 phosphoserine 144 was more abundant in undifferentiated
than in differentiated cells (Figure S3B). Antibodies recognizing
PTK2 phosphotyrosine 576/577 suggested that phosphorylation
of this site was more abundant in differentiated derivatives than
undifferentiated hESCs (Figure S3C), consistent with identification of PTK2 phosphorylated on Tyr576, using MDLC-MS/MS,
only in differentiated derivatives (Table S4A). Phosphorylation
of CDK1/2/3/5 on Thr14 and Tyr15 (two conserved residues in
all four CDK proteins) was more abundant in undifferentiated cells
(Figure S3D), and XIC peak areas suggested that phosphorylation
of CDK1/2/3 on Thr14 and Tyr15 was more abundant in undifferentiated cells (Table S1). CDK1, -2, -3, and -5 phosphorylated
on Thr14 and Tyr15 are recognized in western blots (Supplemental Experimental Procedures), and the corresponding phosphopeptides identified by MDLC-MS/MS (IGEGT*YGVVY and
IGEGTY*GVVY; for brevity, designated as originating from CDK2
in Tables S1 and S5) are identical among CDK1/2/3, whereas the
corresponding peptide from CDK5 differs at two amino acid residues (IGEGT*Y*GTVF), which is easily distinguishable by MS/MS.
The relative abundance of JUN phosphorylated on Ser63, and
HSP27 phosphorylated on Ser82, was similar in undifferentiated
and differentiated cells on western blots (data not shown), and
phosphorylated JUN Ser63 and phosphorylated HSP27 Ser82
were both identified the same number of times in undifferentiated
and differentiated cells (Table S5A), demonstrating further agreement between western blots and MDLC-MS/MS.
If subsequent studies focus on one or a few especially critical
sites of protein phosphorylation, it is advisable to examine the
phosphorylation site using an independent technique. However,
MDLC-MS/MS is reliable for phosphoproteome analysis and can
yield unbiased, large-scale discovery of protein phosphorylation
(Bodenmiller et al., 2007; Brill et al., 2004; Ficarro et al., 2005;
Gruhler et al., 2005; Kruger et al., 2008; Thingholm et al.,
2008a), and our findings support its accuracy. Together, these
results suggest that application of MDLC-MS/MS for identification of phosphopeptides was suitable for phosphoproteomic
analysis of undifferentiated hESCs and their differentiated derivatives.
Phosphopeptide identifications are in Tables S3A–S5B. Each
phosphoprotein, from which phosphopeptides were derived,
was classified as either (1) containing more phosphorylation
site identifications in undifferentiated hESCs, (2) containing
more phosphorylation site identifications in differentiated hESC
derivatives, or (3) containing a similar number of phosphorylation
site identifications in both cell populations. A protein is conservatively defined to contain more phosphorylation site identifications
in a cell population if its phosphorylation was identified exclusively in this population or at least 3-fold more frequently than
in the other population; otherwise, the protein is considered to
contain a similar number of phosphorylation site identifications
in populations from both cell states. Although identification of
protein phosphorylation sites was unlikely to be comprehensive,
as implied by studies using different cell types (Bodenmiller et al.,
2007; Mann et al., 2002), among the 2546 nonredundant phosphorylation sites, 472 were on proteins containing more phosphorylation site identifications in undifferentiated hESCs,
whereas 726 were on proteins containing more phosphorylation
site identifications in differentiated hESC derivatives (Figure 2A).
Of the peptides, 94% were singly phosphorylated, whereas the
rest were doubly phosphorylated, similar to other studies using
IMAC for phosphopeptide enrichment (Bodenmiller et al., 2007;
Thingholm et al., 2008a). Serine, threonine, and tyrosine phosphorylation comprised 82%, 14%, and 4% of the sites,
respectively (Tables S3A–S5B), and tyrosine phosphorylation
was relatively prominent in undifferentiated hESCs (Figure 2C).
Among the 1602 proteins, 389 contained more phosphorylation
site identifications in undifferentiated hESCs, whereas 540 contained more phosphorylation site identifications in differentiated
hESC derivatives (Figure 2B).
Transcription factors can reprogram differentiated cell types
to ESC-like cells when ectopically expressed (Takahashi
et al., 2007; Takahashi and Yamanaka, 2006; Yu et al., 2007)
and were the most abundant known phosphoprotein category
(Figures 2G–2I). This observation, not typical of proteomic analyses, could reflect the growing consensus that many transcription regulators are important in control of ESC state. Among the
Cell Stem Cell
Figure 2. Number of Protein Phosphorylation Sites and Phosphoproteins Identified
in hESCs and Their Differentiated Derivatives, Prominence of Tyrosine Phosphorylation, Predicted Subcellular Location of
the Phosphoproteins, and Phosphoprotein
Categories
(A and B) (A) Total number of nonredundant phosphorylation sites and (B) number of proteins with
more phosphorylation site identifications in undifferentiated hESCs (line H1/WA01) (represented
in red), RA differentiated, H1-hESC derivatives
(represented in gold), or with a similar number of
phosphorylation site identifications in the two cell
populations (represented in gray). The percentage
of the phosphorylation sites and phosphoproteins
in each of the three groups of proteins is shown in
parentheses.
(C) Percentage of nonredundant tyrosine phosphorylation sites, among the sites for which the
phosphorylated residue could be defined as
serine, threonine, or tyrosine (94% of all sites),
that were on proteins containing more identified
sites in undifferentiated hESCs, differentiated
hESC derivatives, or that were on proteins with
a similar number of identified sites between undifferentiated and differentiated cells.
(D–F) The subcellular localization of the phosphoproteins is shown; those widely associated with
more than one subcellular location are designated
as variable.
(G–I) Phosphoprotein categories, among those
whose functions are known, are shown. The
percentage of proteins with known functions are
45.8%, 55.7%, and 57.2% for proteins with more
phosphorylation site identifications in undifferentiated hESCs, differentiated hESC derivatives, or
a similar number of phosphorylation site identifications between the two cell populations, respectively. Each chart progresses from the protein
category containing the most to the fewest entries.
Abbreviations and definitions include the following:
transcript. reg., transcription regulator; enzyme, protein with enzymatic activity outside of the other categories; RNA meta., RNA-binding proteins and proteins
participating in metabolic processes involving RNA; prot. degr., protein degradation; transport., transporter; apop. reg., apoptosis regulator; transmem. recep.,
transmembrane receptor; GEF and GAP, guanine nucleotide exchange factor and GTPase-activating protein; cytoskel., proteins that are components of, closely
associated with, or regulate cytoskeletal function; cell prolif., proteins participating in regulation of cellular proliferation and/or cell-cycle progression; tum. sup.,
tumor suppressor; translat. reg., translation regulator; phosphoinos. sig., proteins participating in phosphoinositide signaling; gen. assem., genome assembly; GF,
growth factor; cell adhes., proteins functioning in cell adhesion; telomere mainten., protein functioning in telomere maintenance; prom. differ., proteins promoting
cellular differentiation; GF buffer, proteins regulating the availability of growth factors; comp. casc., complement cascade; nuc. receptor, ligand-dependent
nuclear receptor; and hormone biosyn., hormone biosynthesis.
158 phosphorylated transcription regulators, 41 contained more
phosphorylation site identifications in undifferentiated hESCs,
46 contained more phosphorylation site identifications in differentiated hESC-derivatives, and 71 contained a similar number
of phosphorylation site identifications in both cell populations.
Most of the transmembrane receptors and predicted extracellular proteins contained more phosphorylation site identifications in either undifferentiated or differentiated hESCs, whereas
fewer of these proteins contained a similar number of phosphorylation site identifications in both cell populations (Figures 2D–2I,
Table S6), implying that growth factors, cytokines, their receptors, and corresponding signaling pathways could participate
in controlling hESC fate. Furthermore, kinases, which are key
players in cell signaling, represented the second-largest cate-
gory of known phosphoproteins (Figures 2G–2I). Phosphorylation of cytoplasmic, cytoskeletal, and cell-adhesion proteins
was identified relatively frequently in differentiated derivatives
(Figures 2D–2I).
Phosphorylated Transcription Regulators
in Undifferentiated hESCs
The transcription regulator ESG1 (official symbol TLE1; Table S7)
is expressed only in preimplantation embryos, ESCs, and
primordial germ cells (Western et al., 2005). ESG1 is coexpressed with OCT4 and SOX2 in both mouse and human
ESCs, suggesting it is a potential pluripotency marker (Western
et al., 2005). In addition, SUPT16H and SSRP1 (Tables S7 and
S8) were phosphorylated in undifferentiated hESCs and are the
Cell Stem Cell
Table 1. Signaling Proteins with More Phosphorylation Site Identifications in Undifferentiated hESCs than in hESC Derivatives
Proteins Participating in Receptor Tyrosine Kinase Signaling
Receptors, Growth
Factors
AREGa, KDR,
IGF2R, EPHA1
Kinases
Phospholipases
Adaptors
Other
LCK, NEK4, MAPK6,
MAPK7, FRAP1 (mTOR),
PIK3C3, PIK3R4, DBF4,
CDC42BPA, CRKL, MINK1,
KIAA1804 (MLK4), CDKL5,
EIF2AK1, CRKRS
PLCG1, PLCG2, PLCH1
SHC1, GAB1, NCK2,
KIAA1303 (RAPTOR),
CNKSR1, CNKSR2, ABI2,
CDC37L1, PLEKHA1
PPAP2B, EPS15L1, TRAF4,
APC, CDH17, IGFBP2,
RAPGEF1,
TRIP10, TSC1, WDR62,
NUMB
Signal Transduction Pathways and Member Proteins
MAPK: JNK, ERK
CRKLb, MINK1, KIAA1804 (MLK4), TRAF4, TRIP10, WDR62, CNKSR1, DBF4, CDC42BPA, RAPGEF1, PLCG1,
SHC1, PLCG2, GAB1, LCK, MAPK6, MAPK7, NEK4, NCK2
PI3K/AKT/mTOR
FRAP1 (mTOR), TSC1, GAB1, PIK3C3, PLCG2, PIK3R4, KIAA1303 (RAPTOR), ANRT (HIF-1b)
a
Official symbols of the proteins, some of which are followed by synonyms in parentheses, are used in this table.
b
Symbols in bold text represent proteins that are relatively specific to JNK signaling.
two subunits of FACT (facilitates chromatin transcription). FACT
destabilizes nucleosomes to allow transcription without disruption of the epigenetic state (Belotserkovskaya et al., 2003) and
promotes initiation of DNA replication in the S phase of the cell
cycle (Tan et al., 2006). CREBBP (Table S7) has histone acetyltransferase activity. Its mRNA is enriched in undifferentiated
hESCs (Brandenberger et al., 2004) (Table S8). AKT (Table
S5A) phosphorylates CREBBP, increasing CREBBP acetyltransferase activity and promoting NF-kB-mediated transcription and
enhanced cell survival (Liu et al., 2006). Furthermore, CREBBP
increases ERK1 expression (Chu et al., 2005). ERK1 activity
contributes to hESC self-renewal in the presence of bFGF (Li
et al., 2007).
At least 18 phosphorylated transcription regulators identified
in undifferentiated hESCs can modify chromatin structure via
histone methylation or acetylation (Table S7) and may contribute
to the epigenetic pattern that is likely to be important to hESCs
(Bernstein et al., 2006; Lee et al., 2006; McCool et al., 2007).
We identified phosphorylation of DNMT3B, MBD3 (Table S3A)
and EZH2 (Table S5A) in undifferentiated hESCs. DNMT3B
encodes a DNA methyltransferase (Table S7), which was expressed in all 59 hESC lines tested (Adewumi et al., 2007), was
enriched in undifferentiated hESCs (Brandenberger et al.,
2004), and was phosphorylated in undifferentiated hESCs (Table
S8). Differential phosphorylation could modulate EZH2 activity.
Phosphorylation at S21 by AKT inhibits the histone H3 Lys27
methyltransferase activity of EZH2 (Cha et al., 2005), and we
identified a phosphorylation site of EZH2 in undifferentiated
hESCs (S371 or T372; Table S5A), a site whose phosphorylation
was also identified in undifferentiated mouse ESCs (L.M.B.,
K.-B.L., W.X., and S.D., unpublished data).
Phosphorylated transcription regulators in undifferentiated
hESCs can participate in transcriptional activation or repression,
histone modification, and more (Table S7). These and other functions may be integrated to favor the undifferentiated state of
hESCs, as implied by the complexity of the phosphoproteome
(Figure 2). Although some of these transcriptional and epigenetic
regulators were previously reported to influence hESCs, the
mechanisms are unclear. The identified phosphorylation sites
provide focused information for future studies of the function
of these factors in hESCs. Furthermore, we also identified
hundreds of phosphoproteins whose presence in hESCs was
unknown, providing a rich resource for further investigation.
For instance, TNRC6A, a factor for gene silencing via RNA interference (Liu et al., 2005), was phosphorylated in undifferentiated
hESCs (Table S3A).
Growth Factor-Mediated Signaling Pathways
Tyrosine phosphorylation, which plays a dominant role in growth
factor/RTK signaling pathways (Schlessinger, 2000), was relatively prominent in undifferentiated hESCs (Figure 2C). Signaling
pathways participating in self-renewal of hESCs include bFGF,
TGF-b/activin, insulin/IGF, EGFR family, PDGF, Wnt, neurotrophin, integrin, and Notch pathways (Beattie et al., 2005; Bendall
et al., 2007; James et al., 2005; Pebay et al., 2005; Wang et al.,
2007; Xu et al., 2005; Yao et al., 2006). However, detailed understanding of the action of these pathways is lacking. The phosphoproteins were grouped into signaling pathways, as described
in the Supplemental Experimental Procedures, to further explore
their functional potential. Forty-one canonical and metabolic
pathways were suggested using the phosphoproteins as input
for pathway analysis (data not shown). Proteins in RTK pathways
were phosphorylated in undifferentiated hESCs, including the
adaptors GAB1, SHC1, and NCK2; the kinases LCK, NEK4,
MAPK6, MAPK7, mTOR, PIK3C3, and PIK3R4; phospholipases
PLC-g1 and PLC-g2; and the phosphatase PPAP2B (Table 1).
Some phosphoproteins are shared among pathways, and
some are more pathway specific, such as APC in Wnt signaling
and NUMB in Notch signaling. Table 1 and Figure 2C imply that
a variety of signaling pathways are important in undifferentiated
hESCs. For example, EGF pathway members ErbB2, AREG,
and EPS15L1 were phosphorylated in undifferentiated hESCs
(Table 1 and Table S5), complementing a report showing that
the ErbB2/ErbB3 ligand heregulin-1b helps support undifferentiated hESCs (Wang et al., 2007). KDR (VEGFR2, FLK1) was
phosphorylated in undifferentiated hESCs (Table 1), and stimulation of hESCs with CM elicits tyrosine phosphorylation (site[s]
undefined) of PDGFRA (Wang et al., 2007). Components of
the VEGF and PDGF pathways were phosphorylated in undifferentiated hESCs, including some proteins in Table 1. We also
identified phosphoproteins from signaling pathways whose
Cell Stem Cell
presence in hESCs has not been reported, and a large number of
proteins not previously known to be phosphorylated (Tables
S3A–S5B).
Molecular profiling studies typically lack biological follow-up
(e.g., Bodenmiller et al., 2007; Boyer et al., 2005; Brandenberger
et al., 2004; Brill et al., 2004; Ficarro et al., 2005; Gruhler et al.,
2005; Lee et al., 2006; McCool et al., 2007; Sperger et al.,
2003; Thingholm et al., 2008a, 2008b; Van Hoof et al., 2006).
However, a few, including transcriptomic (Armstrong et al.,
2006) and proteomic (Bendall et al., 2007; Kratchmarova
et al., 2005; Mukherji et al., 2006; Wang et al., 2006; Wang
et al., 2007) studies, demonstrated that cells responded to stimulation in manners consistent with molecular profiles. To test the
cellular relevance of the phosphoproteomic and pathway analyses, we began by targeting EGF, VEGF, and PDGF pathways
in undifferentiated hESCs using inhibitors of their receptors.
Although specificity of RTK inhibitors is imperfect, we used
some of the widely accepted ones (see the Supplemental Experimental Procedures). Treatment of undifferentiated hESC
cultures with an EGFR inhibitor at 10 mM resulted in extensive
apoptosis (data not shown), similar to another report (Wang
et al., 2007). The hESCs were also treated with 10 mM KDR inhibitor II or 10 mM Gleevec, a PDGFRA inhibitor (Zhang et al., 2003).
Undifferentiated control colonies were compact and expressed
OCT4 and SSEA-4 (Figure 3B and data not shown). In contrast,
most cells differentiated in the presence of KDR or PDGFR
inhibitor, shown by flattening of the colonies, altered cellular
morphology and nearly undetectable OCT4 and SSEA-4
(Figure 3C and data not shown). Vehicle-only controls lacked
any noticeable effect on the cells (Figure 3B). The results were
similar under feeder-free conditions in CM and feeder-free
conditions in chemically defined media (CDM; Yao et al.,
2006). Furthermore, KDR or PDGFR inhibitor, at 10 mM, resulted
in decreased expression of NANOG and OCT4 (Figure 3A).
To further investigate the effect of RTK signaling pathways, we
decreased bFGF to a subthreshold 4 ng/ml (at least 20 ng/ml is
required under feeder-free conditions in CDM [Yao et al.,
2006]) and systematically supplemented cultures with EGF,
PDGF-AA, or VEGF-AA at different concentrations to determine
which trophic factor could complement bFGF deficiency.
Although PDGF-AA without bFGF was unable to maintain longterm cultures of undifferentiated hESCs, PDGF-AA at 10 ng/ml
and the subthreshold concentration of 4 ng/ml of bFGF (subsequently abbreviated PDGF/bFGF) stably maintained undifferentiated hESCs under feeder-free conditions in CDM for >15
passages, and the hESCs remained undifferentiated throughout
all four experiments (Figure 4D). The cells displayed undifferentiated morphology and robust expression of OCT4. In contrast,
when undifferentiated hESCs, which had been stably maintained
in CDM containing PDGF/bFGF for >15 passages, were subsequently cultured for 4 days in CDM containing 4 ng/ml of bFGF
but no PDGF, the cells differentiated (Figure 4B). FACS analyses
demonstrated that 89% of the hESCs in CDM containing
PDGF/bFGF were positive for SSEA-4, comparable to cultures
in CDM containing 20 ng/ml of bFGF (86%; Figure 4). Similar
FACS results were obtained when cells were stained and sorted
for the pluripotency marker Tra-1-60 (data not shown). Moreover, PDGF/bFGF in CDM resulted in sustained expression of
NANOG and OCT4 transcripts, whereas their abundance
Figure 3. Protein Kinase Inhibitors Resulted in Differentiation of
hESCs
(A) Expression of NANOG (Chambers et al., 2003) and OCT4 mRNAs was assessed by RT-PCR, in the presence of protein kinase inhibitors that resulted in
differentiation of hESCs. Cells were cultured with 20 ng/ml of bFGF, and inhibitors (10 mM) were included in the cultures for the final 4 days. Inhibitor identities are indicated in the figure. Slower decline of OCT4 than NANOG was typically observed during hESC differentiation. GAPDH was an internal control.
(B and C) Undifferentiated, vehicle-only control (B) and differentiated, KDR
inhibitor-treated (C) cells are shown under imaging conditions indicated above
the columns. All photomicrographs were at the same magnification, and the
scale bar (bottom right) represents 50 mM. Abbreviations include the following:
i, inhibitor; uhESCs, undifferentiated hESCs.
declined within 4 days in the absence of PDGF-AA or the presence of the PDGFR inhibitor Gleevec (Figures 3A and 4A), further
supporting the proposal that PDGF-AA facilitates maintenance
of undifferentiated hESCs. Together, phosphoproteomic and
pathway analyses suggested that PDGF should favor maintenance of undifferentiated hESCs. PDGFR inhibitor, and separate
use of PDGF-AA, provided clear evidence that PDGF, when
bFGF is at a subthreshold concentration, can promote the
undifferentiated state of hESCs in CDM under feeder-free conditions, insights that derived directly from the phosphoproteomic
analysis.
Our data further suggested that ErbB and VEGFR activation
participate in maintenance of undifferentiated hESCs, because
disruption of these pathways caused apoptosis (data not shown)
and/or differentiation (Figure 3) (although EGF and VEGF-AA
demonstrated limited efficacy at complementing the deficiencies
of 4 ng/ml bFGF). The ErbB2/ErbB3 ligand heregulin-1b contributes to maintenance of undifferentiated hESCs (Wang et al.,
2007). In addition, insulin/IGF pathway members (Bendall et al.,
2007) were phosphorylated in hESCs (including proteins in the
PI3K/AKT/mTOR pathway; Table 1).
Phosphoproteomics, cellular assays, and other reports (Bendall et al., 2007; Wang et al., 2007; Yao et al., 2006) suggest that
multiple RTK pathways are required, although none of them
alone is sufficient to support self-renewal in the absence of
Cell Stem Cell
and pathway analyses also imply that additional pathways could
favor undifferentiated hESCs.
Figure 4. PDGF and a Subthreshold Concentration of bFGF Sustained Long-Term Culture of hESCs
(A) RT-PCR to amplify NANOG and OCT4 transcripts in long-term hESC
cultures (>15 passages) in CDM containing 10 ng/ml of PDGF-AA and 4 ng/
ml of bFGF (lane PDGF, bFGF4). Lanes bFGF20 or bFGF4 refer to 20 or
4 ng/ml of bFGF in the CDM for 4 days, respectively, in the absence of
PDGF, following culture in 10 ng/ml of PDGF-AA and 4 ng/ml of bFGF for
>15 passages.
(B–D) Colony morphology, OCT4 staining, and fluorescence-activated cell
sorting (FACS) demonstrated that PDGF/bFGF in CDM maintained undifferentiated hESCs passaged >15 times. Imaging conditions or FACS analyses of
SSEA-4 expression, detected via Cy3-conjugated secondary antibodies, is
indicated above the columns, and the culture additives that were varied are
indicated beside the rows. In FACS plots, dotted lines delineate boundaries
of fluorescence intensity approximately indicative of cellular identity as undifferentiated hESCs (uhESC) and differentiated hESC derivatives (deriv). Decline
of SSEA-4 is incomplete in differentiated hESCs after 4 days (Figure 1).
Following maintenance of the hESCs in CDM containing bFGF at 4 ng/ml
and PDGF-AA at 10 ng/ml for >15 passages, cells were cultured for 4 days
in CDM lacking PDGF and containing bFGF at 4 ng/ml (B) or 20 ng/ml (C), or
in the continued presence of bFGF at 4 ng/ml and PDGF-AA at 10 ng/ml (D).
All photomicrographs were at the same magnification, and the scale bar
(bottom center panel) represents 100 mM (B–D).
bFGF. Also consistent with our results, although less clear due to
the undefined media that was used, Sphingosine-1-phosphate
plus PDGF contributes to maintenance of undifferentiated
hESCs in the presence of mouse embryonic fibroblasts (MEFs)
or MEF-conditioned media (Pebay et al., 2005). It previously
appeared that bFGF alone might sustain self-renewal of hESCs.
However, as predicted by our phosphoproteomic analysis,
several other factors that exist in serum and/or are secreted by
feeders, acting through autocrine or paracrine effects or as
culture additives, are also important for hESC self-renewal
(Bendall et al., 2007; Wang et al., 2007). Our phosphoproteomic
Phosphorylated Signal Transduction Proteins
PI3K signaling facilitates ESC self-renewal (Armstrong et al.,
2006), and the PI3K pathway is activated by PDGF in mesenchymal stem cells (Kratchmarova et al., 2005), but the mechanism of action of the PI3K pathway has been unclear. PI3K/
AKT/mTOR pathway members were phosphorylated in undifferentiated hESCs. For example, PIK3C3 is enriched in undifferentiated hESCs (Brandenberger et al., 2004), and PIK3C3 was phosphorylated in undifferentiated hESCs (Table 1). mTOR (Table 1)
plays a role in proliferation of undifferentiated hESCs (Wang
et al., 2007) and is phosphorylated at Ser2448 during mitogenic
stimulation (Chiang and Abraham, 2005). mTOR, phosphorylated
at Ser2448 and Ser2454 in undifferentiated hESCs (Figure S3A,
Table S3A) is a protein that enhances cell survival (Peponi et al.,
2006). TSC1 was also phosphorylated in undifferentiated hESCs
(Table 1). TSC1 can limit cell size (Rosner et al., 2003), and its
overexpression caused cells to form compact clusters with
increased reaggregation in vitro (Li et al., 2003), similar to the
small size of undifferentiated hESCs and compact morphology
of hESC colonies. Phosphorylated PI3K/AKT/mTOR pathway
members in undifferentiated hESCs (Table 1) suggest which
pathway members may regulate undifferentiated hESCs.
Phosphoproteins participating in MAPK signaling were identified (Table 1). The ERK pathway contributes to hESC selfrenewal under conditions that include bFGF (Li et al., 2007),
whereas JNK signaling in hESCs has not been reported. Some
phosphoproteins downstream of RTK pathways are relatively
specific to JNK signaling, such as TRAF4, MLK4, CRKL, and
MINK1 (Table 1). To test for JNK signaling in undifferentiated
hESCs, we tested two JNK inhibitors in hESC cultures under
feeder-free conditions in CM. JNK inhibitor II, a small molecule
(SP600125) widely used in JNK studies (Bennett et al., 2001;
Han et al., 2001; Shin et al., 2002), and JNK inhibitor III, a polypeptide (Holzberg et al., 2003), were used. Each inhibitor alone
resulted in cellular differentiation, demonstrated by colony
morphology and decreased OCT4 expression (Figure S4). In
contrast, controls lacking JNK inhibitors, including vehicle-only
controls, remained undifferentiated (Figure S4 and data not
shown). Induction of differentiation by JNK inhibitors was similar
under feeder-free conditions in CDM (data not shown). Furthermore, OCT4 and NANOG mRNA was depleted in the presence
of JNK inhibitor II (Figure 3A). Thus, this phosphoproteomic analysis provides the first suggestion that JNK, an important signal
transduction protein downstream of many RTKs, may facilitate
maintenance of undifferentiated hESCs. Moreover, these experiments further demonstrate agreement between phosphoproteomic and cellular analyses in hESCs.
DISCUSSION
Analysis of molecular mechanisms underlying hESC properties
is essential for optimal use of these cells. Complementing
previous analyses of promoters, transcripts, and protein expression, our phosphoproteomic analysis suggests that multiple
protein phosphorylation events participate in control of hESC
Cell Stem Cell
fate. Application of MDLC-MS/MS-based phosphoproteomics
to pluripotent cells may represent an important tool for stem
cell biologists. While this study focused on its use for hESCs,
one can envision its application to induced pluripotent somatic
cells and other somatic stem cells.
Our phosphoproteomic analyses identified proteins potentially
participating in self-renewal or differentiation of hESCs and
focused attention on pathways heretofore underappreciated
and underexplored. Transcription regulators, including epigenetic and transcription factors, and kinases contained many
phosphorylated members, suggesting that these proteins may
be key determinants of hESC fate decisions. Although a variety
of proteins have been implicated in hESC self-renewal, some
of their functions have been unclear. The identified phosphorylation sites, some on central signaling proteins, expand the knowledge of protein phosphorylation in hESCs. We also identified
many proteins whose potential functions in hESCs had not
been identified previously. In other words, phosphoproteomic
analyses may provide guidance for systematic, rather than solely
serendipitous or overly broad-based, approaches in future
studies of self-renewal and differentiation of pluripotent cells.
Phosphoproteomic analyses identified proteins favoring an
undifferentiated or differentiated state of hESCs. For example,
phosphorylation of proteins in the JNK pathway was identified,
and our cellular follow-up experiments, which are atypical of
molecular profiling studies, suggested that inhibition of JNK
leads to differentiation of hESCs. A role of JNK in undifferentiated hESCs has not been reported. The VEGF and PDGF pathways are candidates to favor maintenance of undifferentiated
hESCs because inhibitors of their receptors resulted in hESC
differentiation. However, the growth factors that were added
singly could not replace bFGF. Together, these results suggested that activation of these pathways is necessary but not
sufficient to sustain self-renewal of hESCs, consistent with
increasing evidence that multiple growth factor-driven pathways
act together to maintain undifferentiated hESCs. For example,
PDGF-AA complemented a subthreshold concentration of
bFGF, shown by long-term maintenance of undifferentiated
cultures under feeder-free conditions in CDM. Use of CDM allowed improved knowledge of the composition of the media,
rather than use of undefined media in the presence of, or conditioned by, feeder fibroblasts (Yao et al., 2006), so the pathways
that were targeted in our cellular assays were more clearly
defined. Together, our results expanded the repertoire of pathways that facilitate hESC culture and support the suggestion
that multiple signaling inputs are needed to maintain undifferentiated hESCs (Wang et al., 2007). Moreover, phosphoproteomic
analyses complement epigenetics, gene expression profiles,
and total protein MS to facilitate an improved understanding of
hESC fate determination.
The functions of most of the phosphorylated proteins in pluripotent cells are unknown and should be evaluated for their influence on stem cell behavior. Application of further advances in
proteomic and allied technologies should enhance future studies
through improved analysis of protein phosphorylation. As phosphoproteins controlling pluripotent behavior are understood
better, methods for developing model systems with stem cells,
and potential therapeutic applications may become increasingly
clear.
EXPERIMENTAL PROCEDURES
Cell Culture, Phosphoproteomic Analysis
Feeder-free cultures were in Matrigel-coated plates in CM containing 8 ng/ml
bFGF (Xu et al., 2001). Differentiation was with 5 mM RA and no added bFGF. In
CDM, hESCs were cultured in Matrigel-coated plates in N2/B27-CDM (Yao
et al., 2006). Phosphoproteomic analyses used cells from CM. Cells were
rinsed with PBS, lysed, and centrifuged, and proteins were precipitated with
(NH4)2SO4 and pelleted by centrifugation.
Proteins were resuspended in 100 mM NH4HCO3, 8 M urea containing phosphatase inhibitors, reduced, alkylated, digested with trypsin, and peptides
desalted. Peptides were separated by SCX, phosphopeptides enriched by
desalt-IMAC (Brill et al., 2004; Ficarro et al., 2005), separated by nanoflow
HPLC, and analyzed by ESI-MS/MS. MS/MS spectra were matched to amino
acid sequences using SEQUEST (Eng et al., 1994). All reported phosphopeptide identifications were manually verified (Bernstein et al., 2008; Brill et al.,
2004; Ficarro et al., 2005).
Normalized XIC peak areas of some phosphopeptides were quantified. For
analyses lacking the identification, MS/MS data were exhaustively searched
for the phosphopeptide, which was rarely found via a poor quality MS/MS
spectrum, and its XIC peak area was quantified.
Phosphoproteins were classified as containing more phosphorylation site
identifications in undifferentiated hESCs or differentiated derivatives, or as
containing a similar number of phosphorylation site identifications in the two
cell populations, as described in the Results.
Western Blot Analysis
Proteins were run on Bis-Tris gels, transferred to PVDF membranes, blocked,
and incubated with antibodies recognizing phosphorylation sites identified
by MDLC-MS/MS. Anti-GAPDH was the loading control. Membranes were
washed, incubated with fluorophore-conjugated secondary antibodies,
washed, imaged, and bands quantified according to the manufacturer (LI-COR).
Phosphoprotein Category, Subcellular Location, and Pathway
Analysis
Ingenuity Pathway Analysis, Metacore, NCBI, Gene Ontology, and peerreviewed literature were used to identify phosphoprotein subcellular location,
category, and signaling pathways.
Cellular Assays, RT-PCR
EGFR, JNK, or PDGFR inhibitors were used. Untreated and vehicle-only
controls were included for each experiment. PDGF-AA/bFGF was used in
cultures for >15 passages.
For immunostaining and DAPI staining, monoclonal mouse anti-OCT4 and
anti-SSEA-4 were used. Secondary antibodies were Cy2-conjugated rabbit
anti-mouse IgM and Cy3-conjugated rabbit anti-mouse IgG. For RT-PCR,
mRNA was isolated and cDNA was synthesized; OCT4, NANOG, and GAPDH
were amplified. For FACS, cells were incubated with mouse monoclonal antiSSEA-4 or anti-TRA-1-60 antibodies, washed with PBS, and incubated with
Cy3-conjugated rabbit anti-mouse IgG.
Details are in the Supplemental Experimental Procedures.
ACCESSION NUMBERS
All supplemental data are deposited in the PRIDE database (http://www.ebi.
ac.uk/pride/) under accession numbers 9253–9257 and 9259–9264.
SUPPLEMENTAL DATA
Supplemental Data include four figures, Supplemental Experimental Procedures, Supplemental References, and 11 tables and can be found with this
article online at http://www.cell.com/cell-stem-cell/supplemental/S19345909(09)00286-0.
ACKNOWLEDGMENTS
We thank Fang C. Kuan, Andrew Su, Jeff Janes, and Ali Iranli for help with
bioinformatics; and Michelle Stettler-Gill, Anthony Boitano, Jacqueline
Cell Stem Cell
Lesperance, and Brandon Nelson for technical assistance. Support was from
a postdoctoral fellowship from the California Institute for Regenerative Medicine (CIRM) (K.-B.L.), the Genomics Institute of the Novartis Research Foundation (GNF), and the 1 P20 GM 075059-01.
Received: October 23, 2008
Revised: May 7, 2009
Accepted: June 9, 2009
Published: August 6, 2009
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Biomed Microdevices (2009) 11:547–555
DOI 10.1007/s10544-008-9260-x
Integrated microfluidic devices for combinatorial cell-based
assays
Zeta Tak For Yu & Ken-ichiro Kamei &
Hiroko Takahashi & Chengyi Jenny Shu &
Xiaopu Wang & George Wenfu He & Robert Silverman &
Caius G. Radu & Owen N. Witte & Ki-Bum Lee &
Hsian-Rong Tseng
Published online: 9 January 2009
# Springer Science + Business Media, LLC 2008
Abstract The development of miniaturized cell culture platforms for performing parallel cultures and combinatorial
assays is important in cell biology from the single-cell level
to the system level. In this paper we developed an integrated
microfluidic cell-culture platform, Cell-microChip (CellμChip), for parallel analyses of the effects of microenvironmental cues (i.e., culture scaffolds) on different mammalian
cells and their cellular responses to external stimuli. As a
model study, we demonstrated the ability of culturing and
assaying several mammalian cells, such as NIH 3T3 fibroblast, B16 melanoma and HeLa cell lines, in a parallel way.
For functional assays, first we tested drug-induced apoptotic
responses from different cell lines. As a second functional
assay, we performed “on-chip” transfection of a reporter gene
encoding an enhanced green fluorescent protein (EGFP)
followed by live-cell imaging of transcriptional activation of
cyclooxygenase 2 (Cox-2) expression. Collectively, our CellμChip approach demonstrated the capability to carry out
parallel operations and the potential to further integrate
advanced functions and applications in the broader space of
combinatorial chemistry and biology.
Keywords Microfluidic devices . Cell-based assay .
Apoptosis . Transfection . Cell culture
Electronic supplementary material The online version of this article
(doi:10.1007/s10544-008-9260-x) contains supplementary material,
which is available to authorized users.
Zeta Tak For Yu and Ken-ichiro Kamei contributed equally to this work.
Z. T. F. Yu
Department of Mechanical and Aerospace Engineering,
University of California,
Los Angeles, CA 90095, USA
C. J. Shu : O. N. Witte
Department of Microbiology, Immunology, and Molecular
Genetics, University of California,
Kamei
Takahashi·: X.
X. Wang
Wang :· G.
G.W.
W.He
He: ·
Z. T. F. Yu : K.-i.
Kamei
· :H.H.Takahashi
C. G. Radu (*) : H.-R. Tseng (*)
Crump Institute for Molecular Imaging, University of California,
e-mail: caiusradu@ucla.edu
e-mail: hrtseng@mednet.ucla.edu
O. N. Witte
The Howard Hughes Medical Institute, University of California,
R.Silverman
Silverman: ·
Kamei
Kamei
Wang·: G.
G. W.
W. He
He :· R.
Z. T. F. Yu : K.-i.
· :X.X.Wang
C. G. Radu : O. N. Witte (*) : H.-R. Tseng
Department of Molecular and Medical Pharmacology,
University of California,
e-mail: owenw@microbio.ucla.edu
K.-B. Lee (*)
Department of Chemistry and Chemical Biology,
Institute for Advanced Materials, Devices and Nanotechnology,
The Rutgers Stem Cell Research Center, Rutgers,
The State University of New Jersey,
Piscataway, NJ 08854, USA
e-mail: kblee@rci.rutgers.edu
DO09260; No of Pages
548
1 Introduction
Conventional cell-based experiments are typically performed on a large cell population. Researchers have begun
to appreciate that there are local variables associated with
the heterogeneous microenvironment in the macroscopic
culture setting, which often leads to experimental inconsistency. On the other hand, differences among individual
cells are often ignored since the conventional assays (e.g.,
Weston blots and microarray analysis) are conducted at a
collective fashion. As a result, it is challenging to elucidate
complex cellular systems and analyze dynamic signaling
pathways (Irish et al. 2006) using the conventional
experiment systems. To overcome these challenges, it is
essential to develop a new technology platform to enable
(1) improved control on the cell culture microenvironment,
(2) precise cell assays with the single-cell resolution, and
(3) sequential and parallel operation by combining those
mentioned in (1) and (2). We envision such a technology
can be applied for screening drug candidates (Dittrich and
Manz 2006; Padron et al. 2000), evaluating biological
pathways (Minor 2003), and understanding pharmacological
effects (Hill et al. 1998; Umezawa 2005), thus constituting
critical technological foundations for a broad spectrum of
biomedical research.
Microfluidic devices (Auroux et al. 2002; Dittrich and
Manz 2006; Dittrich et al. 2006; Reyes et al. 2002) offer a
robust analytical tool that allows rapid analysis of cellular
responses to external stimuli in a parallel way. Moreover,
microfluidics, with its intrinsic advantages of sample/reagent
economy, precise control over physical and chemical microenvironments, high throughput, scalability and digital controllability, whose features allow microfluidics to investigate
complex and dynamic biological processes at the single-cell
level. In addition, microscale cell culture using microfluidics
allows investigating the function of microenvironmental cues at
the single-cell level. Especially, compared to static microfluidic
cell-culture systems, integrated microfluidics allows for the
control of adding and/or removing of biochemical cues at
specific temporal as well as spatial points. This unique
advantage makes it possible to do novel microenvironmental
experiments, such as cell–cell interactions and extracellular
matrix (ECM)–cell interactions. However, there are several
issues associated with the fabrication and control of integrated
microfluidic cell culture systems in order to achieve routine
cellular assays of mammalian cells. Even though microfluidic
cell culture systems have been developed extensively, they can
mainly be described into two ways: what kinds of cell culture
platforms were used, and what types of cells and applications
were tested. Many different cell culture platforms have been
developed, including two-dimensional, three-dimensional
(Cartmell et al. 2003; Toh et al. 2007) and co-culture platforms
(Sin et al. 2004). Those microfluidic approaches enabled to
culture and assay many different cells, such as liver (Kane
et al. 2006; Sin et al. 2004; Zhang et al. 2008), muscle
(Tourovskaia et al. 2005; Tourovskaia et al. 2006), neural
(Millet et al. 2007; Park et al. 2006), and stem cells (Chung
et al. 2005; Gomez-Sjoberg et al. 2007; Kim et al. 2006).
Despite these recent advances, several critical questions and
several challenges should be addressed more explicitly, and
optimized to fully achieve the potential of microfluidic cell
culture and assays. It requires the abilities: (1) to test a robust
and flexible fluidic configuration (e.g., flow through vs.
circulation) for cell culture and (2) to fabricate microfluidic
network capable of performing sequential and parallel
operations such as parallel culture of multiple cell types
and subsequent phenotypic and functional assays in the same
microfluidic chip. In this context, incorporation of isolation
valves (Unger et al. 2000) and peristaltic pumps (Chou et al.
2001) should allow individual addressability and digital
controllability (Lee et al. 2005; Wang et al. 2006) of each
cell culture chamber embedded on a microfluidic device,
which in turn should enable complex manipulations of the
culture microenvironments as well as multiple analytical
measurements.
Over the years, there have been a variety of microfluidic
chips directed for functional biological assays. These
include differentiation of cell through different flow rates
(Gu et al. 2004), fully automated cell culture system by
two-layer PDMS chips (Gomez-Sjoberg et al. 2007),
optimization of drug cocktail to regulate cell activities
through closed-loop control algorithm and microfluidic
platform (Wong et al. 2008), and modelling of galactose
pathway in an alternating culture environment (Bennett et
al. 2008). All these devices or systems have provided
additional modules and thus are superior to the conventional setting on conducting biological research or routine
operations.
In this paper, we describe the design and operation of
polydimethylsiloxane (PDMS)-based Cell-microChip (CellμChip, Fig. 1) envisioned as a digitally controlled platform
for performing parallel cell culture and sequential cellular
assays. The potential of the Cell-μChip to support the
optimal culture and assays of human and murine cell lines
was demonstrated. To determine the optimal culture
conditions, these cells were cultured in six cell culture
chambers embedded on a Cell-μChip under two different
medium supply modes in parallel (i.e., circulation and
direct feeding). The growth and viability of the microcultures were monitored and quantified in real time in the
Cell-μChip using an integrated CCD camera. Sequential
staining with acridine orange (AO) and propidium iodide
(PI) (Hudson et al. 1969; Traganos et al. 1977) allowed
the identification of viable and dead cells, respectively. To
Fig. 1 (a) Schematic representation of an integrated CellmicroChip (Cell-μChip) for
performing multiple cell culture
and assays under a digitally controlled interface. Three pairs of
parallel-oriented cell culture
chambers are incorporated in a
Cell-μChip, where multiple cell
types can be cultured under two
different modes of medium supply, i.e., circulatory (channels i, iii
and v) and direct feeding (channels ii, iv and vi). The operation of
this microchip is controlled by
pressure driven valves with their
delegated functions indicated by
their colors: red for regular valve
(for isolation and gating) and
yellow for pumping valve (for
fluid transport and circulation).
(b) Optical image of the actual
device. The microchip was loaded
with various colors of food dyes
to enhance the visualization of
different parts in the entire system: red and yellow as in (a); blue
indicates the flow channel and the
medium reservoir
549
(a)
Outlet
port
Inlet
port
i
Medium
tubing ii
iii
iv
v
vi
(1) Cell culture
chambers i vi:
100– 150 nL
(2) Metering
pump
Medium
reservoir
5–15µL
Collection
Regular valve
Valve for pump
(b)
demonstrate the ability to perform “on chip” functional assays,
we analyzed drug-induced apoptotic responses. Furthermore,
we showed that the Cell-μChip is amenable to complex
sequential operations such as genetic manipulation and
monitoring of transcriptional activation of gene expression.
2 mm
2 Experimental
outside the culture chambers were removed by washing with
fresh DMEM. The Cell-μChip was placed in an incubator for
6 h. The pump was turned on to introduce the DMEM in a
circulating or feed through fashion in the respective culture
chambers. The flow rate of the medium was controlled in the
range of 0.1–4 nL s−1. Cell growth was monitored by
collecting bright field micrographs of cells inside the
Cell-μChip at 12-h intervals.
2.1 Microfluidic cell culture
2.2 Immunoassay for fibronectin
Using the integrated valves and pumps, bovine fibronectin
(FN) (Sigma) solution (1.0 mg mL−1) filled in Teflon tubing
was introduced into the six cell culture chambers of the
Cell-μChip. The Cell-μChip was kept at 37°C for 30 min
for FN coating. DMEM was then introduced into the cell
culture chambers to extrude the FN solution. The medium
reservoirs and medium tubings were then dead-end filled
with DMEM at 10 psi for 60 min (Song et al. 2008). Then,
individual cell suspensions (NIH 3T3, HeLa and B16) with 2×
106 cells mL−1 were sequentially introduced by gravitation
into the cell culture chambers. After cell loading, cells located
To confirm FN coating efficiency in cell culture chambers,
immunoassay for FN was performed. After FN coating in
a Cell-μChip, blocking solution containing 5% BSA and
0.1% N-dodecyl-β-D-maltoside (DDM) (Pierce) was introduced in a Cell-μChip, and then kept at room
temperature for 1 h. Mouse anti-FN (BD Biosciences)
was loaded into cell culture chambers, and incubated at
room temperature for 2 h. Excess antibody in cell culture
chambers were rinsed with PBS with 0.1% Tween 20
(PBS-T) twice. Then secondary goat anti-mouse IgG
conjugated with Alexa555 (Invitrogen) were introduced in
550
cell culture chambers and incubated at room temperature for
1 h. After washing with PBS-T twice, fluorescent intensity
was measured with a fluorescent microscope, and quantified
with MetaMorph.
2.3 AO/PI fluorescence staining
After culturing the cells in the Cell-μChip for 4 days, a
solution composed of DMEM, AO and PI in a ratio of
10:1:1 was introduced into the cell culture chambers. After
5 min incubation at 37°C, the cells were imaged under a
fluorescence microscope.
3 Results and discussion
3.1 Design of the Cell-μChip
The PDMS-based Cell-μChips (Fig. 1) were fabricated by
multilayer soft lithography approach (see supplementary
information) (Unger et al. 2000; Xia and Whitesides 1998).
It is important to note that the biocompatible and gaspermeable properties (Kim et al. 2007; Korin et al. 2007) of
PDMS matrices help to retain proper physiological
(a)
30 min
2h
1 mg mL-1
NIH 3T3, HeLa and B16 cells were cultured in the cell
culture chambers of the Cell-μChip for 24 h. Staurosporine
(0, 0.1, 1 and 10 μM as final concentrations) or
actinomycin D (0, 0.1, 1 and 10 μM as final concentrations)
in DMEM culture medium were loaded into the cell culture
chambers. After 2 h incubation, the medium in all cell
culture chambers was replaced by the MitoTracker Red
dye (Invitrogen) for staining of viable cells. The CellμChip was incubated for 30 min at 5% CO2, 37°C.
Sequentially, the MitoTracker Red solution was replaced
by a solution containing 100 μL of Annexin binding
buffer and 5 μL of Annexin V-Alexa488 for staining the
apoptotic cells. Following 15 min incubation at RT, cell
culture chambers were flushed with Annexin binding
buffer and the cells were imaged using a fluorescence
microscope.
0.01 mg mL-1
2.4 Apoptosis assay
(b)
NIH 3T3 cells were loaded into the six culture
chambers of the Cell-μChip and were allowed to settle
for 24 h. Cells were transfected with the pCox2-EGFP
plasmid (a kind gift from Prof. Harvey R. Herschman,
UCLA), encoding an enhanced green fluorescent protein
(EGFP) under a murine Cox-2 promoter (Liang et al.
2004). The transfection mixture containing plasmid
(0.5 μg), medium (30 μL) and transfection reagent
(2.5 μL, Superfect reagent, Qiagen) was incubated at RT
for 10 min. The mixture was further diluted with 150 μL of
DMEM and loaded into all culture chambers of the CellμChip. After 3-h incubation at 5% CO2, 37°C, the mixture
was replaced with serum-free DMEM and cells were
incubated for an additional day. To activate the Cox-2
promoter the culture media in three out of six chambers
was replaced with media containing the induction agent
TPA (50 ng mL−1). Following 7 h incubation at 5% CO2,
37°C, EGFP expression was imaged using a fluorescence
microscope.
Fluorescence Intensity (A.U.)
2.5 On-chip transfection and reporter gene imaging
4000
3500
30 min
2h
3000
2500
2000
1500
1000
500
0
0.01 mg mL-1
1 mg mL-1
Fibronectin Concentration
Fig. 2 Fibronectin coating efficiency on the PDMS surface in a CellμChip determined by immunofluorescence assay. (a) Fluorescence
images of immunostained FN on the PDMS surface. (b) Quantitative
analysis of FN coating efficiency determined with fluorescence
images shown in (a)
551
(a)
Day 0
Day 1
Day 2
Day 7
Day 8
200 µm
Day 3
matic valves and peristaltic pumps. This design enabled to
digitally control sequential operations. The chip consisted
of three identical pairs of parallel-oriented culture chambers
with identical dimensions (3×0.5×0.1 mm3, corresponding
to a volume of 150 nL). To allow synchronized pumping,
six internally connected peristaltic pumps were incorporated
at the ends of the six cell culture chambers.
Each pair of culture chambers was configured to have
two types of medium supplies: one allowing media
recirculation through the culture chamber for cellular
auto-conditioning and the other enabling direct feeding of
cells with fresh media. The size of medium reservoir could
accommodate about 10 μL of culture media, a volume
sufficient to sustain continuous on-chip cell culture for
8 days. In contrast, supply Teflon tubings were utilized to
(a)
(b)
(b)
(c)
(c)
200 µm
NIH 3T3
HeLa
B16
(d)
20
(e)
100 µm
Fig. 3 Long term culture of NIH 3T3 cells in a Cell-μChip. (a) Time
lapse images of the NIH 3T3 cell proliferation in the microchip in a
duration of 8 days. (b)–(e) Dead (PI)/live (AO) staining of NIH 3T3
cells cultured in a Cell-μChip for 4 days. (b) A bright field
micrograph of NIH 3T3 cell morphologies. (c) A green fluorescence
micrograph of the live-stained cells. (d) A red fluorescence micrograph of the dead-stained cells. (e) A merged fluorescence image of
(b) and (c)
conditions for a wide range of mammalian cells suitable for
different screening applications. A fluidic network for
individually addressable cell culture chambers and solution/reagent transport was integrated with embedded pneu-
Fold Changes of Cell Number
NIH 3T3
(d)
HeLa
15
with medium
re-circulation
B16
NIH 3T3 with direct
medium feeding
10
5
0
0
2
4
6
Days
Fig. 4 Demonstration of six parallel cell cultures in a closely related
microenvironment. After 3 days culture, the cell morphologies were
shown in (a) NIH 3T3, (b) HeLa and (c) B16. (d) Growth curves of
chip-cultured NIH 3T3, HeLa and B16 cells were quantified by
monitoring the number of cells inside the cell culture chambers over
time. After 5 days culture, we could not count cell number precisely
due to cell confluence and multiple cell layers in the cell culture
chambers
552
Staurosporine (µ
µM)
1
10
0.1
1
10
Apoptosis
NIH3T3
Living
Apoptosis
HeLa
Living
Apoptosis
B16
Living
0.1
ActinomycinD (µM)
Fig. 5 Multiparametric apoptosis assays performed in the CellμChips. NIH 3T3, HeLa and B16 cells were treated with either
staurosporine or actinomycin D to induce apoptosis. Apoptotic cells
were stained with Annexin V conjugated with Alexa488 (green), and
living cells were stained with MitoTracker (red)
store and deliver fresh culture media. This design allowed
us to perform six cell culture experiments in a closely
related microenvironment under two different culture media
supply modes.
searching optimal ECMs, FN is effectively coated on the
PDMS surface in our Cell-μChip (Fig. 2). Several different
conditions, such as different FN concentrations and
incubation time, were tested to optimize the FN coating
condition. We confirm the efficiency and homogeneity of
FN coating on PDMS by immunoassay. As a result, 1 mg
mL−1 of FN for 30 min incubation at 37°C is the optimal
condition in a Cell-μChip.
3.2 Surface modification with fibronectin
We initially used bare PDMS surface to seed cells,
however, we found that cells either could not attach to the
surface well or they detached so easily when new fresh
media were supplied. We reasoned that this problem was
due to the inherent hydrophobicity of PDMS materials.
Thus, we tested to use ECMs in order to make the surface
hydrophilic and biocompatible for cell adhesion. In our
3.3 Cell culture in the Cell-μChip
We initially demonstrated to culture NIH 3T3 cells in all
culture chambers of a Cell-μChip (Fig. 3, Supplementary
Information and Fig. S1). Generally, following loading on
(a)
(b)
20µm
(c)
(d)
553
numbers increased significantly for all three cell lines. To
examine cell viability in the three reservoir-attached cell
culture chambers, we performed sequential PI/AO staining.
As a result, most of the cells are viable, and few dead cells
were observed (data not shown). To quantify the culture
parameters in the Cell-μChip we sought to determine
growth rate for the three cell types (Fig. 4(d)). We observed
that the curves reflected a linear growth phase until the cells
reached confluence followed by a stationary phase. Comparing the cell growth with medium-recirculation or direct
feeding setting, first we used NIH 3T3 cells and continuously monitored them. There was no clear difference in
NIH 3T3 cell growth between the two settings (Fig. 4(d)).
In the case of HeLa and B16 cells, we obtained similar
results as NIH 3T3 cells (data not shown). This result
indicates that a Cell-μChip serves a platform to perform
multiple cell culture in a single device.
3.5 On-chip apoptosis assay
Fig. 6 On-chip transfection and EGFP induction in NIH 3T3 cells.
The plasmid vector which encodes EGFP driven by a Cox-2 promoter
was transfected with NIH 3T3 cells. (a) Bright field and (b)
fluorescence images of NIH 3T3 cells stimulated with TPA for 7 h.
(c) Bright field and (d) fluorescence images of NIH 3T3 cells without
stimulation
the chip, it took about 1 h for cells to attach and spread onto
the FN-coated culture chamber surfaces. NIH 3T3 cells
grew to confluence, and the whole microchannel was fully
occupied at day 8. During this period, we continuously
monitored cell growth inside the Cell-μChip using a CCD
camera (see ESM Movie 1). Interestingly, cells in a CellμChip grew on the bottom as well as ceiling. This
phenomenon is not allowed under conventional culture
conditions using Petri dishes. This observation indicated
that a Cell-μChip could provide unique and intrinsic
characteristics of cell culture manipulations. To determine
cell viability we performed AO/PI fluorescence staining
(Fig. 3(b)–(e)). AO staining (Fig. 3(c)) indicated that the
majority of cells in the chamber were viable. PI staining
(Fig. 3(d)) showed a small percentage of dead cells.
3.4 Parallel culture of multiple cell lines in a single
Cell-μChip
To demonstrate this concept, we cultured NIH 3T3, HeLa
and B16 cells in a single Cell-μChip (Fig. 4). NIH 3T3,
HeLa and B16 cells were sequentially loaded into the pairs
of culture chambers; i–ii, iii–iv and v–vi accordingly. After
cell adhesion, the six peristaltic pumps were turned on to
feed cells with the medium, in either recirculation or direct
feeding setting. The results were consistent with those
observed for the parallel culture of NIH 3T3 cells. Cell
Apoptosis is not only fundamentally involved in developing cells and maintaining tissue homeostasis, but also can
be closely related to several diseases including cancer,
autoimmune, and neurodegeneration. Even though, extensive studies have been reported to dissect apoptosis’s
molecular basis, more system level as well as single-cell
level analysis by using integrated microfluidics would bring
new insights for the underlying mechanisms of these
biological processes. To demonstrate the capability of “on
chip” functional analyses with the Cell-μChip, we performed a drug-induced apoptosis assay (Fig. 5). We used
two kinds of apoptosis inducers, staurosporine (ST)
(Rajotte et al. 1992; Tafani et al. 2001; Wang et al. 1996)
and actinomycin D (AD) (Martin et al. 1990). NIH 3T3,
HeLa and B16 cells were treated with apoptosis inducers at
four concentrations (0, 0.1, 1 and 10 μM) in the CellμChips. ST or AD treated cells increased apoptotic cell
population with a dose-dependent manner of apoptosis
inducers. Although cell viability among untreated and
treated cells appeared to be the same based on the
MitoTracker staining, the cells underwent the apoptosis
process at various speeds according to the drug concentration, as demonstrated by the Annexin V staining. We
conclude that the Cell-μChip served as a platform to
perform multiparametric functional assays.
3.6 On-chip transfection and monitoring of reporter gene
expression
Plasmid DNA transfection is one of the common methods
to manipulate gene expression in mammalian cells.
However, the detailed conditions of optimal transfection
for different cell lines are variable. Combinatorial approach
554
from parallel microfluidic cellular assays can help to
identify the optimal condition. To demonstrate the feasibility
of performing the other assay in the Cell-μChip, gene
transfection experiments were carried out (Fig. 6). For
proof-of-concept, we used a plasmid vector with an
enhanced green fluorescent protein (EGFP) driven by a
cyclooxygenase-2 (Cox-2) promoter. Therefore, EGFP
expression serves as a reporter of Cox-2 transcription. The
basal expression level of Cox-2 in NIH 3T3 cells is very low.
Tetradecanoylphorbol acetate (TPA) can activate Cox-2
transcription. EGFP expression in these cells was monitored
under a fluorescence microscope. As shown in Fig. 6, even
though weak EGFP signals were detected in the negative
control chambers, strong EGFP signals were observed in
TPA-induced transfected cells in the corresponding cell
culture chambers.
4 Conclusion
In summary, we developed a fully digitally controlled
microfluidic cell culture and assay platform that could
support parallel cell culture and sequential cell assays.
Through the integration of isolation valves, murine and
human cells lines could be cultured in different cell culture
chambers and tested for different conditions of cell culture
and assays in a single Cell-μChip. Real-time monitoring of
cell morphology and numbers, viability assay, apoptosis
assay and transfection to monitor expression of a reporter
gene vector were also performed using the same platform.
Our results indicate that intrinsic advantages of microfluidic
devices enable the execution of complicated and integrated
biological operations in stand-alone devices such as the
Cell-μChip. We envision that this platform will be further
integrated with advanced functions and utilities for more
sophisticated cell culture applications.
Acknowledgements This research was supported by the NIH
NanoSystems Biology Cancer Center, the DOE-UCLA Institute of
Molecular Medicine and the NIH-UCLA Center for In Vivo Imaging
in Cancer Biology and Siemens Medical Solutions USA Inc. We thank
Stephanie M. Shelly, Dan Rohle, Shirley Quan and Mireille Riedinger for
the outstanding technical support with conventional cell culture conditions. ONW is an Investigator of the Howard Hughes Medical Institute.
CGR was supported by a Developmental Project Award (ICMIC, NIH/
NCI grant no. CA08630). C.J.S. was supported by a National Institutes of
Health (NIH) Research Training in Pharmacological Sciences training
grant PHS T32 CM008652.
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COMMUNICATION
www.rsc.org/loc | Lab on a Chip
Microfluidic image cytometry for quantitative single-cell profiling of human
pluripotent stem cells in chemically defined conditions†
Ken-ichiro Kamei,‡*abcde Minori Ohashi,‡abcde Eric Gschweng,ef Quinn Ho,abcdeg Jane Suh,abcde Jinghua Tang,e
Zeta Tak For Yu,abcde Amander T. Clark,eh April D. Pyle,ef Michael A. Teitell,ei Ki-Bum. Lee,j
Owen N. Witteacefk and Hsian-Rong Tseng*abcde
Received 4th November 2009, Accepted 11th February 2010
First published as an Advance Article on the web 16th March 2010
DOI: 10.1039/b922884e
Microfluidic image cytometry (MIC) has been developed to study
phenotypes of various hPSC lines by screening several chemically
defined serum/feeder-free conditions. A chemically defined hPSC
culture was established using 20 ng mL1 of bFGF on 20 mg mL1 of
Matrigel to grow hPSCs over a week in an undifferentiated state.
Following hPSC culture, we conducted quantitative MIC to perform
a single cell profiling of simultaneously detected protein expression
(OCT4 and SSEA1). Using clustering analysis, we were able to
systematically compare the characteristics of various hPSC lines in
different conditions.
Human pluripotent stem cells (hPSCs) such as human embryonic
stem cells (hESCs)1 and human induced pluripotent stem cells
(hiPSCs)2–6 exhibit unique characteristics and may provide great
opportunities for cell-based therapy and regenerative medicine. These
characteristics include unlimited propagation capacity in the undifferentiated stage with a normal euploid karyotype and the ability to
differentiate into all cell types in the human body.
a
Department of Molecular & Medical Pharmacology, University of
California, Los Angeles, CA, 90095, USA. E-mail: kkamei@mednet.
ucla.edu; hrtseng@mednet.ucla.edu
b
Crump Institute for Molecular Imaging, David Geffen School of Medicine,
University of California, Los Angeles, CA, 90095, USA
c
California NanoSystems Institute, University of California, Los Angeles,
CA, 90095, USA
d
Institute for Molecular Medicine, University of California, Los Angeles,
CA, 90095, USA
e
Eli and Edythe Broad Center of Regenerative Medicine and Stem Cell
Research, University of California, Los Angeles, CA, 90095, USA
f
Department of Microbiology, Immunology and Molecular Genetics,
g
Department of Biological Chemistry, University of California, Los
Angeles, CA, 90095, USA
h
Department of Molecular Cell and Developmental Biology, Los Angeles,
CA, 90095, USA
i
Department of Pathology and Laboratory Medicine, Los Angeles, CA,
90095, USA
j
Department of Chemistry & Chemical Biology, Institute for Advanced
Materials, Devices and Nanotechnology, Rutgers Stem Cell Research
Center, Rutgers, The State University of New Jersey, Piscataway, NJ,
08854, USA
k
Howard Hughes Medical Institute, David Geffen School of Medicine,
† Electronic supplementary information (ESI) available: Fabrication of
microfluidic hPSC array, generation of hiPSC (i.e., hiPSA1 and
hiPSB2), microscopy settings and image processing are available. See
DOI: 10.1039/b922884e
‡ These authors contributed equally to this work.
Typically, hPSC culture conditions contain serum such as
KnockOut serum replacement (KSR) and feeders such as mouse
embryonic fibroblasts (MEFs). Although these conditions can
successfully maintain pluripotency of hPSCs, these animal products
could cause xenogenic contamination and immunorejection in
patients after transplantation of hPSCs, posing a major challenge to
the use of hPSCs in cell-based therapy applications. Additionally,
these factors are undefined and some are proprietarily formulated,
forming an obstacle in being able to systematically study the regulation of stem cell biology. Therefore, it is essential to develop serum/
feeder-free culture methods for hPSCs in order to define culture
elements and later apply them to effective therapeutic use.
Currently, there is an ongoing trend towards establishing chemically defined conditions for hPSC culture. Several chemically defined
culture systems have been introduced to maintain hESCs in combination with (i) growth factors/cytokines (e.g., basic fibroblast growth
factor (bFGF), nodal, transforming growth factor-b1 (TGF-b1),
activin A and insulin-like growth factor-1 (IGF-1) analog (heregulin1b)) and (ii) supplements (e.g., GABA, pipecolic acid and lithium
chloride,7,8 and N2/B279) on extracellular matrices (ECM) such as
Matrigel or other ECM components. A chemically defined culture
system with serum/feeder-free conditions is ideal since it excludes the
unknown factors and enhances the reproducibility and robustness of
hPSC propagation. Thus, to facilitate practical applications involving
hPSCs, optimal chemically defined culture conditions must be
established that will not only maintain phenotypically and karyotypically stable cells for extended periods but will also retain the
ability for directed and reproducible differentiation.
Until now, even with the conventional culturing methods,
controlling hPSC fate (e.g., self-renewal, differentiation, apoptosis
and quiescence) has been challenging and underlying mechanisms are
mostly unidentified. However, recent studies have uncovered some
extrinsic factors that can influence state stability of hPSCs and
contribute to fate decisions.10 These extrinsic factors include various
soluble factors, cell-cell interactions, and ECM, which are key
components of the hPSC microenvironment by definition (Fig. 1a).11
Additionally, soluble factors such as growth factors added to the
culture or secreted by stem cells are often potent in their effects on cell
fate.12 Indeed, undifferentiated hPSCs are highly sensitive to the
soluble growth factors that are usually contained in these media.
However, the effects of various defined media for maintaining selfrenewal states over extended periods have not been fully studied and
optimally defined culture conditions have yet to be further refined.
Therefore, screening chemically defined media (CDM) to evaluate the
influence of these factors will also be essential for acquiring more
Lab Chip, 2010, 10, 1113–1119 | 1113
Fig. 1 (a) The extrinsic factors such as soluble growth factors, cell-cell interactions, and ECM play an important role in controlling stem cell fate in the
microenvironment. (b) Schematic illustration of a microfluidic hPSC array for hPSC culture and phenotype assay. (c) Microfluidic image cytometry
(MIC) was conducted followed by segmentation and quantification analysis.
1114 | Lab Chip, 2010, 10, 1113–1119
qualified and defined culture methods that support self-renewal of
hPSCs.
Still, there are several other parameters that must be addressed in
the study of hPSCs. First, although hPSCs can self-renew indefinitely,
it is known that there is enormous variation between different PSC
lines with regard to expression of pluripotency and differentiation
markers.13 This is likely due to that fact that hES cell lines have been
derived from embryos with different characteristics and further isolated by different procedures.14 In the case of hiPSCs, there is also
variation due to (i) the factors used for reprogramming, (ii) the
methods to deliver these factors, (iii) the source of the original cell
lines, (iv) the expression levels of delivered factors, (v) the culture
conditions for obtained hiPSCs and (vi) the methods to identify
obtained hPSCs.15 Second, various commercially available hPSC
defined culture media and ECM7–9,16–18 contain different components
that may cause variable effects depending on the cell line and culture
periods. Thus, taking into consideration all of these parameters as
influential factors, there is also a need to systematically compare the
differences between hPSC lines in order to comprehend their
fundamental biology.
However, there are some disadvantages in conventional experimental settings for hPSCs. Especially, when screening the characteristics among various hPSC lines, conventional analyses such as
flow cytometry, microarray or RT-PCR require large amounts of
cells, resulting in high costs in maintenance.19 On the other hand, the
introduction of microfluidics can allow major advances in stem cell
research. While there are tremendous efforts to compare the similarities and differences of various hPSC characteristics worldwide,13 it
is highly important to establish a standardized hPSC culture condition, which causes less deviation and uncertainty. In microfluidics,
miniaturization of cell culture platforms not only allows us to observe
cellular behavior on the scale found in living systems but also
provides a means to engineer miniaturized cell culture platforms that
are more in vivo-like than conventional dish cultures,20,21 thereby
fostering robust, reproducible and uniform culture conditions.
Additionally, with the ability to manipulate the fluid flow precisely,
microfluidics can make excellent perfusion cell-culture devices, which
are powerful tools to control the soluble and mechanical parameters
of the cell culture environment.22 These aspects are extremely essential
since hPSCs interact strongly with their microenvironmental factors,
which can directly influence the fate decisions. Furthermore, microfluidic technology can be integrated with a variety of biological assays
and is compatible with Micro Electro Mechanical System (MEMS)
technology for further applications including electrophoresis and cell
sorting.19 A microfluidic device is made out of polydimethylsiloxane
(PDMS), which is an elastomeric material utilizing the process of soft
lithography for fabrication. Its beneficial features for stem cell
biology include biocompatibility, gas permeability and durability. It
is also safe and easy to handle within general laboratories performing
biological research. Additionally, with its scalability and automation,
it has more potential for clinical applications. Ultimately, since
microfluidic devices can perform standard tissue culture in a more
rapid, controllable and reproducible fashion with considerably low
costs in a high-throughput fashion,23,24 microfluidic technology is
well-suited for evaluating multiple hPSC culture conditions and
simultaneously observing their responses.
Previously, Villa-Diaz et al. and we have reported maintaining
hESCs in conventional KSR/MEF conditions inside a hESCmChip.25,26 However, to date, it has not yet been reported that hPSCs,
especially hiPSCs, can be cultured in chemically defined culture
conditions and quantitatively studied in a microfluidic device. More
importantly, there has not been a systematic comparison of the
similarities or differences of each of these hPSCs cultured in various
chemically defined culture conditions.
Therefore, we have developed a microfluidic hPSC array (Fig. 1b)
to perform hPSC culture and phenotype assay. Subsequently,
microfluidic image cytometry (MIC) was conducted (Fig. 1c) followed by segmentation and quantification analysis. In this study, we
have reported (i) a microfluidic platform to optimize ECM and
various CDM for evaluating optimal culture conditions in hPSCs,
combined with (ii) a systematic and quantitative analysis and smallscale screening of the hPSCs cultured in various CDM using multiparallel detected protein expressions. Using this array, we have also
performed (iii) a side-by-side comparison of the hPSC phenotypic
responses across available stem cell lines and CDM. This analysis
allowed for examination of the cell fate of a single hPSC in a hPSC
colony in each condition and demonstrated the sensitivity and
effectiveness of our microfluidic hPSC array for use in quantification
of multiple stem cell culture parameters.
For fabrication of a PDMS-based microfluidic hPSC array, we
used the process of soft lithography (ESI Fig. S1a †). The PDMS was
mounted and assembled on a glass slide. During the hPSC culture
assays, this array was set on an inverted microscope stage for routine
monitoring of hPSCs. Our PDMS-based microfluidic hPSC array
was comprised of 24 cell culture chambers (700 mm (W) 900 mm (L)
100 mm (H), Total volume 630 nL). For on-chip cell culture, each
chamber was used for the static culture conditions. Using an electrical
pipette (0.5–12.5 mL, Thermo Fisher Scientific) capable of handling
precise volume and flow rates, four mL of solution containing hPSCs
or reagents were filled into the tip. The tip was gently inserted into the
inlet of a microfluidic hPSC culture array and solution was dispensed
at 6 mL sec1 with accurate piston movement (ESI Fig. S1b†). A few
hours after hPSC loading, the medium was changed every 12 h (see
also in ESI Methods and Fig. S2†). In this array, each chamber can
perform immunocytochemical analysis under discrete hPSC culture
conditions to determine the levels of protein expression (see also in
ESI Methods†). For cell line study, we examined 5 lines including
(i) OCT4-enhanced green fluorescent protein (EGFP) knock-in
HSF1 cell line (HSF1-OCT4-EGFP),25,27 (ii) hESC lines (HSF1 and
H1) and (iii) hiPSC lines (iPSA1 and iPSB2, ESI Fig. S3†). The
OCT4-EGFP cell line is unique in that it allows live cell monitoring of
its pluripotency status in real-time. We therefore used it to optimize
the defined culture conditions. Other cell lines were used to further
make comparisons between their protein expressions.
For the purpose of establishing optimal culture conditions in
a microfluidic hPSC array, we began with examining the optimal
concentration of ECM by using MEF-conditioned medium (CM).
We chose to use hESC qualified Matrigel, (see also in ESI Methods†)
since this is commonly used for feeder-free hPSC culture in current
stem cell research. As mentioned, we used HSF1-OCT4-EGFP cell
lines to monitor the morphology of hPSC colonies and EGFP
expression levels during culturing periods. The results showed that
HSF1-OCT4-EGFP colonies were unable to attach, spread out and
grow well on the substrate coated with 100 mg mL1 (Fig. 2a). On the
other hand, the HSF1-OCT4-EGFP colonies extended well and
maintained their growth in an undifferentiated state for 7 days with
20 mg mL1. Thus, we determined that 20 mg mL1 of Matrigel was an
optimal ECM condition for hPSC culture.
Lab Chip, 2010, 10, 1113–1119 | 1115
Next, using an optimized ECM condition, we then screened CDM
for optimal culture conditions. We tested three CDM (StemPro,16
mTeSR7,8,32 and N2B279), which had been published to support
undifferentiated growth of hPSCs with defined components. Each
medium was also supplemented with varying bFGF concentrations
and the morphology of hPSC colonies and EGFP expression levels
were then monitored over 5 days (Fig. 2b). After five days in culture,
HSF1-OCT4-EGFP cells were able to form colonies and express
EGFP driven by an OCT4 promoter in all three CDM conditions.
We also found that bFGF concentrations did not influence cell
viability and pluripotency of hPSCs. In a previous study, we used
100 ng mL1 of bFGF with KSR/MEF conditions.15 However, at this
time, we observed that 20 ng mL1 bFGF in feeder-free chemically
defined hPSC culture conditions was sufficient to grow undifferentiated hPSCs. Here, we confirmed that all three chemically defined
conditions were able to sustain the growth of hPSCs with undifferentiated states using optimized ECMs and accordingly established
optimal defined culture conditions in a microfluidic hPSC array.
Interestingly, within optimal conditions we now found a variation in
physical and biochemical characteristics in hPSCs cultured with
different media. We then compared the effects of these media on
various phenotypes across the cell lines including morphology,
growth rates and expression level of pluripotency protein markers. A
recent study showed that colony morphology was an important
parameter to determine characteristics of hPSCs and molecular
phenotype and differentiation potential could vary within morphologically different hPSC colonies.28 According to the results of DAPI
nuclear staining, we found that HSF1-OCT4-EGFP cells cultured in
StemPro formed colonies with sharp pointed edges (Fig. 2c). These
cells also had a tendency to form a relatively larger nuclear size than
those cultured in the other CDM. Additionally, HSF1-OCT4-EGFP
cells cultured in mTeSR represented more dense and tight colonies.
We then conducted growth assays to examine the average growth
rate of colonies cultured in each medium by measuring the surface
area of HSF1-OCT4-EGFP colonies (Fig. 2d). We found that
although all the conditions were able to support self-renewal of
hPSCs and maintain pluripotency marker protein expression over
four days, the growth rate of colonies differed depending on culturing
media. Among three CDM, HSF1-OCT4-EGFP cultured in StemPro showed the fastest growth rate. Compared to N2B27, StemPro
and mTeSR conditions showed 2.65 and 1.85-fold changes in their
average colony size, respectively. We speculated that the components
heregulin-1b and activin A were responsible for promoting proliferation of HSF1-OCT4-EGFP cells in StemPro.16
To further characterize the effects of these media on a collection of
hPSC lines, we performed immunocytochemistry to evaluate expression of pluripotency markers in hES and hiPS cells quantitatively
(Fig. 3 and ESI Fig. S2†). The pluripotent markers we used were
OCT4, NANOG, SSEA4, TRA-1-60 and TRA-1-80. Here, we
introduced one more condition where we induced differentiation by
Fig. 2 The establishment of serum/feeder-free chemically defined hPSC
culture conditions in a microfluidic hPSC array. Bright field (BF) and
fluorescence images of hPSCs are shown on the top and bottom,
respectively. (a) Optimization of Matrigel coating conditions. HSF1OCT4-EGFP cells were cultured on ECM with two concentrations (20 or
100 mg mL1) using MEF-CM. Scale bar represents 50 mm. (b) Screening
of CDM (StemPro, mTeSR and N2B27) with different concentrations of
1116 | Lab Chip, 2010, 10, 1113–1119
bFGF using HSF1-OCT4-EGFP cells. HSF1-OCT4-EGFP cells were
cultured on the glass slide coated with an optimal Matrigel concentration
(20 mg mL1). Scale bar represents 100 mm. (c) Morphologically different
HSF1-OCT4-EGFP colonies cultured in three CDM. (d) Quantitative
comparison of the growth curves of HSF1-OCT4-EGFP cells cultured in
three CDM. Each dot represents mean S.D. (*p < 0.05, ***p < 0.001).
Scale bar represents 50 mm.
Fig. 3 Evaluation of pluripotency/differentiation marker expression in
a microfluidic hPSC array using MIC. (a) Bright-field (BF) images, DAPI
nuclear fluorescence images and other fluorescence images of HSF1 cells
cultured in StemPro immunostained against pluripotent markers (OCT4,
NANOG, SSEA4, TRA-1-60 and TRA-1-80). Scale bar represents
50 mm. (b) BF images, DAPI nuclear fluorescence images, OCT4 and
SSEA1 fluorescence images of HSF1 cells cultured in StemPro or
Differentiation condition (Diff). Scale bar represents 50 mm. (c,d) Singlecell based immunofluorescent histograms of (c) OCT4 expression and
(d) SSEA1 expression in individual HSF1, H1, iPSA1 and iPSB2 cells
cultured in KSR/MEF, StemPro, mTeSR, N2B27 and Differentiation
condition. (e,f) Heat maps based on the quantified (e) OCT4 expression
and (f) SSEA1 expression. The protein expression level was normalized
among samples of H1, HSF1, iPSA1 and iPSB2 cultured in KSR/MEF,
StemPro, mTeSR, N2B27 and Differentiation condition and analyzed by
Euclidean distance hierarchical clustering to categorize similar groups
adding 10% fetal bovine serum to DMEM medium as a negative
control. The differentiation marker we used was SSEA1. As the results
indicated, all of the hPSCs cultured in chemically defined conditions
uniformly expressed these pluripotency markers (Fig. 3a). After
confirming their pluripotency, we further quantified OCT4 and
SSEA1 expression at the single-cell level based on immunofluorescence imaging (Fig. 3b). Image cytometry is an image-based
measurement that allows quantitative analysis of these marker
expressions at the single cell level by using software such as CellProfiler, which can generate flow-cytometry-like data. Single-cell based
immunofluorescent histograms presented a variable distribution in
OCT4 and SSEA1 expression (Fig. 3c,d, respectively) after each cell
line in each condition was co-stained with OCT4 and SSEA1 and
analyzed in a single cell. In general, this visually expressed how
heterogeneous/homogeneous each colony was within the undifferentiated and differentiated conditions. As an illustration, when a histogram of protein expression showed the broad distribution,
populations were more heterogeneous and vice versa. The histograms
of OCT4 expression in HSF1, H1 and iPSB2 cultured in KSR/MEF
and three CDM conditions had a homogenous distribution with the
higher level of OCT4 expression compared to Differentiation condition. The iPSA1 cells cultured in mTeSR and N2B27 had two
subpopulations with both high and low OCT4 expression. The
histograms for SSEA1 expression in HSF1, iPSA1 and iPSB2 cultured
in KSR/MEF and three CDM conditions exhibited a homogeneous
distribution with the low SSEA1 expression level, indicating that most
of the hPSC populations remained undifferentiated. On the other
hand, all the cell lines in Differentiation condition exhibited the broad
distribution of the histogram for SSEA1 expression, revealing various
responsiveness and sensitivity of the highly heterogeneous cells upon
the serum inducement. This could also be attributed to the weak and/
or short-time period of treatment, but still enabled visual dynamics of
the protein expression during the differentiation process. Additionally, while the H1 cells cultured in mTeSR and KSR/MEF conditions
had low SSEA1 expression, some populations in StemPro and N2B27
conditions showed relatively high SSEA1 regardless of the fact that
they simultaneously expressed high OCT4.
For systematic analysis, we further conducted Euclidean distance
hierarchical clustering29 based on the mean values of OCT4 and
SSEA1 expression and generated the heat maps, which represented
values of data in two-dimensional maps as colors (Fig. 3e,f, respectively. ESI Fig. S2), to compare the distinct protein expression
resulting from the various cell lines and CDM. Hierarchical clustering
generates a hierarchy of sample groups represented by a dendrogram
(a tree-like diagram). To determine the similarities of two groups, we
used Euclidean distance calculated with the eqn (1),
sffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi
n
X
d ¼ jp-qj ¼
ðpi qi Þ2
(1)
i¼1
if p and q are the two points (here, samples) in Euclidean n-space
for finding nearest neighbors of sample groups. The heat maps
together. Each row represents marker expression in each cell line. Each
column represents a particular condition. (g,h) Evaluation of MIC in
terms of the reproducibility. The quantification of (g) OCT4 fluorescence
intensity and (h) SSEA1 fluorescence intensity between the two experiments is shown.
Lab Chip, 2010, 10, 1113–1119 | 1117
generated based on Euclidean distance hierarchical clustering are
commonly used for, for instance, microarray analysis to reduce data
dimension, categorize samples, and show a multidimensional data set
in 2-D maps in colors. In terms of the different media treatments for 4
days, according to the heat map, hPSCs cultured in StemPro and
mTeSR expressed high OCT4 and low SSEA1 across all the cell lines.
The heat map also showed a similar expression pattern across all the
cell lines in the OCT4 level between hPSCs cultured in StemPro and
mTeSR conditions. In contrast, hPSCs cultured in N2B27 condition
rendered relatively lower OCT4 expression across the cell lines,
exhibiting its tendency to direct differentiation during the culturing
periods. Therefore, N2B27 condition was categorized as similar to the
hPSCs cultured in Differentiation condition based on the clustering.
In the case of SSEA1 expression, all cell lines cultured in Differentiation condition showed strong SSEA1 expression. Additionally, we
observed that some populations of H1 and iPSB2 cells cultured in
N2B27 condition also expressed relatively high SSEA1. In terms of
cell lines within the same condition, each cell line responded differently and resulted in various phenotypes. The iPSB2 line especially
appeared to have different OCT4 and SSEA1 expression compared
to the other three lines. However, there seems to be no clear trend in
cell lines concluded based on the level of pluripotent marker
expression.
Finally, we evaluated the robustness of MIC to confirm the fidelity
of our study (Fig. 3g,h). Two microfluidic chips that cultured H1 cells
in Stem Pro (Exp1 and Exp2) were randomly chosen and both OCT4
(Fig. 3g) and SSEA1 (Fig. 3h) expression were quantified. Between
the two chips, there were no significant differences in OCT4 or
SSEA1 fluorescence intensity value therefore we concluded that our
microfluidic hPSC array in conjunction with MIC was precise and
reproducible.
conventional 6 well plates. According to the results, we found that
culturing in various CDM resulted in different phenotypes in each
hPSC line including morphology, growth rate and pluripotent
marker expression. We speculated that over culturing periods,
heterogeneous cell populations within a single colony showed varied
growth factor responsiveness and protein expressions by intricately
interplaying with the microenvironmental factors at the single cell
level. In general, the final phenotype in a single cell relies on the
current state of the cell and the microenvironment that is composed
of the extrinsic factors such as soluble factors in media and output
signals of hPSCs.11,31 Here, by presenting the detail phenotypic
analysis, we have also demonstrated the ability of our device to study
the heterogeneity of hPSCs and the interaction of different hPSC lines
with the microenvironment, which will have an overall effect in
governing stem cell fate. With a carefully selected set of markers (e.g.
pluripotency, apoptosis, differentiation and cell cycle), this tool can
be applied to conduct more phenotype studies when combined with
signaling cascades transduced by extrinsic factors using its multiplexity to determine the hPSC molecular signatures. Because of these
unique features, we envision that this microfluidic platform will be
beneficial to investigate stem cell biology in a wide range of
biomedical settings and applications in regenerative medicine.
Conclusions
Notes and references
We developed a simple microfluidic platform to optimize ECM,
screen CDM and establish the optimal chemically defined culture
system for both human ESCs and iPSCs. By using this microfluidic
platform, we were also able to study hPSC phenotypic response by
comparing the effects of various CDM and hPSC lines. Although we
cannot ignore the fact that PDMS may absorb molecules from
solution due to their characteristics (e.g., highly porous and hydrophobic material)30 and release them during culturing periods,
according to the results, not only this microfluidic platform can
effectively maintain pluripotency of hPSCs over a week in CDM with
20 ng mL1 of bFGF but all the results were consistent and reproducible across the hPSC lines. Also, our concentration of bFGF was
the original concentration7–9,16 found in other studies with the
conventional macro-scale settings. Thus, we considered this PDMS
effect was negligible. Additionally, we found that the condition with
StemPro medium on the ECM of 20 mg mL1 of Matrigel for
culturing hPSCs generally provides high OCT4 and low SSEA1
expression across the cell lines including hiPSCs. In this work, we
have demonstrated that a microfluidic hPSC array can achieve robust
and reproducible hPSC cultures on a simple setting when combined
with highly quantitative single-cell profiling methods. Furthermore,
not only can this array be utilized for real-time live cell monitoring of
hPSCs, but this platform can also perform small-scale screening with
multi-parallel detection system, using a small amount of samples and
reagents that are roughly 3 orders of magnitude less than the
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This work was supported by the Eli and Edythe Broad Center of
Regenerative Medicine and Stem Cell Research at the Institute of
Molecular Medicine at University of California, Los Angeles.
H.R.T., M.A.T., A.D.P., A.T.C., and O.N.W. were supported by
California Institute of Regenerative Medicine. O.N.W. is an investigator of the Howard Hughes Medical Institute.
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Lab Chip, 2010, 10, 1113–1119 | 1119
Stem cell differentiation
Controlling Differentiation of Neural Stem Cells
Using Extracellular Matrix Protein Patterns
Aniruddh Solanki, Shreyas Shah, Kevin A. Memoli, Sung Young Park,
Seunghun Hong, and Ki-Bum Lee*
The ability of stem cells to differentiate into specialized lineages within a specific microenvironment is vital for
regenerative medicine. For harnessing the full potential of
stem cells for regenerative therapies, it is important to investigate and understand the function of three types of microenvironmental cues—soluble signals, cell–cell interactions,
and insoluble (physical) signals—that dynamically regulate
stem cell differentiation.[1] Neural stem cells (NSCs) are
multipotent and differentiate into neurons and glial cells,[2]
which can provide essential sources of engraftable neural
cells for devastating diseases such as Alzheimer’s disease,[3]
Parkinson’s disease[4] and spinal cord injury.[5] One of the
major challenges involved in the differentiation of NSCs is
to identify and optimize factors which result in an increased
proportion of NSCs differentiating into neurons as opposed
to glial cells. To this end, soluble cues such as brain-derived
neurotrophic factor (BDNF),[6] sonic hedgehog (Shh),[7]
retinoic acid (RA),[6c] and neuropathiazol[8] have been
shown to significantly increase neuronal differentiation of
NSCs in vitro. However, the research toward studying the
function of the other two microenvironmental cues (cell–
cell interactions and insoluble cues) during the neurodifferentiation of NSCs is limited, mainly due to the lack
of availability of methods for the investigation.[9] While
various aspects such as cell–cell interactions,[10] combinations of extracellular matrix (ECM) proteins,[1a,11] and
physical properties of substrates have been shown to play
a vital role in determining the fate of other adult stem cells
such as mesenchymal stem cells (MSCs),[12] cardiac stem
A. Solanki, S. Shah, K. A. Memoli, Prof. K.-B. Lee
Department of Chemistry and Chemical Biology, Rutgers
The State University of New Jersey
Email: kblee@rutgers.edu, http://rutchem.rutgers.edu/∼kbleeweb/
S. Y. Park
Interdisciplinary Program in Nano-Science and Technology
Seoul National University
Seoul, South Korea
Prof. S. Hong
Department of Physics
Department of Biophysics and Chemical Biology
Seoul, South Korea
DOI: 10.1002/smll.201001341
small 2010, 6, No. 22, 2509–2513
cells,[13] and hematopoetic stem cells,[14] little is known
about the influence of such factors on the neuronal differentiation of NSCs. Therefore, there is a pressing need
to develop methods for investigating the role of cell–cell
interactions and insoluble signals in selectively inducing the
differentiation of NSCs into specific neural cell lineages.
Herein, we demonstrate how ECM protein patterns
can be used to investigate the effect of physical cues combined with cell–cell interactions on the differentiation of
NSCs. Bio-surface chemistry combined with soft lithography was used to generate combinatorial patterns with
varying geometries and dimensions of ECM proteins (e.g.,
laminin, fibronectin, and collagens) to study the influence
of surface features and ECM compositions on the differentiation of NSCs. We hypothesized that the ECM protein
patterns with variant geometries and dimensions would
provide physical cues (e.g., mechanical or topographical
cues), as well as guide cell–cell and cell–ECM interactions
in a controlled manner, both of which would ultimately lead
to a pattern geometry-dependent and pattern dimensiondependent neuronal and glial differentiation (Figure 1).
Our data confirmed that the difference in the extent of neuronal and glial differentiation of NSCs on the ECM protein
patterns was entirely due to the pattern geometry and dimension, as all the experiments were carried out in the absence of
exogenous factors that promote neurogenesis; this suggests
that NSCs can undergo differentiation by purely sensing the
difference in ECM pattern geometries and dimensions.
Extracellular matrix protein patterns with variant
geometries and dimensions were fabricated by initially patterning octadecanethiol (ODT, 5 mm in ethanol), a hydrophobic alkanethiol, which formed self-assembled monolayers
(SAMs) of squares, stripes, and grids on glass substrates
coated with a thin film (12 nm) of gold. In order to minimize
the nonspecific attachment of laminin, the background of the
substrates was passivated by incubating in a solution (5 mm
in ethanol) of tetraethylene glycol terminated alkanethiol
[EG4-(CH2)11-SH, 12 h] (See Supporting Information for
synthesis and characterization). After passivating the background, a solution of ECM protein [e.g., laminin (10 μg mL−1)
in phosphate buffered saline (PBS) buffer, pH = 7.4] was
added onto the substrates (3 h) and was preferentially
adsorbed onto the hydrophobic regions (ODT patterns).
The selective adsorption of laminin on hydrophobic regions
was consistent with the results of other groups[15] and was
also confirmed by immunostaining using anti-laminin IgG
© 2010 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
wileyonlinelibrary.com
2509
communications
A. Solanki et al.
interactions of NSCs with the passivated
areas, and then incubated in fresh basal
medium. The media was exchanged with
pattern geometry
fresh media every other day. During our
screening approach to investigate the
squares
stripes
grids
function of physical cues on neuronal difH H H H H H
O O O O O O
O O O O O O
ferentiation of NSCs, we monitored the
O O O O O O
differentiation on ECM protein patterns
O O O O O O
O O O O O O
by using two orthogonal assays, namely
immunocytochemical and morphological
assays. To assess the differentiation of
S S S S S S S S S S S S
NSCs, the down-regulation of the NSC
Au Thin Film
marker (Nestin) and the geometryECM
Protein
ECM Protein Patterns
dependent expression of the neuronal
marker (β-III Tubulin, TuJ1) and glial
marker (glial fibrillary acidic protein,
GFAP) were monitored. In addition, the
development of branches or spindle-like
morphologies, and neurite outgrowths
were observed by using an inverted phase
contrast microscope (Zeiss Axiovert
squares
stripes
grids
200M equipped with AxioCam CCD).
Patterns of ECM proteins with different geometries contributing to adheFigure 1. A schematic diagram of our approaches. A) The fabrication of ECM protein patterns sion, proliferation, growth and migration
and their application for NSC differentiation. B) The selective attachment of NSCs on the protein
of various cells (including stem cells) have
patterns and differentation into two different kinds of neural cells. C) The differentiation of
been reported.[16] In addition, reports from
NSCs into either neurons (red) or astrocytes (green) on the protein patterns. D) Increased
the literature have shown cell–cell interacneuronal differentiation on the grid patterns, as compared to the stripes and squares.
tions to play a critical role in the differentiation of adult stem cells. For instance,
(See Supporting Information, Figure S1). Only the patterned it was recently shown that cell–cell interactions played an
regions, coated with ECM proteins, promoted cell adhesion important role in the osteogenic (bone) differentiation of
and growth whereas the rest of the substrate remained inert MSCs.[10] To study the influences pattern geometries and cell–
(Figure 1). We similarly patterned several different ECM pro- cell interactions on the differentiation of NSCs, we initially
teins including fibronectin and collagen, but found that lam- patterned the NSCs on stripes of laminin, which promoted
inin provided the optimum microenviromental cues for NSC one-way interactions in a controlled manner (Figure 2.A1).
adhesion and growth. Hence, all our differentiation studies We found that after six days, 36% of NSCs on the isolated
were carried out using laminin patterns.
stripes differentiated into neurons (Figure 2.A2 and Figure 3).
To examine the effect of the ECM protein patterns on At the same time we observed that 64.3% of NSCs on
stem cell differentiation, primary rat hippocampal neural these stripes differentiated into astrocytes (Figure 2.A3 and
stem cells (Millipore) were first expanded and maintained Figure 3).
in an undifferentiated state in a homogeneous monolayer on
To further confirm the influence of such interactions
a polyornithine and laminin-coated Petri dish in a defined on the differentiation of NSCs, we used square patterns of
serum-free growth medium [DMEM/F12 supplemented with laminin to isolate NSCs and restrict their growth within the
B27 and basic fibroblast growth factor (bFGF, 20 ng mL−1)]. square patterns (Figure 2.B1). We hypothesized that the difFor obtaining reproducible and consistent results, all ferentiation behaviour of NSCs can be considerably influexperiments were carried out using NSCs from passages enced by limiting cell–cell interactions. We observed that
2–5 at a constant cell density of 150 000 cells per substrate NSCs patterned on squares, having the same dimensions and
(1.5 cm × 1.5 cm), which was optimum for cell growth spaces as the stripes, differentiated into neurons to a considwithout clustering. Arresting the proliferation of NSCs and erably lesser extent (28.1%, Figure 2.B2 and 3) as compared
initiating their spontaneous differentiation was achieved to the NSCs involved in one-way interactions on the striped
by withdrawing bFGF from the culture medium (resulting laminin patterns. At the same time, the number of NSCs that
in basal medium), without the additional treatment with differentiated into astrocytes increased considerably on
exogenous factors (proteins and small molecules). The basal squares –76.9% on squares as compared to 64.3% on stripes
medium (2 mL) containing the NSCs (75 000 cells mL−1) (Figure 2.B3 and 3). Thus, the reduced cell–cell interactions
was put in a single well of a 6-well plate containing a sub- with the NSCs on the surrounding patterns may have led to
strate with laminin patterns. After the NSCs attached onto reduced neuronal differentiation and increased glial differenthe laminin patterns (1 h), the substrates were rinsed with tiation of the NSCs. Based on the observed differentiation of
copious amounts of media in order to minimize nonspecific NSCs on stripes and squares, we further hypothesized that using
neurons
B
pa
tte
rn
di
m
en
sio
n
A
astrocytes
C
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Figure 2. Growth and differentiation of NSCs on the laminin patterns.
Phase contrast images show NSC attachment and growth on stripes
(A1), squares (B1), and grids (C1) on Day 2 after seeding. Fluorescent
images of cells stained for the neuronal marker TuJ1 (red) and nucleus
(blue) show the extent of neuronal differentiation of NSCs on stripes
(A2), squares (B2), and grids (C2) on Day 6 after seeding. Similarly, cells
stained for astrocyte marker GFAP (green) and nucleus (blue) show the
extent of glial differentiation on stripes (A3), squares (B3), and grids
(C3) on Day 6 after seeding. Scale bars: 50 μm.
specific pattern geometries promoting cell–cell interactions
could lead to higher neuronal differentiation. For this purpose,
we used grid patterns of laminin, having the same dimensions as the stripe and square patterns, for NSC growth and
differentiation. The grid patterns were specifically designed to
increase cell–cell interactions in a controlled manner (by promoting two-way interactions, Figure 2.C1). After six days in
basal medium, as compared to the NSCs patterned on stripes
and squares of laminin, we observed a remarkable increase
in the number of NSCs that underwent neuronal differentiation (45.6%, Figure 2.C2 and 3) and a decrease in the number
of cells that underwent glial differentiation on grid patterns
of laminin (49.6%, Figure 2.C3 and 3). All the experiments
were repeated several times under the same conditions. To
maintain consistency and minimize the effects from other
variables, we fabricated and used PDMS stamps to generate
ECM protein patterns of all the three geometries (having the
same dimensions and spacing) on the same substrate. Using
this method, we could reproduce and confirm our results
with relative ease. Neuronal and glial differentiation of NSCs
was also monitored on control substrates which included
substrates coated with laminin (unpatterned) and substrates
without laminin. The NSCs on substrates without laminin did
not attach and failed to survive, whereas 32.5% of the NSCs
on the unpatterned substrates coated with laminin differentiated into neurons and 71.2% of the NSCs differentiated into
astrocytes six days after seeding.
In addition to investigating the effect of pattern-geometry,
we also studied the effect of dimensions on NSC differentiation. To this end, we generated ten different dimensions for
each of the geometries, ranging from sizes as small as 10 μm
and as large as 250 μm (Figure 4B). Interestingly, for the
three different geometries above 50 μm, we observed little
difference in the percentage of NSCs undergoing neuronal
% of Cells Expressing Neural Markers
90
TuJ1 (Neurons)
80
70
60
50
40
30
20
10
0
Squares
Stripes
Grids
Pattern Geometry (10-50 µm)
Figure 3. Quantitative comparison of the percentage of cells expressing
the neuronal marker TuJ1 and astrocyte marker GFAP on laminin patterns
of squares, stripes and grids. Six days after seeding, the differentiated
cells were counted and plotted as a ratio of TuJ1-positive cells or GFAPpositive cells to the total number of cells (n = 3). Student’s unpaired
t-test was used for evaluating the statistical significance for cells
stained for TuJ1 on stripes and squares, compared to those on grids.
(∗ = P < 0.01, ∗∗ = P < 0.001).
small 2010, 6, No. 22, 2509–2513
Figure 4. NSC alignment and differentiation on combinatorial ECM
patterns. A) NSCs on grids of laminin express the neural stem cell
marker, nestin (purple) on Day 2 after seeding, thus confirming that
the NSCs are undifferentiated. B) NSCs stained for actin (green) show
extensive spreading and cell–cell interactions on grid patterns of
laminin on Day 2 after seeding, confirming that the NSCs, while still in
the undifferentiated state, extensively interact with each other. C) SEM
image of NSCs on Day 2 after seeding, showing the early alignment and
extension of processes on grid patterns of laminin. D) NSCs previously
shown to extend and grow on the grid patterns of laminin undergo
neuronal differentiation and express the neuronal marker synapsin
(pseudocolored yellow) on Day 6 after seeding. Scale bars: 20 μm.
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A. Solanki et al.
and glial differentiation. The result observed for pattern
dimensions above 50 μm was similar to that observed with
unpatterned substrates. We believe the cells may not be able
to sense the difference in pattern geometries above 50 μm
and thus show similar behaviour to the cells on unpatterned
substrates. Since the NSCs showed remarkable difference in
differentiation on patterns ranging from 10–50 μm, all of our
statistical analysis and investigation was done using pattern
features within this range.
We observed that the laminin patterns of all three
geometries enabled the NSCs to attach and grow within
a day or two day after seeding. By staining for actin using
phalloidin and using field emission scanning electron microscopy (FESEM, Zeiss Gemini), we further observed that
the cytoskeleton of the NSCs aligned well within the laminin patterns, guiding cellular morphology and interactions
(Figure 4B,C). To confirm that the laminin patterns influenced
morphological changes before differentiation (as opposed to
an early differentiation of NSCs which might have caused
a change in alignment and morphology), the NSCs were
immunostained for the neural stem cell marker nestin two
days after seeding in basal medium. We observed that most
of the NSCs that aligned along the patterns, stained positive
for nestin (Figure 4A), confirming that cells were in an undifferentiated (multipotent) state when they aligned along the
patterns (See Supporting information, Figure S2 for NSCs on
squares and stripes stained for actin and nestin). We further
confirmed neuronal differentiation of NSCs on the laminin
patterns using synapsin as another neuronal marker in addition to TuJ1. After six days in basal medium, a remarkably
high number of the NSCs growing along the grid patterns of
laminin expressed synapsin (Figure 4D). In addition, colocalization of TuJ1 and synapsin was observed within the NSCs
differentiated on the grid patterns, confirming that the neurons expressed both neuronal markers (Supporting information, Figure S3).
In summary, we fabricated and utilized patterns of ECM
proteins for modulating the extent of neuronal and glial differentiation of NSCs in the absence of soluble cues such as
small molecules and exogenous proteins. Potentially, our
approach and methodology can be helpful for deconvoluting
physical cues and cell–cell interactions from complex microenvironmental cues. More detailed mechanistic studies on
how physical cues modulate the signaling cascades and the
signaling pathways that are primarily involved in stem cell
differentiation induced by such factors are currently under
investigation. The implications of our results could also
potentially be significant for tissue engineering for brain
and spinal cord injuries, where NSCs or NSC-based differentiated cells can be transplanted into the damaged regions
with scaffolds. For example, scaffolds having patterns promoting cell–cell interactions in a controlled manner could
potentially lead to increased neuronal differentiation in vivo.
Even though we have explored only proof-of-concept experiments focusing on differentiation of NSCs, a similar strategy
could be extended to study and control the fate of other
stem cells, such as MSCs and embryonic stem cells (work in
progress). Our results substantiate the importance of pattern
2512 www.small-journal.com
dimensions, pattern geometries, and cell–cell interactions in
controlling stem cell fate.
Supporting Information
Supporting Information is available from the Wiley Online Library
or from the author.
Acknowledgements
A. S. and S. S. have contributed equally to the authorship of this
paper. We acknowledge John Kim and Neal Patel for their help in
the experimental procedures. This work was supported by the NIH
Director’s Innovator Award [(1DP20D006462–01), K.-B. L.] and the
N.J. Commission on Spinal Cord grant [(09–3085-SCR-E-0), K.-B. L.].
S. H. acknowledges the support from the NRF grant (2009–
0079103) and the System 2010 program. K.-B. L. acknowledges
partial support from Bioforce Nanosciences.
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Received: August 4, 2010
Published online: September 21, 2010
2513
Biomedical Applications
Carbon Nanotube Monolayer Cues for Osteogenesis
of Mesenchymal Stem Cells
Ku Youn Baik, Sung Young Park, Kwang Heo, Ki-Bum Lee, and Seunghun Hong*
Recent advances in nanotechnology present synthetic bioinspired materials to create new controllable microenvironments for stem cell growth, which have allowed directed
differentiation into specific lineages.[1,2] Carbon nanotubes
(CNTs), one of the most extensively studied nanomaterials,
can provide a favorable extracellular environment for intimate
cell adhesion due to their similar dimension to collagen. It has
been shown that CNTs support the attachment and growth of
adult stem cells[3–6] and progenitor cells including osteoblasts
and myoblasts.[7,8] In addition, surface-functionalized CNTs
provide new opportunities in controlling cell growth. Surface
functionalization improves the attachment of biomolecules,
such as proteins, DNA, and aptamers, to CNTs.[9] Zanello
et al. cultured osteoblasts on CNTs with various functional
groups and showed reduced cell growth on positively charged
CNTs.[10] Recent reports have shown that human mesenchymal stem cells (hMSCs) formed focal adhesions and grew
well on single-walled CNTs (swCNTs).[5,6] However, the effect
of naïve swCNT substrates on the differentiation of stem cells
has not been reported before. Herein, we report the osteogenic differentiation of hMSCs induced by swCNT monolayer cues without any chemical treatments. Interestingly, the
surface treatment of swCNTs via oxygen plasma showed synergistic effects on the differentiation as well as the adhesion
of hMSCs. The stress due to the enhanced cell spreading on
swCNT layers was proposed as a possible explanation for the
Dr. K. Y. Baik, Prof. S. Hong
Department of Physics and Astronomy
Seoul, 151–747, Korea
Dr. K. Y. Baik
Plasma Bioscience Research Center
Kwangwoon University
S. Y. Park, K. Heo, Prof. S. Hong
Prof. K.-B. Lee
Department of Chemistry and Chemical Biology, Rutgers
NJ 08854, USA
Prof. S. Hong
Department of Biophysics and Chemical Biology (WCU Program)
E-mail: seunghun@snu.ac.kr
DOI: 10.1002/smll.201001930
small 2011, 7, No. 6, 741–745
enhanced osteogenesis of hMSCs on the swCNT monolayers.
Previous reports showed that the stress to stretch stem cells on
microscale molecular patterns generated the tension on actin
filaments, which eventually enhanced the osteogenesis.[11,12]
Since our method relies on monolayer coating of swCNTs, it
can be applied to a wide range of substrates including conventional scaffolds without any complicated fabrication processes.
Figure 1 shows a schematic diagram depicting our experimental procedure. In this experiment, three substrates were
used: glass as a control, a pristine swCNT monolayer adsorbed
on a glass surface, and an oxygen-plasma-treated swCNT
(O-swCNT) monolayer on a glass surface (Figure S1a in the
Supporting Information (SI)).[13] swCNTs had the average
diameter of approximately 1–2 nm, and their average length
was 1.5 μm. When the cleaned cover slips were dipped in a dispersed swCNT solution, swCNTs were adsorbed onto the glass
surface to form a monolayer. Oxygen plasma treatment was
performed to modify the surface chemistry of swCNTs, which
is known to generate hydroxyl or carboxyl groups on the surface.[14,15] Atomic force microscopy (AFM) analysis revealed
that the swCNT layer maintained its surface roughness even
after 40 s of the oxygen plasma treatment, while its contact
angle decreased abruptly after 20 s of exposure to oxygen
plasma (Figure S1b in the SI). Our control experiments show
that, as the time for plasma treatment increased, the abrupt
change of contact angle or surface roughness occurred at around
20 or 40 s, respectively. Due to such an abrupt transition, the
properties of swCNT layers treated with oxygen plasma for
20 or 40 s usually exhibited rather large variations, which
reduced the reproducibility of the following stem cell growth
experiments. On the other hand, we could reproducibly obtain
similar surface properties for the swCNT layers after 30 s of
plasma treatment, thus enabling reliable stem cell growth experiments. Consequently, in our experiment, we utilized swCNT
monolayers treated with oxygen plasma for 30 s, which reproducibly exhibited hydrophilic properties while maintaining the
roughness of pristine swCNT monolayers. The hMSCs from
bone marrow were seeded on these substrates, and then their
adhesion, proliferation, and differentiation were examined.
Figure 2 shows the adhesion and proliferation of hMSCs
cultured on glass (Figure 2a), a swCNT monolayer (Figure 2b),
and an O-swCNT monolayer (Figure 2c). Their actin filaments and nuclei were visualized after 24 h from seeding. It
is notable that hMSCs spread wider, and their actin fibers
look thicker on swCNT monolayers than on glass substrates.
For quantitative analysis of cell adhesion, the area of 200
individual cells on each substrate was estimated from that of
stained actin fibers (see Figure S2 in the SI) and utilized to
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K. Y. Baik et al.
in hMSCs cultured on an innate swCNT film.[6] Other
researchers also have observed wider adhesion of various
cells on swCNT substrates than on conventional substrates
such as glass. Osteoblast-like cells and hMSCs spread better
on a swCNT film than on glass substrates, resulting in the
larger cell area and higher occurrence of filopodia at the cell
boundaries.[6,17]
Besides the effects of nanoscale surface roughness,[16]
our results also indicate that surface chemistry plays a role
in cellular interaction between cells and swCNTs. Note
that even though our swCNT monolayers with or without
oxygen plasma treatment had similar surface roughness
Figure 1. Schematic diagram depicting hMSC growth on swCNT
(Figure S1 in the SI), hMSCs on O-swCNT substrates exhibmonolayers. a) Glass substrate was used as a control. b) swCNTs
were adsorbed onto the glass substrate to form a swCNT monolayer. ited an enhanced cell area and proliferation compared with
c) Oxygen plasma treatment was applied to modulate the swCNT surface those on pristine swCNT substrates (Figure 2). Presumably,
the chemical changes of the O-swCNT layer, such as enhanced
properties.
hydrophilicity and surface oxygen content, increased the procalculate the average area per cell (Figure 2d). The results liferation and adhesion of hMSCs on it.[18]
The morphological change of hMSCs has been reported to
show that the adherent area of individual hMSCs is higher on
swCNT or O-swCNT monolayers compared with that on the be related to their capacity for multipotentiality. For example,
glass substrate. At the same time, the ratio of long and short spindle-shaped hMSCs have high potential for adipogenesis,
axial lengths of the hMSCs is smaller on swCNT monolayers while flat hMSCs have high potential for osteogenesis.[19,20]
and much smaller on O-swCNT monolayers than on glass Similarly, the actin cytoskeleton changed from thin and parsubstrates (Figure 2e). The MTS (3-(4,5-dimethylthiazol-2- allel microfilament bundles to thick and crisscrossed bundles
yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetra- under the osteogenic differentiation process, which is conzolium) assay results show that the proliferation of hMSCs sistent with Figure 2b,c.[21,22] This implies that the swCNT
substrates could enhance the osteogenesis of hMSCs.
was enhanced on the O-swCNT substrate (Figure 2f).
In order to check the osteogenic induction by the swCNT
One possible explanation regarding the enhanced cell
area on the swCNT substrate can be the nanoscale rough- monolayer, the osteogenic proteins and corresponding genes
ness of the swCNT monolayers. Zhang et al. reported that were tested. Core binding factor alpha1 (CBFA1), osteocalcin
nanoscale surface roughness of swCNTs can deform the cell (OCN), and alkaline phosphatase (ALP) were used as ostemembrane and hinder the motion of vesicles inside cells.[16] ogenic markers. CBFA1 is the main transcription factor for
This membrane deformation may affect the distribution committing hMSCs to the osteoblastic lineage; OCN is a difand diffusion of membrane proteins including focal adhe- ferentiated osteoblast-specific gene for mineralizing the bone
sion proteins, which are critical factors in cell adhesion. Tay matrix, and ALP is an early osteoblastic marker.[23,24] hMSCs
et al. showed increased number of focal adhesion proteins were cultured on three different substrates with and without
osteogenic induction media for 17 days.
Immunostaining was performed at day
12, and the quantitative gene analysis was
performed at day 7 and 14 from seeding.
Figure 3a shows that the hMSCs filled
the whole area on all substrates at day
12. Also, note that the cells cultured in
osteogenic induction media were slightly
detached from the glass due to strong
cell–cell interactions. The detachment was
retarded on swCNT substrates (data not
shown). The osteogenic protein OCN was
detected and visualized with fluorescent
dyes (Figure 3b). The hMSCs grown on
swCNT substrates exhibited brighter
OCN immunofluorescence than those on
glass substrates, but it was not statistically
Figure 2. Adhesion and proliferation of hMSCs on various substrates. Fluorescence images significant (Figure S3 in the SI). However,
of actin filaments show the morphology of hMSCs on a) a glass substrate, b) a swCNT
the hMSCs grown on O-swCNT substrates
monolayer, and c) an oxygen-plasma-treated swCNT monolayer (O-swCNT). Scale bars are
definitely exhibited brighter immunofluo100 μm. Quantitative analysis was visualized with d) the averaged value of area per cell
(number of cells, n = 200), e) the averaged value of the ratio of long and short axial lengths rescence than those on glass substrates,
(a/b in Figure 2b; n = 200), and f) the averaged MTS assay value at day 6 (n = 3). In all and quantitative statistical analysis showed
analyses, the student’s t-test was utilized for the significance calculation (∗: p < 0.05).
much more brightness on O-swCNT
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Carbon Nanotube Monolayer Cues for Osteogenesis of Mesenchymal Stem Cells
Figure 3. Immunohistochemistry of an osteogenic protein and mRNA analysis of hMSCs on glass substrates, swCNT substrates, and O-swCNT
substrates with normal culture media; and hMSCs cultured on glass substrates with osteogenic induction media. Scale bars are 100 μm. a) Brightfield images, b) osteocalcin (OCN) protein immunostaining at day 12, c) quantitative polymerase chain reaction (qPCR) data of mRNA extracted from
hMSCs on each substrate at day 7 and 14. An early marker of osteogenic commitment (CBFA1), a late marker of developing osteoblasts (OCN), and
an early osteoblastic marker (ALP) were tested (n = 5). The results indicate the enhanced osteogenic differentiation of hMSCs on swCNT monolayer
and on an O-swCNT monolayer without any differentiation-inducing chemicals.
substrates. This high protein expression was confirmed in
mRNA expression. The mRNA of beta-actin was incorporated as an endogenous housekeeping gene for all the test
substrates, and the relative transcription level to that of
the hMSCs cultured on glass is shown in Figure 3c. After
14 days of culture, the osteogenic genes such as CBFA1 and
OCN were upregulated on both swCNT and O-swCNT substrates.[24] In the case of ALP, the expression was enhanced
only on the O-swCNT substrates in the early days of culture
(Figure 3c; Figure S4 in the SI).
The brighter fluorescence and enhanced mRNA expression indicate the enhanced commitment of hMSCs on
O-swCNT monolayers for osteogenesis without any differentiation-inducing chemicals. Although the enhanced osteogenic function of osteoblast on CNTs have been reported
previously,[25,26] this is the first report regarding the enhanced
commitment of hMSCs to osteoblast lineage on a CNT
monolayer.
A possible explanation for the enhanced osteogenesis can
be the stress on cells due to the enhanced cell spreading on the
swCNT monolayers. It was reported that the stress to stretch a
stem cell generated the tension on actin filaments and eventually enhanced the osteogenesis.[11,12,27] In order to verify this
hypothesis, we fabricated differently sized swCNT square patterns. The size of the patterns was determined considering a
previous report utilizing 12 μm extracellular matrix (ECM)
molecular patterns to enhance adipogenesis and 100 μm
small 2011, 7, No. 6, 741–745
patterns to osteogenesis.[11] We fabricated patterns larger than
100 μm to see only the effect of enhanced spreading on osteogenesis. Figure 4a shows the bright field images of hMSCs
cultured for 12 days. The hydrophobic molecules efficiently
blocked the hMSC adhesion, resulting in the confined growth
of hMSCs only in the swCNT regions.[3] A closer look at
cell morphologies showed that the nuclei were located at
the center of the pattern, and cytoplasm was extended to the
boundaries. Actin fibers were anchored and stretched to the
focal adhesions located at the boundaries (Figure S5 in the SI).
This indicates that the hMSCs tended to spread out to the
boundaries of swCNT patterns using focal adhesion, and thus
actin fibers were stretched as the square size increased.
The DAPI stained nuclei count 2, 13, 17, and 17 for 100,
200, 300, and 400 μm square patterns, respectively (Figure 4b).
The spreading area per cell was calculated by dividing the
actin stained area by the cell number. Data from 20 square
swCNT patterns were plotted in Figure 4d. As the pattern
size increased, the cell density decreased and the averaged
area per cell increased. This implies that the hMSCs in larger
swCNT patterns were spread wider. The OCN immunofluorescence value per single cell was averaged from nine of each
swCNT pattern size (Figure 4c), and the values were plotted
in Figure 4e. The results indicate that hMSCs on larger square
patterns exhibited easier commitment to the osteogenic
lineage. This is consistent with our hypothesis that larger cell
spreading in swCNT patterns enhanced the osteogenesis.
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K. Y. Baik et al.
differentiation-inducing media. In addition, we found that simple oxygen plasma
treatment amplifies the adhesion, proliferation, and even osteogenic differentiation
of hMSCs by adding chemical effects
on the main topographical effects. The
stress on hMSCs due to the enhanced cell
spreading on swCNT monolayers was proposed as a possible explanation. However,
a more detailed mechanism, involving such
details as the signal pathways in the stem
cells, is not yet clear and requires further
study. In any case, this work suggests that
the swCNTs can be a powerful scaffold in
mesenchymal stem cell engineering.
Experimental Section
Preparation of CNT Substrates: swCNTs
in powdered form (Carbon Nanotechnologies, Inc.) were sonicated in dichlorobenzene
(0.05 mg mL−1), and cleaned glass cover slips
(piranha solution; H2SO4:H2O2 = 3:1) were
dipped in dispersed swCNT solution so that
swCNTs were adsorbed onto the glass surface.
In this case, the first adsorbed swCNTs blocked
the additional adsorption of swCNTs, resulting
in monolayer coverage with a certain maximum
density.[28] After 2 min, the swCNT-coated glass
Figure 4. Immunohistochemistry and fluorescence quantification of confined hMSCs in was rinsed with dichlorobenzene vigorously
differently sized square patterns of swCNT monolayers. Scale bars are 100 μm. a) Bright-field
to remove any weakly adsorbed swCNTs, and
images of hMSCs on the square patterns of a swCNT monolayer after 12 days. b) Fluorescence
images of nuclei stained with 4’,6-diamidino-2-phenylindole (DAPI). The panels show 2, 13, then dried with N2 gas. Oxygen plasma treat17, or 17 cells in 100, 200, 300, or 400 μm sized square patterns, respectively. c) Fluorescence ments were performed to modify the surface
images of OCN immunostaining, indicating osteogenic differentiation. d) Area per cell in chemistry of the swCNTs (Expanded Plasma
differently sized square patterns. Measurements were averaged over 20 squares for each Cleaner (PDC-002) from Harrick Plasma,
size. e) Intensity of OCN immunostaining per cell in differently sized square patterns. Values radio frequency (RF) power ≈ 30 W, pressure
were averaged over 9 squares for each size. The results indicate that enhanced osteogenesis
≈ 120 mTorr; 1 mTorr = 0.133 Pa).
is accompanied with enlarged cell area.
Cell Culture and Reagents: hMSCs from
human bone marrow (purchased from
As previously reported, the enhanced adhesion might be Lonza, Walkersville, USA) were expanded in MSC growth medium
linked to Rho-family GTPase (guanine triphosphatase) sign- (MSCGM) and used for our experiments at passage 4–6 in culaling and nonmuscle myosin contraction within the cell. They ture medium (high-glucose Dulbecco’s Modified Eagle Medium
showed that the overexpression of ras homolog gene family (Gibco) + 10% fetal bovine serum (FBS; Gibco) + 1% penicillinmember A (RhoA) or Rho-associated coiled-coil-containing streptomycin (Gibco)). All the substrates were cleaned with 70%
protein kinase 1 (Rock1) stimulated myosin contraction and ethanol and phosphate buffered saline (PBS) to remove residual
promoted osteogenesis.[11] Microarray analysis and pathway toxic solvents. The cells were seeded with a density of about
inhibition studies should be followed to elucidate a more 3000 cells cm−2 on the prepared substrates, and culture media
detailed mechanism. Furthermore, the effect of plasma was changed every 2–3 days. For osteogenic differentiation,
treatment on swCNT should be studied systemically. As the hMSCs were cultured in osteogenic differentiation media (100 nM
plasma treatment was known to enhance protein adsorption dexamethasone, 50 μM ascorbic acid, and 10 mM glycerol 2onto swCNT surfaces, it might have an influence on the cel- phosphate in culture medium).[11]
Immunohistochemistry: To stain actin fibers, cells were fixed in
lular interaction with the nanostructured surface. Interestingly, surface chemical modulation with –OH and –COOH 4% formaldehyde solution, permeabilized with 0.1% Triton X-100,
groups itself was reported to downregulate the osteogen- and then stained by tetramethylrhodamine isothiocyanate (TRITC)esis.[28] This implies that there is a synergistic effect of both conjugated phalloidin (1:100, Molecular Probes). For osteocalcin
staining, cells were fixed with 4% formaldehyde solution in PBS, persurface roughness and chemistry on O-swCNT substrates.
In summary, we showed that the osteogenic differentiation meabilized with 0.1% Triton X-100 in PBS, blocked with 10% normal
of hMSC was promoted from swCNT monolayers without goat serum for 1 h at room temperature, and incubated with mouse
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small 2011, 7, No. 6, 741–745
Carbon Nanotube Monolayer Cues for Osteogenesis of Mesenchymal Stem Cells
anti-human osteocalcin IgG (1:100 dilution, 50114, QED Bioscience
Inc.) for 1 h at room temperature. The second antibody (1:500 dilution, Alexa-Fluor 488 conjugated anti-mouse IgG, Sigma) was then
adhered for immunofluorescence. For vinculin staining, monoclonal
anti-vinculin antibody produced in mouse (1:100, Sigma) was used.
After counterstaining the nuclei with DAPI (prolong gold antifade
reagent with DAPI, Invitrogen), fluorescence images were obtained
using a Nikon Eclipse TE2000-U microscope and a complementary
metal-oxide semiconductor camera (INFINITY1–1C, Lumenera Corp.).
Cell Proliferation Test: The cells were seeded at the same density, and the number of cells on each substrate was determined
using CellTiter 96 Aqueous One Solution Cell Proliferation Assay
(G3580, Promega) and spectrometer (HP845, Hewlett-Packard).
Cell Area Measurement: Actin filaments of cells were stained
with phalloidin. The fluorescence signal was converted to the
values of ‘0’ or ‘1’ (black and white) by subtracting the averaged
background value. The number of pixels whose value is ‘1’ was
counted to calculate the area of each cell. The area of 200 cells for
each case was measured.
qPCR Analysis: Total RNA was extracted from hMSCs using
RNeasy Mini Kit (74104, Qiagen), and converted to cDNA using
reverse transcriptase and random primers (ImProm-II Reverse Transcription System, Promega). The same amount of total RNA was
used in cDNA synthesis. Resulting cDNAs was used in qPCR (7300
Real Time PCR system, Applied Biosystems) with the primers
for β-Actin (NM_001101.3), CBFA1 (NM_001015051), OCN
(NM_199173.3), and ALP (NM_000478.3).
Preparation of Micropatterned Substrates: Micropatterned substrates were prepared by photolithography. Photoresist (AZ 5214)
patterns were first prepared via photolithography, and the substrate
was immersed in octadecyltrichlorosilane (OTS, Aldrich) solution
(1:250 v/v in anhydrous hexane) for 5 min to cover bare SiO2 regions
with OTS molecules. The substrate was then sonicated in acetone
and methanol solution to remove the photoresist, resulting in
hydrophobic OTS self-assembled monolayer (SAM) patterns. When
the patterned substrate was immersed in swCNT (Carbon Nanotechnologies, Inc.) solution (0.05 mg mL−1 in 1,2-dichlorobenzene) for
1 min, swCNTs were adsorbed only on the bare surface regions,
while the CH3-terminated SAM prevented their adsorption. After
that, the substrate was immersed in the same SAM solution to passivate remaining bare surface regions in the CNT patterned region.
or from the author.
Acknowledgements
This work was supported by the National Research Foundation
grant (No. 2010–0000799) and the International Research & Development Program from the MEST (No. 2010–00293). SH acknowledges support from the Converging Research Center Program (No.
2010k001138) and the Happy tech. program (No. 20100020821)
from the MEST. KBL acknowledges the NIH Directors’ Innovator
small 2011, 7, No. 6, 741–745
Award (1DP20D006462–01) and is also grateful to the NJ commission on Spinal Cord grant (09–3085-SCR-E-0). KYB acknowledges
the support from the National Research Foundation of Korea Grant
funded by the Korean Government (No.2010–0029418). We thank
Jaehyuk Choi, Aniruddh Solanki, Birju Shah, and Shreyas Shah for
helpful discussions.
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Received: October 29, 2010
Revised: December 12, 2010
Published online: February 7, 2011
745
ARTICLE
Polarization-Controlled Differentiation
of Human Neural Stem Cells Using
Synergistic Cues from the Patterns of
Carbon Nanotube Monolayer Coating
Sung Young Park,†,‡ Dong Shin Choi,† Hye Jun Jin,‡ Juhun Park,‡ Kyung-Eun Byun,‡ Ki-Bum Lee,§,*
and Seunghun Hong†,‡,^,*
†
Interdisciplinary Program in Nano-Science and Technology and ‡Department of Physics and Astronomy, Seoul National University, Seoul 151-747, Korea,
Department of Chemistry and Chemical Biology, Rutgers, The State University of New Jersey, Piscataway, New Jersey 08854, United States, and ^Deparment of
Biophysics and Chemical Biology, Seoul National University, Seoul 151-747, Korea
§
D
ue to the ability to generate the
main phenotypes in the nervous systems, neural stem cells (NSCs) offer
great potential in regenerative medicine.1
For therapeutic applications, such as rebuilding damaged nerves, one should be
able to precisely control the direction and
structural polarization of individual axonal
growth.2 Previously, attempts have been
made to control the structural polarization
of cultured neurons by using several key
strategies such as molecular cues of diffusible gradient or substrate-bound chemical/
extracellular matrix (ECM) protein patterns
and topographical cues.3 12 However, they
have several disadvantages. For example, a
diffusible gradient is not suitable for the
membrane/matrix proteins due to the difficulty of maintaining it over time. In the case
of printed protein patterns, the protein
molecules may undergo conformational
changes during the process of stamping,
which often leads to the denaturation and
the loss of biological activities.13 Besides,
to create optimal nanotopographical cues
which can help cell growth, various microor nanofabrication techniques are required,
such as electron-beam lithography or chemical/reactive-ion etching.14 On the other
hand, various synthetic nanomaterials such
as biocompatible nanofibers and carbon nanomaterials have been recently proposed for
novel nanostructured scaffolds.14 19 However, neuronal polarization control of NSCs,
especially at the level of individual axons or
dendrites, has not been demonstrated
using these nanomaterials.
Herein, we report a method for the
structural-polarization-controlled neuronal
PARK ET AL.
ABSTRACT We report a method for selective growth and structural-polarization-controlled
neuronal differentiation of human neural stem cells (hNSCs) into neurons using carbon nanotube
network patterns. The CNT patterns provide synergistic cues for the differentiation of hNSCs in
physiological solution and an optimal nanotopography at the same time with good biocompatibility.
We demonstrated a polarization-controlled neuronal differentiation at the level of individual NSCs.
This result should provide a stable and versatile platform for controlling the hNSC growth because
CNT patterns are known to be stable in time unlike commonly used organic molecular patterns.
KEYWORDS: neural stem cells . carbon nanotubes . polarization . nanotopography .
micropattern
differentiation of human NSCs (hNSCs) using
the patterns of CNT network structures with
good biocompatibility. In this strategy, the
CNT network patterns provided synergistic
cues of selective laminin adsorption and
optimal nanotopography, which resulted
in selective adhesion and growth of hNSCs
on them. CNT network structures were
found to induce the enhanced adhesion
and growth of hNSCs even better than
conventional cell-culture substrates, such
as glass. CNT patterns with various geometries were utilized to explore their effect on
the outgrowth of hNSC during the growth
and differentiation process. As a proof of
concept, a structural-polarization-controlled
neuronal differentiation using CNT network
patterns was demonstrated at the level of
individual axons and neurites. Furthermore,
we applied our strategy for the controlled
hNSC growth on flexible and biocompatible
polymer substrates such as polyimide. Since
CNT monolayer coatings can be applied to
versatile substrates and provide stable microenvironments for hNSC growth control
even better than commonly used organic
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* Address correspondence to
seunghun@snu.ac.kr,
kblee@rutgers.edu.
Received for review February 14, 2011
and accepted May 9, 2011.
Published online May 13, 2011
10.1021/nn2006128
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Figure 1. CNT network patterns for selective hNSC growth and polarization. (a) Schematic diagram showing structural-polarization-controlled neuronal differentiation using CNT patterns. CNT monolayer patterns were fabricated on a substrate
using a previously reported method,22 and laminin was absorbed selectively on the CNT-coated regions. This structure induced preferential adhesion of hNSCs, finally achieving structural-polarization-controlled neuronal differentiation. (b) SEM
image of CNT patterns (dark spots). Scale bar represents 40 μm. (c) Immunofluorescence image of anti-laminin (green) bound
to the laminin which was selectively adsorbed on the CNT patterns. The scale bar represents 200 μm. It confirms the selective
adsorption of laminin on the CNT. The inset shows the AFM topography image of the laminin-coated CNT monolayer in phosphate buffered saline (PBS). The scale bar in the inset represents 2 μm. (d) Cell viability assay of hNSCs on CNT patterns for
3 day proliferation. The viability was measured by flow cytometry. The obtained data in the graph clearly indicate that 98% of
hNSCs grown on the CNT layer were alive (red).
molecular patterns, our work should provide a simple
but efficient way to control the structural polarization
of NSCs and may open up various applications in
neural engineering and regenerative medicine.
RESULTS AND DISCUSSION
Figure 1a shows a schematic diagram illustrating our
basic experimental procedure. CNT patterns were prepared according to previously reported methods.20,21
Briefly, a self-assembled monolayer (SAM) of methylterminated 1-octadecanethiol (ODT) was first patterned on thin Au films on cover glass substrates by
microcontact printing, while leaving some bare Au
surface regions unaltered (see method in Supporting
Information). When the patterned substrate was placed
in CNT suspensions (0.05 mg/mL in 1,2-dichlorobenznene), CNTs were selectively adsorbed onto bare Au
regions, forming CNT monolayer patterns. The CNT patterns were then placed in laminin solution (10 20 μg/mL)
for 10 30 min so that laminin molecules were selectively adsorbed onto the CNT patterns. Laminin is one
of the ECM components that is helpful for hNSC
adhesion and growth. After cell seeding, the hNSCs
grew preferentially along these laminin-coated CNT
patterns in the culture media with growth factors, such
PARK ET AL.
as basic fibroblast growth factor (bFGF) and epidermal
growth factor (EGF). For hNSCs, the growth factors
(bFGF and EGF) are known to enhance hNSC growth
and proliferation, while blocking the differentiation
process. Afterward, the substrate was placed in culture
media without bFGF and EGF for 2 weeks to study
the differentiation of hNSCs on laminin-coated CNT
patterns.
Figure 1b shows the scanning electron micrograph
(SEM) image of the prepared CNT patterns. It shows the
well-defined CNT regions (darker square regions) as
well as ODT-coated area (lighter region). The highresolution atomic force microscopy (AFM) image also
confirmed the highly selective adsorption of CNTs on
bare Au regions (Figure S1a in Supporting Information).
When placed in laminin solution, CNT patterns selectively adsorb laminin molecules from solution. This was
verified by immunochemistry (Figure 1c). For this purpose, after the laminin adsorption, the substrate was
placed in the fluorescent-labeled anti-laminin solution
so that the anti-laminin molecules would bind to the
laminin molecules on the substrate. The fluorescence
image shows much stronger fluorescence intensity in
the CNT regions (brighter green regions in Figure 1c) than
on ODT regions, confirming the high-density adsorption
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Figure 2. hNSC growth and differentiation depending on the size of CNT patterns. The phase contrast images of hNSCs grown
for 1 day (a,d,g) and those of the differentiated cells for 2 weeks (b,e,h), and the immunofluorescence images of the differentiated cells (c,f,i) are shown. The immunofluorescene markers are Hoechst for nuclei, glial fibrillary acidic protein (GFAP) for
astroglial cells, and TUJ1 and neurofilament light (NF-L) for neuronal cells. All scale bars represent 200 μm, unless otherwise
noted. The dotted black squares (a,b,d,e) indicate some of the CNT-coated regions. (a c) hNSC growth and differentiation on
rather large square-shape CNT patterns (300 μm 300 μm, 200 μm spacing). Note that neural networks were constructed in
arbitrary manner after differentiation. The immunofluorescence image (c) shows the differentiated cells positive for the
astroglial marker, GFAP (green). (d f) Restrictive neurite growth of hNSCs in individual CNT square patterns (50 μm 50 μm,
50 μm spacing). We did not observe any indication of neurite outgrowth of hNSC after the growth and differentiation from the
immunostaining image of NF-L (red). (g i) Outgrowths of hNSCs directed by rather small square-shape CNT patterns (5 μm 5 μm, 5 μm spacing). The inset figure (g) shows that a single hNSC was attached on seven individual CNT square patterns. The
immunofluorescence image (i) indicates that the differentiated cells are positive for neuronal cell marker, TUJ1 (red). The scale
bar in the phase contrast image (g) represents 100 μm.
of laminin molecules on the CNT patterns. It is also
consistent with previous reports regarding the preferential adsorption of protein molecules to CNT sidewalls22,23
and the resistance of alkyl chains of the ODT SAM to
laminin adsorption.24 The CNT patterns with laminin
coating were also investigated via an AFM topography
image (inset in Figure 1c and Figure S1b in Supporting
Information). It exhibited the average roughness of
26 nm, which is in the optimal range of surface roughness
(20 50 nm) promoting the adhesion and longevity of
primary neurons.25,26 This result indicates that the nanotopographic cues of CNT network structures as well as
laminin molecules adsorbed on the CNT patterns can
synergistically induce the selective growth of hNSCs.
The biocompatibility of the CNT network structure as
a substrate for hNSC growth was investigated via cell
PARK ET AL.
viability assay using flow cytometry. For the assay, the
adherent hNSCs were detached from the CNT patterns
after 3 day growth and 3 day differentiation, respectively. After the 3 day growth period, 98% of the cells
were found to be viable (Figure 1d). The assay result of
a 3 day differentiation also exhibited nearly 97% cell
viability (Figure S2 in Supporting Information). This
suggests the good biocompatibility of CNT patterns
for hNSC growth and differentiation. Furthermore, we
utilized the Western Blot method to confirm the protein expression of hNSCs before and after the differentiation (Figure S3 in Supporting Information). The
results show that the hNSCs grown with the growth
factors (EGF and bFGF) were positive for neural stem
cell markers (nestin and SOX2), which shows that they
just proliferated and undifferentiated. Meanwhile, those
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Figure 3. Control of hNSC orientation using line shape CNT patterns. (a c) hNSC growth on CNT line shape patterns (30 μm
width). The hNSCs inside individual 30 μm wide line patterns were observed to grow, extending their neurites in the same
direction along the predefined CNT line patterns. (d f) Individual hNSC growth on each CNT line pattern (5 μm width). The
hNSCs were aligned to form a bipolar shape on the CNT line patterns during the growth and differentiation. (g i) Neural
network formed on narrow line shape CNT patterns combined with large square-shape ones. Note that highly oriented hNSC
growth was induced by the predefined CNT patterns, and eventually well-organized neural networks were formed after differentiation. (a,d,g) Phase contrast images of hNSC grown for 1 day, and the scale bars in the phase contrast images are
200 μm. (b,e,h) Phase contrast images of the differentiated cell. (c,f,i) Immunofluorescence images of the differentiated cells.
The scale bars in the phase contrast images are 50 μm. The dotted black squares indicate some of the CNT-coated regions.
grown without these growth factors were positive for
glial fibrillary acidic protein (GFAP) and neuron-specific
class III β-tubulin (TUJ1), which indicates that they
differentiated.
When the hNSCs were seeded on the laminin-coated
CNT patterns in the culture media with the growth
factors, they selectively adhered onto the CNT pattern
regions and grew along the patterns (Figure 2a,d,g).
In this stage, the growth factors blocked the differentiation of hNSCs. When the substrate was placed in
the culture media without the growth factors, the
hNSCs started to differentiate (Figure 2b,e,h). The
differentiation was confirmed by immunocytochemistry (Figure 2c,f,i). Here, we used three different markers
to look at cytoskeletal distributions on the CNT patterns after the differentiation: GFAP as an astroglial cell
marker (Figure 2c), neurofilament light (NF-L, Figure 2f),
and TUJ1 (Figure 2i) as neuronal cell markers. We also
performed an experiment to investigate the hNSC
PARK ET AL.
growth on CNT networks compared with that on
conventional substrates such as coverglass (Figure S5
in Supporting Information). After the hNSC seeding on
the laminin-coated CNT patterns that were prepared
on coverglass (Figure S5A), we observed that the
hNSCs grew selectively in the CNT regions (Figure
S5B D). This result clearly indicates that the CNT network can provide a better extracellular environment
for hNSC growth than conventional cell-culture substrates such as coverglass.
Depending on the geometries of CNT patterns, the
hNSCs exhibited significantly different outgrowing behaviors during growth and differentiation (Figure 2a c).
When the size of the CNT square patterns was large
enough (300 μm 300 μm, 200 μm spacing) to hold
multiple cells, the hNSCs in the CNT patterns could
maintain their cell cell interactions and proliferated
very well (Figure 2a). Eventually, they outgrew over the
200 μm wide ODT SAM regions toward the adjacent
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Figure 4. Control of hNSC growth and differentiation on biocompatible and flexible polyimide (PI) substrate. (a) Optical image of a polyimide membrane with CNT patterns, which is flexible and transparent. (b) Immunofluorescence image of
antilaminin (green). It confirms that the laminin was selectively adsorbed onto the CNT patterns on PI substrate. Scale bar
represents 200 μm. (c) Phase contrast image of selective hNSC adhesion on CNT patterns on PI after cell seeding. Scale bar
represents 200 μm. (d) Immunofluorescence image of the differentiated hNSCs on CNT patterns on PI (TUJ1 for neural cells
and GFAP for astroglial cells). The inset shows the magnified image of the region marked by the white solid square. Scale bar
represents 200 μm, and that of the inset represents 50 μm. It should be noted that the orientation-controlled neural networks
were constructed along the CNT patterns on the PI membrane.
CNT square patterns and formed the neural networks,
where the cells grown on the distanced CNT square
patterns were connected (Figure 2b). The fluorescence
image clearly shows that the outgrowing astrocytes
(green regions marked as GFAP) were connecting the
hNSC population on the distanced patterns after differentiation (Figure 2c).
We then reduced the size of CNT square patterns
(50 μm 50 μm, 50 μm spacing) such that each square
could hold only a single hNSC (Figure 2d f). In this
case, the hNSC outgrowth was extremely restricted
during the growth and differentiation process
(Figure 2d). Even after we removed the growth factors
to induce differentiation, the hNSCs did not exhibit any
indication of major outgrowth over the ODT regions
(Figure 2e). The fluorescence image of neuronal cytoskeletons (NF-L, red) does not show any outgrowing
hNSCs from the patterns (Figure 2f). This result clearly
shows that the cell cell interaction can be controlled
by the geometries of CNT patterns, which can be
critical for hNSC growth and differentiation.
We also tested hNSC behaviors on CNT square
patterns smaller (5 μm 5 μm, 5 μm spacing) than
individual hNSCs (Figure 2g i). Here, the hNSCs first
adhered and outgrew over several CNT square patterns
(Figure 2g). Note that each cell was bound strongly on
the small CNT pattern regions and outgrew and extended over the ODT regions. In this case, the spacing
of the CNT square patterns should significantly affect
PARK ET AL.
the cytoskeletal tensions of the individual hNSCs,
which probably should affect the differentiation of
stem cells.27 After the differentiation, we could observe
that the neuronal outgrowths extended and bound on
the nearby CNT patterns (Figure 2h). The fluorescence
image clearly shows the neuronal cytoskeletal marker
(TUJ1, green) indicating the connected neural networks bound on the small CNT patterns. Overall, the
results in Figure 2 clearly show that the size and
spacing of CNT patterns can play a critical role in
controlling the hNSC outgrowths during the growth
and differentiation process, which can possibly affect
cell cell interactions or cytoskeletal tensions.
Furthermore, line shape CNT patterns can be utilized
to control the neuronal orientation with high precision
(Figure 3). In the line shape CNT patterns with a line
width (30 μm width, 60 μm spacing) that can hold
two or three cells, the hNSCs adhered (Figure 3a) along
the line pattern. We observed that they differentiated to form neural networks along the inside of the
line patterns (Figure 3b). Here, the differentiation was
also confirmed by immunocytochemistry with the
TUJ1 marker. When the CNT line width was narrowed
down to about 5 μm, which can hold only a single
hNSC, the hNSCs grew and differentiated into bipolar shapes along the individual CNT line patterns
(Figure 3f). Significantly, this result indicates that we
can control the orientation of hNSCs with single-celllevel precision.
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Figure 5. Structural-polarization-controlled neuronal differentiation of individual hNSCs using CNT patterns. (a) SEM image
of a CNT pattern with a single narrow strip as shown in dark gray. (b) Phase contrast images of hNSC adhesion on CNT patterns.
The dotted square and line (red) represent the CNT patterns. Scale bar represents 50 μm. After cell seeding, the cell bodies of
hNSCs were attached within the CNT square region. (c) Phase contrast images of the differentiated cells on the CNT patterns.
Note that the growing parts in the hNSCs were observed along the CNT single narrow strip regions during the differentiation.
(d) Immunofluorescence images of growth-associated protein 43 (GAP 43, green) and Hoechst (blue, for nucleus). Scale bar
represents 50 μm. It should be noted that the GAP 43 (green dots) was distributed along the narrow strip region. (e) Immunofluorescence image of GFAP (green), TUJ1 (red), and Hoechst (blue). Scale bar represents 50 μm. It should be noticed that
the differentiated neuronal cells (TUJ1, red) were surrounded by astroglial cells (GFAP, green) on the structural-polarizationcontrolled CNT pattern, where the neuronal polarization was also directed by the CNT narrow strip region.
When circle-shape patterns are connected with
narrow line shape ones, we observed quite an interesting
hNSC behavior during the growth and differentiation
(Figure 3g i). After being seeded in the culture media
with growth factors, the cell bodies of hNSCs tended to
adhere and proliferated on the circle-shape pattern regions (Figure 3g). After withdrawal of the growth factors
in the culture media, they started to differentiate and
the outgrowing neurites were observed mostly along
the narrow line shape CNT pattern regions (Figure 3h).
The differentiation was also confirmed via immunostaining (Figure 3i). Since the hNSCs first adhered and grew on
the circle-shape patterns, their nuclei (blue regions) were
mostly located on the circle regions, while the long
neurites (red regions) extended along the line shape
regions (Figure 3i). This result indicates that the CNT
patterns can be utilized to control both of the locations
of cell nuclei and the direction of neurite growth,
thus allowing us to control the structural polarization of
the neuronal differentiation of hNSCs. Furthermore, the
synapse formation of the neurons was checked by a
neuronal presynaptic vesicle marker, synaptophysin
(Figure S6 in Supporting Information). The results clearly
show that the neurons differentiated from the hNSCs
grown on the CNT patterns can also form the synapses,
which are important for a neuron to pass a chemical/
electrical signal to another neuron.
PARK ET AL.
For future therapeutic applications, such as regenerative medicine, it would be crucial to apply our
strategy to a flexible and biocompatible substrate such
as polyimide (PI) (Figure 4), which has been widely
utilized for implantable neural devices such as threedimensional artificial nerve conduits28 and stimulating
electrodes.29,30 We prepared CNT patterns on thin Aufilm-coated PI substrates and performed the experiments of hNSC growth and differentiation on them
(Figure 4a). We could achieve high-quality CNT patterns on the Au-coated PI substrates, as shown in the
SEM images (Figure S7a in Supporting Information).
The immunofluorescence image indicates the highly
selective adsorption of laminin onto the CNT patterns
on the PI substrate (green regions in Figure 4b). After
being seeded on it, the hNSCs adhered selectively onto
the CNT pattern regions on PI substrates and proliferated (Figure 4c). Eventually, we achieved the orientation-controlled growth and differentiation of hNSCs
along the CNT patterns on the flexible PI substrate
(Figure 4d and Figure S7b in Supporting Information).
Finally, the structural-polarization-controlled differentiation of individual hNSCs can be achieved by CNT
patterns composed of one square and one line shape
(Figure 5a). Here, the width of the line shape region is
much smaller than the size of an individual hNSC. After
cell seeding, we were able to observe the selective
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EXPERIMENTAL METHODS
Fabrication of CNT Monolayer Patterns. CNT (multiwalled CNT,
98% purified, NanoLab, MA, USA) patterns were fabricated on
Au-coated glass substrate according to the methods described
previously (Supporting Information).20,21 To prepare a polymer
substrate, polyimide (PI, VTEC Polyimide 1388, Richard Blaine
International, Inc., PA, USA) in solution was coated on a cover
glass by spin coating at 1000 rpm for 1 min and then cured on a
hot plate (Supporting Information). CNT patterns on Au-coated
PI were generated by the same method as before.20,21
hNSC Culture. Immortalized human NSCs (ReNcell VM, Millipore, Temecula, CA, USA) were purchased and maintained
according to the manufacturer's protocol.32 Differentiation was
initiated by removal of growth factors such as basic fibroblast
growth factor (bFGF) and epidermal growth factor (EGF) from
the culture media, and the cells were allowed to differentiate
usually for 2 weeks. For the hNSC culture, the prepared CNT
patterns were incubated in laminin solution (20 μg/mL, Sigma,
MO, USA) for 30 min. The laminin-coated CNT patterns were
washed with PBS several times and subsequently seeded with
suspensions of hNSC at a cell density of 105/mL. All of the hNSC
experiments were carried out between passages 3 and 10.
Cell Viability Assay. The hNSCs were either grown for 3 days or
subsequently differentiated for 3 days, and then they were used
for cell viability assay. The NSCs were first detached and made
into 106/mL cell suspensions, of which only a fraction was used
for counting cell viability. The cells were incubated with a
reagent composed of a mixture of a cell permeant and a noncell
permeant dye (ViaCount Reagent, Millipore, Heyward, CA, USA)
according to the manufacturer's protocol, and the viability was
determined using a single-laser four-color flow cytometry detection system (EasyCyte Plus, Millipore, Heyward, CA, USA) at
500 cells per one flow rate with predefined gating.
PARK ET AL.
CONCLUSION
In summary, we demonstrated a structural-polarization-controlled neuronal differentiation of hNSCs
using the patterns of CNT monolayer coating. Due to
the synergistic effect of CNT network structures for
selective laminin adsorption and optimal nanotopography, we could effectively promote the selective
growth of hNSCs on the CNT patterns. The result of
the cell viability assay (>97%) suggested the good
biocompatibility of CNT patterns for hNSC growth
and differentiation. We also confirmed that CNTs
could induce the adhesion and growth of hNSCs
even better than conventional cell-culture substrates such as bare glass. Importantly, the structural-polarization-controlled neuronal differentiation
was demonstrated at the level of an individual axon
or neurite. Furthermore, we also applied it to flexible
and biocompatible PI substrates, which should significantly expand the possible therapeutic applications of our method. Since CNT monolayer coatings
can be applied to versatile substrates including
flexible ones and provide a better cell-growth environment than conventional cell-culture substrates
such as glass, our strategy should provide many
new opportunities in various areas such as neural
engineering, stem cell therapy, and regenerative
medicine.
ARTICLE
hNSC adhesion inside the square regions (Figure 5b).
Then, the hNSCs on the square region outgrew along
the narrow line shape regions during the growth
and differentiation stages (Figure 5c). The neuronal
differentiation was confirmed by growth associated
protein 43 (GAP 43, green in Figure 5d), which is
known to be expressed in the growth cone regions
of neural cells. We observed that GAP 43 was also
highly expressed on the line shape CNT regions,
indicating that the neurites outgrew along the line
shape regions (Figure 5d and Figure S8 in Supporting
Information).
We carried out immunocytochemistry to check the
neural lineages of the differentiated cells on these CNT
patterns (Figure 5e). Here, GFAP and TUJ1 indicate
astroglial and neural cells, respectively. To confirm
their lineages, the relative fluorescence intensities of
GFAP and TUJ1 from the cell nuclei on the square
pattern regions were quantified using a method similar
to that reported previously (Figure S9 in Supporting
Information).31 The result shows that 20% of them
were TUJ1-positive, whereas another 20% were
GFAP-positive. It should be noted that the hNSCs were
differentiated with controlled structural polarity on the
CNT patterns, while maintaining their capabilities to
differentiate into the main phenotypes in the nervous
system, such as neuronal or astroglial cells.
Immunocytochemistry. The hNSCs were fixed for 15 min in 4%
paraformaldehyde in PBS and permeabilized with 0.1% Triton
X-100 in PBS for 15 min, followed by overnight incubation at
4 C in the following primary antibodies: TUJ1 (1:500; clone
SDL.3D10, Sigma, MO, USA), GFAP (1:1000; Dako, Glostrup,
Denmark), NF-L (1:200; Millipore, Temecula, CA, USA), GAP 43
(1:200; Millipore, Temecula, CA, USA), and synaptophysin
(Millipore, Temecula, CA, USA). Cells were washed with PBS,
incubated with either goat anti-mouse FITC (1:200; Sigma, MO,
USA) or goat anti-rabbit TRITC (1:500; Sigma, MO, USA), then
counterstained with 10 mM Hoechst 33342 (Sigma, MO, USA).
The mounted samples were imaged using an inverted fluorescence microscope (Nikon, TE2000, Tokyo, Japan) with an
EMCCD monochrome digital camera (DQC-FS, Nikon, Tokyo,
Japan). ImageJ software (freely downloadable from National
Institutes of Health Web site, http://rsbweb.nih.gov/ij/) was
used for subsequent processing of the fluorescence images.
Acknowledgment. We appreciate J.H. Yi and E. Miljan for the
fruitful discussion to evaluate NSC culture and differentiation.
We also thank A. Solanki, B. Shah, and S. Shah for helpful
discussions. This project has been supported by the NRF Grant
(No. 2011-0000390), and partial support from the Happy Tech
Program (No. 20100020821). S.H. acknowledges the support
from the Converging Research Center program (No. 2010K001138) and the System 2010 program of the MKE. K.-B.L.
acknowledges the NIH Directors' Innovator Award (1DP20D006462-01) and is also grateful to the NJ commission on Spinal
Cord Grant (09-3085-SCR-E-0).
Supporting Information Available: Supplementary methods,
additional details on fabrication method, and supplementary
figures. This material is available free of charge via the Internet
at http://pubs.acs.org.
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REFERENCES AND NOTES
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BRIEF ARTICLE
pubs.acs.org/molecularpharmaceutics
Synergistic Induction of Apoptosis in Brain Cancer Cells by Targeted
Codelivery of siRNA and Anticancer Drugs
Cheoljin Kim,† Birju P. Shah,† Prasad Subramaniam, and Ki-Bum Lee*
Department of Chemistry and Chemical Biology, Rutgers, The State University of New Jersey, Piscataway, New Jersey 08854,
United States
bS Supporting Information
ABSTRACT: Multiple dysregulated pathways in tumors
necessitate targeting multiple oncogenic elements by combining
orthogonal therapeutic moieties like short-interfering RNAs
(siRNA) and drug molecules in order to achieve a synergistic
therapeutic effect. In this manuscript, we describe the synthesis
of cyclodextrin-modified dendritic polyamines (DexAMs) and
their application as a multicomponent delivery vehicle for
translocating siRNA and anticancer drugs. The presence of βcyclodextrins in our DexAMs facilitated complexation and
intracellular uptake of hydrophobic anticancer drugs, suberoylanilide hydroxamic acid (SAHA) and erlotinib, whereas the cationic polyamine backbone allowed for electrostatic interaction with
the negatively charged siRNA. The DexAM complexes were found to have minimal cytotoxicity over a wide range of concentrations
and were found to efficiently deliver siRNA, thereby silencing the expression of targeted genes. As a proof of concept, we
demonstrated that upon appropriate modification with targeting ligands, we were able to simultaneously deliver multiple payloads
—siRNA against oncogenic receptor, EGFRvIII and anticancer drugs (SAHA or erlotinib)—efficiently and selectively to
glioblastoma cells. Codelivery of siRNA-EGFRvIII and SAHA/erlotinib in glioblastoma cells was found to significantly inhibit
cell proliferation and induce apoptosis, as compared to the individual treatments.
KEYWORDS: RNA interference, codelivery, cyclodextrins, SAHA, brain tumor cells, targeted delivery
’ INTRODUCTION
Advances in the field of chemical genetics and molecular cell
biology have triggered a surge in development of genetic
manipulation based therapies for cancer.1,2 Such genetic manipulation methods typically rely on either the traditional smallmolecule/protein modalities3 or the newly discovered RNA
interference (RNAi) based modalities,4 each having their own
advantages and disadvantages. For example, RNAi therapeutics
can provide attractive solutions to the major shortcomings of the
conventional therapeutics, including difficulty in lead identification and complex synthesis of small organic molecules and
proteins, and potentially can be applicable to all molecular targets
for cancer therapy.5 However, RNAi-based therapeutics, such as
small interfering RNA (siRNA) and micro RNA (miRNA), are
inherently antagonistic and their downstream effects (i.e., genesilencing) are delayed, compared to those of conventional smallmolecule/protein-based therapeutics.6 Additionally, owing to
their short serum half-life and poor cellular uptake, successful
clinical application of siRNA requires appropriate chemical
modifications and better delivery vehicles to overcome the
numerous cellular barriers.4 On the other hand, small organic
molecules can act as both antagonists and agonists for molecular
targets and their drug effects can be much faster than siRNA with
minimal problems during their intracellular uptake.5 Hence, from
a biological perspective, it would be beneficial to combine the
r 2011 American Chemical Society
advantages of these therapeutic modalities to potentially enhance
their individual efficacy. For example, it was recently demonstrated that simultaneous delivery of siRNA against multidrug
resistance genes in cancer cells led to the enhanced efficacy of the
codelivered anticancer drugs.7,8 These studies show that it would
be desirable to target multiple oncogenic signaling elements
using different therapeutic modalities for cooperative effect,
especially considering the molecular heterogeneity of tumors.
However, to achieve this goal, the primary requirement is to
develop noncytotoxic codelivery platforms capable of efficient
translocation of siRNA and small molecules with specificity as
well as identify the right combination of siRNA and small
molecules for a cooperative therapeutic effect.
To address the aforementioned need for cooperative chemotherapeutics, herein we describe the synthesis of a multifunctional delivery platform consisting of a dendritic polyamine
backbone conjugated with β-cyclodextrin (β-CD) moieties
[henceforth referred to as DexAMs] and its application for
target-specific codelivery of two orthogonal chemotherapeutic
molecules (siRNA and anticancer drug). We hypothesize that
Received: December 27, 2010
Accepted: July 27, 2011
Revised:
July 11, 2011
Published: July 27, 2011
1955
dx.doi.org/10.1021/mp100460h | Mol. Pharmaceutics 2011, 8, 1955–1961
Molecular Pharmaceutics
BRIEF ARTICLE
Scheme 1. (A) General Scheme Showing Codelivery of Small Molecules Like Anticancer Drugs and siRNA to Cancer Cells Using
Cyclodextrin Modified Polyamines (DeXAMs). (B) Chemical Structure of the Delivery Vehiclea
a
See Supporting Information for other DexAM generations.
codelivery of siRNA and anticancer drugs will have a cooperative
therapeutic effect against the target oncogenic signaling pathway
(EGFRvIII-PI3K/AKT), resulting in the selective induction of
apoptosis in brain tumor cells (Scheme 1). Additionally, conjugation of targeting ligands against receptors overexpressed in
brain cancer cells (EGFR) would allow for selective uptake of our
complexes into glioblastoma cells, thereby minimizing toxic side
effects on normal cells.
Additionally, our delivery platform and synthetic methods
have several advantages, as compared to conventional carrier
molecules (e.g., polyethyleneimine (PEI) and polyamidoamine
(PAMAM)). These include (i) minimal cytotoxicity and high
transfection efficiency of siRNA/drugDexAM constructs, (ii)
significantly higher yields and purity of DexAMs and increased
aqueous solubility of DexAM constructs, and (iii) capability of
simultaneously delivering nucleic acids, small organic molecules
and proteins, thereby achieving cooperative therapeutic effects.
’ MATERIALS AND METHODS
Starting materials, reagents, and solvents were purchased from
commercial suppliers (Sigma-Aldrich, Acros, and Fisher) and
used as received unless otherwise noted. All reactions were
conducted in flame-dried glassware with magnetic stirring under
an atmosphere of dry nitrogen. Reaction progress was monitored
by analytical thin layer chromatography (TLC) using 250 μm
silica gel plates (Dynamic Absorbents F-254). Visualization was
accomplished with UV light and potassium permanganate stain,
followed by heating. Proton nuclear magnetic resonance (1H NMR)
spectra were recorded on either a Varian-300 instrument (300 MHz),
a Varian-400 instrument (400 MHz) or a Varian-500 instrument
(500 MHz). Chemical shifts of the compounds are reported in
ppm relative to tetramethylsilane (TMS) as the internal standard.
Data are reported as follows: chemical shift, integration, multiplicity (s = singlet, d = doublet, t = triplet, q = quartet, br = broad,
m = multiplet), and coupling constants (Hz).
Quantification of siRNA Loading Efficiency. The complexes
were prepared at various charge ratios by mixing equal volumes
of DexAM with siRNA in PBS. Charge ratios (N/P) were calculated as a ratio of the number of primary amines in the polymer,
determined from 1H NMR spectra, to the number of anionic
phosphate groups in the siRNA. The samples were then incubated at room temperature for 30 min to ensure complex formation.
The complexes were prepared at a final siRNA concentration of
0.2 μg of siRNA/100 μL of solution. 100 μL of each complex was
transferred to a 96-well (black-walled, clear-bottom, nonadsorbing)
plate (Corning, NY, USA). A total of 100 μL of diluted PicoGreen dye (1:200 dilution in Tris- EDTA (TE) buffer) was
added to each sample. Fluorescence measurements were made
after 10 min of incubation at room temperature using a M200 Pro
Multimode Detector (Tecan USA Inc., Durham, NC, USA), at
excitation and emission wavelengths of 485 and 535 nm, respectively. All measurements were corrected for background fluorescence from a solution containing only buffer and PicoGreen dye.
Particle Size and Zeta Potential Analysis. Dynamic light
scattering (DLS) and zeta potential analyses were performed
using a Malvern Instruments Zetasizer Nano ZS-90 instrument
(Southboro, MA) with reproducibility being verified by collection
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dx.doi.org/10.1021/mp100460h |Mol. Pharmaceutics 2011, 8, 1955–1961
and comparison of sequential measurements. Polymer/siRNA
complexes (siRNA concentration = 330 nM) at different polymer concentrations were prepared using purified water (resistivity =
18.5 MΩ-cm). DLS measurements were performed at a 90°
scattering angle at 25 °C. Z-average sizes of three sequential
measurements were collected and analyzed. Zeta potential
measurements were collected at 25 °C, and the Z-average
potentials following three sequential measurements were collected and analyzed.
Cell Culture. Cells were cultured in the following growth
media: DMEM (Dulbecco’s modified Eagle’s medium) with high
glucose (Invitrogen), 10% fetal bovine serum (FBS), 1% streptomycinpenicillin, 1% glutamax (Invitrogen), and selection
markers, G418 (100 μg/mL) and hygromycin B (30 μg/mL)
for U87-EGFP and U87-EGFRvIII respectively. PC-12 cells were
cultured in DMEM with 10% horse serum, 5% FBS and 1%
streptomycinpenicillin. For the knockdown experiment and
targeted delivery, passaged cells were prepared to 4060%
confluency in 24-well plates. For the knockdown experiment,
targeted delivery and cell viability assay, medium was exchanged
with serum-free basal medium (500 μL) and siRNADexAM
solution (50 μL) was added after 2030 min. After incubation
for 12 h, medium was exchanged with normal medium. Fluorescence measurement and cellular assays were performed after
4896 h from the starting point.
Cytotoxicity Assays. The percentage of viable cells was
determined by MTS assay following standard protocols described by the manufacturer. All experiments were conducted in
triplicate and averaged. The quantification of polymer-mediated
toxicity was done using MTS assay after incubating the glioblastoma cells in the presence of varying concentrations of only
polymer vehicle for 4896 h. The data is represented as formazan
absorbance at 490 nm, considering the control (untreated) cells
as 100% viable.
Quantification of Knockdown of EGFP Expression (ImageJ).
Following siRNA treatment, cells were washed with DPBS and fixed
with 24% paraformaldehyde solution prior to imaging. For the
fluorescence, DIC and phase contrast images were obtained using
the Zeiss Axio observer inverted epifluorescence microscope. Each
image was captured with different channels and focus. Images were
processed and overlapped using Image-Pro (Media Cybernetics)
and ImageJ (NIH).
Targeted Delivery. Highly tumorigenic U87-EGFP cells and
low-tumorigenic PC-12 cells were cultured in 24-well plates, at a
density of 5 104 cells per well. For PC-12 cells, the normal
growth medium was DMEM (with high glucose, Invitrogen), 5%
horse serum, 10% FBS, 1% Glutamax, and 1% penicillinstreptomycin. For the delivery of EGFR-Ab conjugated DexAM
polyplexes, medium was exchanged with serum free DMEM
medium. The cells were incubated in the Ab-conjugated polyplex
medium for 68 h. Fluorescence images were taken after
replacing the serum-free medium with regular medium.
Apoptosis Assay. Cells were harvested by trypsinization and
stained using an Annexin V FITC Apoptosis Detection kit
(Roche, Cambridge, MA) according to the manufacturer’s protocol. The stained cells were immediately analyzed by flow
cytometry (FACScan; Becton Dickinson, Franklin Lake, NJ).
Early apoptotic cells with exposed phosphatidylserine but intact
cell membranes bound to Annexin VFITC but excluded
propidium iodide. Cells in necrotic or late apoptotic stages were
labeled with both Annexin VFITC and propidium iodide.
BRIEF ARTICLE
’ RESULTS AND DISCUSSION
Using multistep solution-phase and solid-phase synthesis, we
generated a series of highly water soluble dendritic polyamine
compounds conjugated to one or more β-cyclodextrin (β-CD)
molecules, referred to as DexAMs, with higher yield and purity as
compared to reported syntheses (Scheme 2). The first step for
synthesizing DexAM involved generating a dendritic polyamine
backbone by Michael addition of tris(2-aminoethyl)amine and
methyl acrylate, followed by amidation of the amino esters generated after Michael addition. The use of tris(2-aminoethyl)amine
as the core initiator yielded higher surface amine groups and
hence more compact dendrimers as compared to the reported
synthetic methods (for, e.g., ethylenediamine, ammonia) for
PAMAM dendrimers.9 The conjugation of β-cyclodextrin to
the polyamine backbone involved tosylation of β-cyclodextrin,
followed by nucleophilic addition with amine group. Compared
to the previously reported protocol,10 where tosyl chloride was
used for regioselective tosylation of β-cyclodextrin resulting in
very low yields, we improved the synthetic yield (∼50%) and
purity by using tosylimidazole, instead of tosyl chloride, under
reflux conditions to generate 6-monotosylated β-cyclodextrin
(see Supporting Information). In the final step, polyamine backbone was conjugated to tosylated CD via nucleophilic addition to
generate cyclodextrin conjugated polyamines, resulting in a 25fold increase in the aqueous solubility of CD (>50 g/100 mL) as
compared to that of CD alone (<1.8 g/100 mL), owing to
generation of an aminium salt (see Supporting Information for
the detailed synthesis).
The first component of our delivery vehicle—β-CD—has
been extensively used in pharmaceutical applications to improve
solubility of hydrophobic moieties, such as anticancer drugs.11
Many anticancer drugs are known to have poor aqueous solubility, thereby necessitating the use of toxic organic solvents like
dimethyl sulfoxide (DMSO), which can be detrimental in
biological applications.12 The presence of β-CD in our DexAM
moiety and the optimized drug loading would not only prevent
the use of such toxic solvents but also improve the water
solubility of CDdrug complex for the optimal cellular uptake
and drug efficacy. In our study, two hydrophobic anticancer
drugs [erlotinib and suberoylanilide hydroxamic acid (SAHA)]
were synthesized and loaded into the β-CD cavity by using our
optimized protocols (Figure 2b).13,14 For instance, by utilizing
the pH-dependent solubility of erlotinib, we could load drug up
to almost 50% of the molar ratio of β-CD, resulting in a significant
increase in its aqueous solubility (178 mg/100 mL).15,16 Similarly, we complexed SAHA with β-CD under reflux conditions to
obtain highly water soluble SAHACD complexes (solubility:
175 mg/100 mL) (see Supporting Information for the more
detailed synthesis and experimental protocols).17 The second
component of our DexAMs—dendritic polyamine backbone—
provides a positive surface charge which can interact electrostatically with the negatively charged nucleic acids, condensing
them into cationic complexes (known as polyplexes), thus facilitating their intracellular uptake and endosomal escape.1820
However, these primary/tertiary amines are also responsible for
cytotoxicity by interacting with the cellular components and
interfering in the cellular processes.21 Hence, there is a clear need
to develop synthetic chemistry to control the ratio of electrostatic
properties and the size of polymer structures. Our synthetic
methods enabled us to precisely control the number of primary
amine head groups from 4 to 48 leading to four different generations
1957
BRIEF ARTICLE
Scheme 2. (A) Schematic Representation of Synthesis of DexAMs. (B) Conjugation of Drugs and Antibodies to DexAMs
of DexAMs molecules (D1D4), thereby allowing us to achieve
an optimal balance between cytotoxicity and complexation
ability.
We assessed the capability of our four different generations of
DexAMs (D1D4) to spontaneously form complexes with the
negatively charged siRNA using a well-established dye exclusion
assay. As the number of amine groups increased from DexAM-1
(D1, 4 primary amines) to DexAM-4 (D4, 48 primary amines),
the amount of free/unbound siRNA decreased correspondingly
at a given DexAM concentration (see Figure S1 in the Supporting
Information). Since we found that the complexation ability of
DexAM-4 is higher than that of the other generations with
minimal cytotoxicity, we proceeded with using DexAM-4 for
the subsequent experiments. Additionally, the hydrodynamic
diameters of the resultant polyplexes could be controlled from
250 to 400 nm with polydispersity index of 0.81.0 by increasing
the polymer concentration (see Figure S2a in the Supporting
Information). The zeta potentials of the resulting polyplexes
were in the range of 810 mV at pH 7.4 (see Figure S2b in the
Supporting Information), demonstrating the cationic nature of
the polyplexes. Cytotoxicity of the DexAM molecules was
assessed using MTS assay. First we confirmed the effect of the
β-CD moiety on the cytotoxicity of DexAMs by comparing the
cytotoxicity of the DexAM (containing CD) to that of the
DexAM without CD. Our cytotoxicity assay data clearly shows
that the DexAM constructs with CD show significantly less
cytotoxicity as compared to those without CD (Figure 1a). We
believe this is due to the presence of CDs on a polycationic
backbone in DexAM, which can potentially reduce nonspecific
binding of the DexAM constructs with proteins or cellular
structures.2023 We also compared the cytotoxicity of our DexAMs
with the commercially available transfection agents, polyethyleneimine (PEI), Lipofectamine 2000 (LF) and X-tremeGENE (Xgene)
at the recommended concentrations for transfection, and found
that those agents were significantly more cytotoxic at those
concentrations as compared to DexAMs (Figure 1b).
The optimization of gene silencing with our siRNADexAM
constructs and assessment of knockdown efficiency were performed by measuring the suppression of enhanced green fluorescent protein (EGFP) in glioblastoma cell lines (U87-EGFP),
which were genetically modified to constitutively express EGFP.
The decrease of green fluorescence intensity due to siRNAmediated EGFP silencing was monitored over a time period of
4896 h to quantify the knockdown efficiency of our DexAM/
siRNA constructs (see Figure S4 in the Supporting Information).
Approximately 70% of the U87-EGFP cells showed no EGFP
signal after 96 h of siRNA treatment as compared to the control
cells at a polymer concentration of 100 μM (Figure 1c) with
negligible cytotoxicity (∼95% cell viability). In parallel, we
compared the transfection efficiency and the corresponding
cytotoxicity of our delivery platform with that of the commercially available transfection agent (X-tremeGENE) under the same
condition, in which X-tremeGENE-based transfection demonstrated similar levels of EGFP knockdown (∼70% knockdown
efficiency), albeit with significant toxicity (∼30% cell viability)
(see Figure S5 in the Supporting Information).
In addition to efficient translocation of siRNA across the cell
membrane with minimal cytotoxicity, successful therapeutic application of siRNA also requires the siRNA to interact with the RNAi
machinery within a target cell, thereby minimizing off-target effects.24
Brain tumor cells, particularly glioblastoma cells, are known to
present high levels of epidermal growth factor receptors (EGFRs)
on their cell surface, thus making it a specific biomarker for cellspecific delivery toward brain tumor cells.25 For targeted delivery
to glioblastoma cells, we modified our DexAM-4 with appropriate ratios of EGFR antibodies (DexAM-4:EGFR-Ab = 1:5)
1958
BRIEF ARTICLE
Figure 1. Cytotoxicity and transfection efficiency of DexAM-4 (D4). (A) Effect of cyclodextrin grafting on polymer-mediated toxicity. (B) Comparison
of toxicities of DexAM-4 with commercially used transfection agents at optimized concentrations of delivery agent and siRNA. (LF 2000, Lipofectamine
2000; and Xgene, X-tremeGENE). (C) Phase contrast (C1, C2) and fluorescent (C3, C4) images showing siRNA-mediated decrease in green
fluorescence in treated and control (untreated) U87-EGFP cells.
and incubated them in U87 (glioblastoma cell line, target cells)
and other less-tumorigenic PC-12 cells (control cells) which tend
to have low levels of expression of EGFRs. The DexAM-4 constructs were also labeled with a fluorescent dye (Alexa Fluor 594)
to monitor their intracellular uptake using fluorescence microscopy.
From our data we could see that EGFR-antibody modified DexAM-4
were selectively translocated into U87 (target glioblastoma cells)
with high efficiency as compared to the PC-12 (control cells)
(Figure 2a).
Having demonstrated the target-specific delivery and efficient
gene silencing capability of the siRNADexAM constructs, we
then focused on our main goal of codelivering siRNA and
anticancer drugs for targeting key oncogenic signaling pathways
(e.g., EGFRvIII-(phoshphatidylinositol-3-kinase)PI3K/AKT) to
achieve a cooperative chemotherapeutic effect. Tumors harbor
multiple dysregulated signaling pathways, thus limiting the
clinical utility of single target agents.26 Hence, combining approaches targeting multiple oncogenic elements, using a single
delivery platform, can not only increase the likelihood of blocking
tumor survival and metastasis, as compared to individual treatments, but also simplify clinical applications. For this purpose, we
focused on developing a combinatory therapeutic approach
based on siRNA and anticancer cancer drugs targeting oncogenic
pathways in glioblastoma multiforme (GBM), an extremely
aggressive and difficult-to-treat form of primary brain tumor.
We aimed at downregulating the EGFRvIII-PI3K/AKT pathway,
implicated in the proliferation and apoptosis of brain tumor cells,
by delivering siRNA against epidermal growth factor receptor
variant III (EGFRvIII), which is known to enhance the tumorigenicity of GBM.2729 However, due to tumor molecular
heterogeneity, only siRNA-based downregulation of a single
oncogenic target (EGFRvIII) may not be efficacious. Histone
deacetylase (HDAC) inhibitors like suberoylanilide hydroxamic
acid (SAHA) and EGFR tyrosine kinase inhibitors like erlotinib
have been reported to enhance the efficacy of other EGFR
antagonists.26,30 To this end, we used either SAHA or erlotinib
for codelivery with siRNA against EGFRvIII oncogene to
deactivate the target signaling pathway in a selective and efficient
manner. These drugs have already shown some promising results
for GBM therapy, but have met with limited success since they
require higher doses and longer exposures, which may lead to
increased toxic side effects.31
Our hypothesis is that combination of anticancer drugs against
complementary therapeutic targets with siRNA therapeutics
against EGFRvIII would have a cooperative effect on induction
of apoptosis in brain tumor cells. To test this hypothesis, we
initially compared the antiproliferative capability of anticancer
drugs (SAHA and erlotinib) and siRNA against EGFRvIII in
glioblastoma cells, either individually or in combination by using
cell viability assay (Figure 2b). From the data, we could clearly
observe a cooperative inhibition of glioblastoma cell proliferation
when SAHA (5 μM) was codelivered with the siRNA (200 nM;
polymer concentration 100 μM), as compared to treating the
cells with only SAHA at the same concentration (5 μM). This can
be attributed to the fact that SAHA is known to significantly
enhance the efficacy of agents targeting EGFR signaling pathway
by modulating several indirect downstream targets, which in turn
are key regulators of EGFR pathways. Similarly, codelivery of
erlotinib (30 μM) and siRNA (200 nM) also inhibited tumor cell
proliferation to a higher extent (Figure 2b). Additionally, we also
monitored the effect of codelivery of both siRNA and anticancer
drugs on inducing cell death in glioblastoma cells using the
apoptosis assay (Annexin-V/propidium iodide assay). A significantly higher proportion of cell population treated with both
siRNA and SAHA was Annexin-VFITC-positive as compared
to the individual treatments as well as untreated cells. These
results indicate greater induction of apoptosis in cells treated with
both siRNA and SAHA, as compared to those with only SAHA
and only siRNA treatment (Figure 2c). A similar trend in the
cooperative induction of apoptosis was seen in the case of
1959
BRIEF ARTICLE
Figure 2. Targeted delivery of DexAMs and cooperative effect of anticancer drugs and siRNA on glioblastoma cells. (A) Targeted delivery of DexAMs
modified with EGFR antibodies in highly tumorigenic U87-EGFP cells and less-tumorigenic PC-12 cells. (B) Viability of glioblastoma cells following
individual treatments and codelivery of drugs and siRNA, based upon MTS assay. (C) Flow cytometry based Annexin-V/PI assay demonstrating the
apoptotic effect of combined and individual siRNA and drug treatments. Percentages represent Annexin-V-positive (apoptotic cells). For all
experiments, the polymer concentration was kept constant (100 μM), whereas the concentrations of SAHA, erlotinib and siRNA were 5 μM,
30 μM and 200 nM respectively.
combined erlotinib/siRNA treatment (Figure 2c). We also
found that complexation of SAHA and erlotinib within the
CD cavity improved their aqueous solubility and hence
increased their potency, measured as IC 50 values, by approximately 2-fold as compared to its DMSO solution (see Figure
S6 in the Supporting Information). Thus, these results show
the cooperative effect on selectively inducing the apoptosis of
brain tumor cells by the right combination of siRNA and
anticancer drugs and the capability of our delivery molecules
(DexAMs) for target-specific delivery and improved chemotherapeutic efficacy.
In conclusion, we synthesized a multimodal delivery platform
to simultaneously deliver two orthogonal therapeutic modalities
having cooperative therapeutic efficacy in an efficient and
selective manner to the target brain tumor cells. As a proof-ofconcept experiment, we demonstrated that target-specific codelivery of siRNA and anticancer drugs having complementary
therapeutic results would be a novel method to enhance the
apoptotic signaling pathways and inhibit the proliferation signaling pathways in brain tumor cells. Potentially, our approach and
methodology can be beneficial for introducing exogenous siRNA
combined with small molecules into other mammalian cells,
which can represent a powerful approach for the optimal
manipulating signal transduction. Our synthetic techniques
afforded facile manipulation of the polymer structure to achieve
efficient transfection with minimal polymer-mediated cytotoxicity. The strategy of codelivering anticancer drug with therapeutic siRNA is particularly advantageous for in vivo applications, so
that both the moieties are delivered to the target cells using a
single delivery platform. Our versatile delivery platform can also
be used to codeliver different kinds of small molecules and
nucleic acids to regulate cancer cell fate such as proliferation,
migration and apoptosis by targeting multiple signaling pathways. Collectively, our DexAM-based codelivery strategy has
significant potential for cancer therapy as well as regulating cell
fate by modulating key signaling cascades.
’ ASSOCIATED CONTENT
bS
Supporting Information. Detailed synthesis of DexAMs;
NMR characterization of synthesized compounds; conjugation
of targeted moieties to DexAMs, complexation of drugs, particle
size and zeta potentials of DexAM polyplexes, siRNA loading
efficiency. This material is available free of charge via the Internet
at http://pubs.acs.org.
’ AUTHOR INFORMATION
Corresponding Author
*Department of Chemistry and Chemical Biology, The State
University of New Jersey, Piscataway, NJ 08854, United States.
Tel: (+1) 732-445-2081. Fax: (+1) 732-445-5312. E-mail: kblee@
rutgers.edu. Homepage: http://rutchem.rutgers.edu/∼kbleeweb/.
Author Contributions
†
These authors contributed equally to this work.
’ ACKNOWLEDGMENT
The authors thank Joan Dubois for assisting with the flow
cytometry measurements and Kevin Memoli for providing us
with Erlotinib and SAHA. We are also grateful to KBLEE group
members for their valuable suggestions for the manuscript. This
work was supported by NIH Director’s Innovator Award
(1DP20D006462-01) and N.J. Commission on Spinal Cord
Research grant (09-3085-SCR-E-0).
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1961
APPLIED PHYSICS LETTERS 98, 173702 共2011兲
ZnO thin film transistor immunosensor with high sensitivity and selectivity
Pavel Ivanoff Reyes,1 Chieh-Jen Ku,1 Ziqing Duan,1 Yicheng Lu,1,a兲 Aniruddh Solanki,2
and Ki-Bum Lee2,a兲
1
Department of Electrical and Computer Engineering, Rutgers University, 94 Brett Road, Piscataway,
New Jersey 08854-8058, USA
2
Department of Chemistry and Chemical Biology, Rutgers University, 610 Taylor Road, Piscataway,
New Jersey 08854-8058, USA
共Received 11 January 2011; accepted 3 April 2011; published online 29 April 2011兲
A zinc oxide thin film transistor-based immunosensor 共ZnO-bioTFT兲 is presented. The back-gate
TFT has an on-off ratio of 108 and a threshold voltage of 4.25 V. The ZnO channel surface is
biofunctionalized with primary monoclonal antibodies that selectively bind with epidermal growth
factor receptor 共EGFR兲. Detection of the antibody-antigen reaction is achieved through channel
carrier modulation via pseudo double-gating field effect caused by the biochemical reaction. The
sensitivity of 10 fM detection of pure EGFR proteins is achieved. The ZnO-bioTFT immunosensor
also enables selectively detecting 10 fM of EGFR in a 5 mg/ml goat serum solution containing
various other proteins. © 2011 American Institute of Physics. 关doi:10.1063/1.3582555兴
The ion-selective field effect transistors 共ISFETs兲 has
been used popularly as a sensitive pH sensor and various
biochemical sensors.1–3 Recently the ISFET structure has
been integrated with poly-Si thin film transistors 共TFTs兲 and
GaN/AlGaN high electron mobility transistors for detection
of DNA, penicillin, and cellular potentials.4,5 However, the
sensing procedure using the ISFET can be invasive as its
entire gate serves as the sensing area which contains both the
analyte solution and the reference electrode. Another class of
FET-type biosensors is based on organic field-effect transistors 共OFETs兲.6–9 The general structure of an OFET consists
of a back-gate metal-oxide semiconductor field-effect transistor 共MOSFET兲 with the conducting channel made of organic semiconductors. The OFET has the advantage of being
easily controlled through biasing due to the back-gate configuration. However, OFETs require high bias voltages, and
suffer from low channel mobility. Currently, nanowire-based
FET sensors are demonstrated with high sensitivity reaching
the order of fM.10,11 However, these prototypes of sensors
generally involve a complex fabrication process as they are
constructed individually by manipulating and aligning a
single strand of semiconducting nanowire such as TiO2 or Si
as the FET channel between the source and drain patterns. It
is difficult to achieve repeatability and manufacturability in
fabrication and integration of these devices for larger sensor
arrays.
ZnO is emerging as a wide band gap semiconductor oxide with multifunctional properties that makes it an attractive
sensor material. ZnO and its nanostructures are compatible
with intracellular material and ZnO–based sensors have been
demonstrated for detection of biochemicals such as enzymes,
antibodies, DNA immobilization, and hybridization.12–15 In
this letter, we report the highly sensitive and selective immunosensing ability of a ZnO based TFT biosensor 共ZnObioTFT兲. The epidermal growth factor receptor 共EGFR兲 is
used as the example because the sensing of EGFR-antibodies
reacting with EGFR proteins has its implications in cancer
related studies and drug screening for cancer, as EGFR is
a兲
Authors to whom correspondence should be addressed. Electronic addresses: ylu@rci.rutgers.edu and kblee@rutgers.edu.
0003-6951/2011/98共17兲/173702/3/$30.00
well-known to be overexpressed in solid tumors, especially
breast cancers. The ZnO-TFT devices possess excellent and
repeatable characteristics. It can be fabricated using conventional microelectronic process and can be integrated into a
large-scale at sensor arrays low cost, which will benefit further development of a device platform not only for diagnosing cancers, but also for monitoring a patient’s response to
therapy in real-time.
The device schematic is shown as the inset of Fig. 1共a兲.
It follows a back-gate inverted-staggered configuration. A Si
substrate was covered with 1 ␮m layer of SiO2 through wet
oxidation followed by e-beam deposition of a layer of Au 共50
nm兲/Cr 共100 nm兲 that serves as the gate electrode. A 70 nm
layer of SiO2 serving as the gate oxide was then deposited
through plasma enhanced chemical vapor deposition with
substrate temperature of 250 ° C and using SiH4 and N2O as
the source gases. A 50 nm ZnO thin film was grown using
metalorganic chemical vapor deposition on the top of the
FIG. 1. 共a兲 Transconductance curve of the ZnO-bioTFT and its vertical
structure schematic 共inset兲; 共b兲 transistor characteristic curves for various
gate bias, and the top view of the device 共inset兲.
98, 173702-1
© 2011 American Institute of Physics
173702-2
Reyes et al.
Appl. Phys. Lett. 98, 173702 共2011兲
FIG. 3. Drain current vs gate bias for various Molar concentrations of pure
EGFR proteins detected by the ZnO-bioTFT to demonstrate sensitivity.
FIG. 2. 共Color online兲共a兲 Drain current vs gate bias for fixed drain bias of
10 V. Step 1: bare device, step 2: EGFR-antibody immobilization, and step
3: EGFR protein detection; 关共b兲–共d兲兴 schematic of the carrier modulation
mechanism for steps 1 to 3, respectively.
SiO2 to serve as the n-type conduction channel, with substrate temperature at 350 ° C and using diethyl zinc 共DEZn兲
as the metal precursor and ultrahigh purity O2 as oxidizer. Au
共50 nm兲/Ti 共100 nm兲 was deposited through e-beam evaporation for the source and drain Ohmic contacts. The exposed
ZnO channel acts as the sensing area and has a dimension of
200 ␮m ⫻ 400 ␮m, giving a W/L ratio of 2. Shown in the
inset of Fig. 1共b兲 is the top view of the TFT device. The
electrical characteristics of the ZnO-bioTFT are shown in
Figs. 1共a兲 and 1共b兲. The transconductance curve 关drain current 共ID兲 versus gate voltage 共VGS兲兴 in Fig. 1共a兲 shows that
the bioTFT is a normally-OFF enhancement mode transistor
with a threshold voltage of 4.25 V and an ON-OFF ratio of
⬃108. The high ON-OFF ratio of the device provides the
high sensitivity of the device to the charge modulation within
the ZnO channel. Figure 2共b兲 shows the transistor characteristic curves with drain current versus drain voltage for various gate-biasing of the device.
To realize the immunosensing ability of the ZnObioTFT, the exposed ZnO channel was functionalized using
linkage chemistry, which involves three basic steps. First, the
ZnO channel was functionalized with trimethoxysilane aldehyde 共having a reactive aldehyde end group兲 by incubating
the device in 1% v/v solution of the silane-aldehyde in 95%
ethanol for 30 min. The device was then cured at 120 ° C for
15 min. Second, the aldehyde groups were coupled to the
amine groups of the monoclonal EGFR antibodies 共1:50兲
through reductive amination in the presence of 4 mM sodium
cyanoborohydride in PBS 共pH 7.4兲 for two hours. Third,
unreacted aldehyde groups were blocked using 100 mM
ethanolamine in a similar manner to prevent nonspecific in-
teractions of proteins. Finally, the device was rinsed in a
continuous flow of PBS, pH 7.4 for 10 min.
The biofunctionalization enables the exposed ZnO channel direct interaction with the biochemical species being detected. The mechanism of detection of antibody-antigen reaction is illustrated in Figs. 2共a兲–2共d兲. In the first step 关Fig.
2共b兲兴 the unfunctionalized ZnO-bioTFT is positively biased
at the drain and gate electrode. The positive voltage at the
gate causes the majority carriers of the n-type ZnO channel
to accumulate near the base of the ZnO layer to facilitate a
conduction path for the current flow from drain to source.
The positive voltage at the drain causes some of the carriers
to also accumulate near the side of the drain electrode forming a wedge-shaped conduction path. The bias at the drain
also acts as the electron pump to drive the current to flow.
For the second step 关Fig. 2共c兲兴, the exposed ZnO channel is
functionalized with EGFR monoclonal antibodies 共mAbs兲
having free lysine groups. The immobilized antibody molecules caused significant decrease in conductivity of the ZnO
surface layer, thus, reducing the drain current. In the third
step 关Fig. 2共d兲兴, the EGFR protein captured by the EGFR
mAbs forms a polarized molecule with a dominant partiallypositive charged tip16 which led to the accumulation of negative carriers within the ZnO channel to accumulate near the
exposed surface where the antibody-protein pairs were
present. This carrier accumulation was in addition to the conduction path created near the gate. The combined amount of
accumulation layer caused an increase in the current flow.
The top molecule layer 共reacted protein兲 acted as a virtual
top gate and the antibody layer acted as a virtual insulator
layer, thus forming a pseudodouble gated field-effect conduction scheme for the ZnO-bioTFT. The actual measured drain
currents that confirmed each step of the detection process are
shown in Fig. 2共a兲. The drain voltage is fixed to 10 V and the
gate voltage is varied from ⫺5 to +15 V, and the drain
current is measured using an HP4156C semiconductor parameter analyzer and Cascade Microtech probe station.
To demonstrate the high sensitivity of the ZnO-bioTFT,
solutions of pure EGFR 共in PBS兲 were prepared with four
different Molar concentrations using serial dilutions, namely,
10 nM, 100 pM, 1 pM, and finally 10 fM. Each EGFR solution 共2 ␮l兲 was introduced to a separate but similar ZnObioTFT fabricated on a single chip that were simultaneously
functionalized with EGFR mAbs. The drain current was
monitored as a function of gate voltage with a fixed drain
voltage of 10 V, for each concentration. Figure 3 shows the
measured drain current versus gate voltage of the bioTFT. An
increase in drain current was measured as the EGFR concen-
173702-3
Appl. Phys. Lett. 98, 173702 共2011兲
Reyes et al.
device. Moreover, the device was able to discern as low as
10 fM of EGFR protein concentration in the serum solution.
The sensitivity plot of the device for both pure and in-serum
detection is shown in Fig. 4共b兲 which exhibits linearity in the
x-y logarithmic scale.
In summary, we have demonstrated a ZnO bioTFT that
has the ability to perform immunosensing with high sensitivity and selectivity. The channel of the bioTFT is functionalized with amine-terminated EGFR mAbs. EGFR proteins
with the lowest concentration of 10 fM were detected by the
device in both pure state and selectively in a concentration
serum solution containing various other protein species. The
ZnO-bioTFT enables bias-controlled operation though its
bottom gate configuration. The high sensitivity of the device
is attributed to its high on-off ratio, and the output current
trend is explained by the pseudo-double gating electric field
effect. The realization of the ZnO-bioTFT functionalized
with EGFR mAbs reacting with EGFR proteins has potential
applications in cancer diagnosis and treatment.
FIG. 4. 共a兲 Drain current vs gate bias for various molar concentrations of
EGFR-proteins in a serum solution containing many different proteins. 共b兲
Sensitivity plot of the device for pure protein and protein in serum detection.
tration was increased and the graph also shows that the device was able to detect as low as 10 fM of EGFR concentration. The trend in the current readings agrees with the
hypothesis provided by the pseudodouble gating effect discussed above.
The highly selective sensing of EGFR using the ZnObioTFT was also demonstrated. In this experiment, a 5
mg/ml 共in PBS, pH 7.4兲 goat serum solution was prepared,
which contains many different species of proteins. As mentioned above, different EGFR solutions were prepared,
namely, 100, 1, and 10 fM, using this serum solution as the
solvent and not pure PBS. For all the concentrations, the
total amount of serum present remained approximately the
same. Each of the different solutions 共2 ␮l兲 was introduced
onto a chip containing multiple similar bioTFT devices that
were biofunctionalized with EGFR mAbs. The drain current
of each device was measured as a function of gate voltage,
with a fixed drain voltage of 10 V. As a control, we first
introduced serum solution without the EGFR proteins to the
ZnO-bioTFT. Figure 4共a兲 shows no change in the drain current for the pure serum confirming that there were no EGFR
molecules in the solution. The drain current increased as a
function of EGFR concentration. The bio-TFT detected only
the EGFR proteins out of the many different proteins present
in the serum solution introduced onto the sensing area of the
This work has been supported in part by the AFOSR
under Grant No. FA9550-08-01-0452, and by the NSF under
Grant No. ECCS 1002178.
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www.advmat.de
www.MaterialsViews.com
Sung Myung, Aniruddh Solanki, Cheoljin Kim, Jaesung Park, Kwang S. Kim,
and Ki-Bum Lee*
Nanomaterials such as silicon nanowires (SiNWs),[1,2] carbon
nanotubes (CNTs),[3–6] and graphene,[7,8] have gained much
attention for use in electrical biosensors due to their nanoscopic and electrical properties. For instance, SiNWs and CNTs
can be integrated into field-effect transistors (FETs) to detect
small amounts of target biomolecules with high sensitivity and
selectivity by measuring electrical disturbances induced by the
binding of these biomolecules to the surface of the nanostructure.[9,10] The detection of biomarker proteins with high sensitivity and selectivity is vital for the early diagnosis of many
diseases including cancer and HIV. For this purpose, carbonbased nanomaterials such as CNTs and graphene have attracted
significant attention for fabricating highly sensitive FET-based
biosensors.[6,8,9,11–15] In particular, the use of graphene in FETbased biosensors is becoming more and more appealing not
only due to its unique properties, such as higher 2D electrical
conductivity, superb mechanical flexibility, large surface area,
and high chemical and thermal stability, but also due to its
ability to overcome the limitations of CNTs, such as variations
in electrical properties of CNT-based devices and the limited
surface area of CNTs.[16–24] Nevertheless, there have been only a
few reports on the development of graphene FET-based biosensors,[14,25] and their potential as biosensors has not been fully
explored. It is therefore critical to develop nanoscopic graphenebased biosensors that are simple in device structure, small in
size, and allow label-free detection and real-time monitoring of
biomarkers, all of which are essential criteria for biosensors. A
key challenge in the above requirements is the achievement of
both, well-organized 2D or 3D graphene structures, in microscopic and nanoscopic biosensing devices and well-defined bioconjugation chemistry on graphene.
Dr. S. Myung, A. Solanki, C. Kim, Prof. K.-B. Lee
Department of Chemistry and Chemical Biology
Rutgers, The State University of New Jersey
Prof. K.-B. Lee
Institute for Advanced Materials
Devices and Nanotechnology (IAMDN)
J. Park, Prof. K. S. Kim
Center for Superfunctional Materials
Department of Chemistry
Pohang University of Science and Technology
Pohang, Korea
DOI: 10.1002/adma.201100014
Adv. Mater. 2011, 23, 2221–2225
Here, we demonstrate a novel strategy for the fabrication and
application of a reduced graphene oxide (rGO)[26,27] encapsulated
nanoparticle (NP)-based FET biosensor for selective and sensitive detection of key biomarker proteins for breast cancer. It is
important to note that we used Human Epidermal growth factor
Receptor 2 (HER2) and epidermal growth factor receptor (EGFR),
which are known to be over-expressed in breast cancers,[28–30]
only as a proof-of-concept to demonstrate the high sensitivity and
selectivity of the graphene-encapsulated NP biosensor. This biosensor could be used to detect any important cancer markers with
relative ease. In the typical experiments for fabricating grapheneencapsulated NP-based biosensors, individual silicon oxide NPs
(100 nm) functionalized with 3-aminopropyltriethoxysilane
(APTES) were first coated with thin layers of graphene oxide
(GO), which prevent aggregation and maintain high electrical
conductivity (Figure 1). This was mainly achieved via the electrostatic interaction between the negatively charged GO (see Supporting Information, Figure S1) and the positively charged silicon
oxide NPs. The GO solution (0.05 mg mL−1 in deionized water)
was simply injected into the NP solution (5 mg mL−1), wherein
the negatively charged GO assembled on the positively charged
NP surface until equilibrium coverage was reached.[31–33] The
transmission electron microscopy (TEM) image of the GO-coated
NPs clearly shows the uniform assembly and saturation density
of GO on the NP surface (Figure 1a). The GO thickness on the
surface of the NPs was 5 nm, as measured from high-resolution
TEM (HR-TEM). As seen in the image, the NPs were connected
through a film of GO that was used as an electrical carrier after
its reduction to rGO. For efficient use of NP junctions as electrical channels, it was imperative to assemble the NPs with high
density on the device. The scanning electron microscopy (SEM)
images show well-defined, dense rGO-NP patterns uniformly
covering a large area of the silicon oxide substrate (Figure 2a,b).
Furthermore, the modified self-assembly method (using centrifugation) allowed us to assemble NPs with high density in a
short span and by using minimal amount and concentration of
the NP solution. Importantly, the high surface-to-volume ratio of
the GO-encapsulated NPs can generate 3D electrical surfaces that
significantly enhance detection limits and enable label-free, highly
reproducible detection of clinically important cancer markers.
One of the most attractive and advantageous aspects of the
graphene-encapsulated NP biosensor is the ease of fabrication
and measurement (Figure 2). The device was fabricated using
photolithography, followed by a lift-off process, both of which
are well-established.[33,34] We first generated gold electrodes on
a silicon oxide substrate using photolithography and lift-off.
To generate arrays of GO-NPs, we patterned the photoresist
(AZ 5214) on the substrates with gold electrodes using
© 2011 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
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Graphene-Encapsulated Nanoparticle-Based Biosensor for
the Selective Detection of Cancer Biomarkers
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Figure 1. Fabrication process of biomolecular sensor based on graphene-coated NPs. a) Schematic diagram of GO assembly on amine-functionalized
NPs and TEM image of NPs coated with GO. b) Fabrication of a metal electrode on the oxide substrate and surface modification for the assembly of
GO-NP. c) Photoresist (PR) patterns on the metal electrodes. d) GO-NP assembly in the centrifuge tube. e) Removal of PR patterns and reduction of
GO coated on the NP surface.
Figure 2. Reduced GO-NP patterns and the electrical property of the rGO-NP device. a) SEM images of rGO-NPs assembled on a large area. b) The
SEM images of biosensors consisting of a rGO-NP array with gold electrodes. c) The schematic diagram of measuring process of the gate effect utilizing
ionic liquid (left) and the gate effect of ten rGO-NP junctions with a 50 mA channel length (right).
2222
Adv. Mater. 2011, 23, 2221–2225
www.advmat.de
One of the key barriers to using graphene FET-based biosensors is to operate the device under physiological conditions (e.g.,
different pH and salt concentrations), in which different ionic
environments affect the conductivity of graphene FET-based biosensors.[36] To study the working conditions of the graphene FETbased biosensors in aqueous solutions, we measured the gating
effect of the rGO-NP-based biosensor using an ionic liquid gate
(Figure 2c). A typical source–drain current versus gate potential
plot was obtained in an ionic liquid, 1-butyl-3-methylimidazolium
tetrafluoroborate (BmimPF6) (see Supporting Information for the
synthesis of BmimPF6). In an ionic liquid, the high concentration
of ions renders the thickness of the diffusion layer negligible, thus
making it useful as a gate insulating layer. Silver (Ag) wire was
used as the reference electrode for the measurements in the ionic
liquid. The gating effect observed in the GO-NP devices was similar to that observed in rGO thin-film transistors that have ambipolar conduction and p-type behavior near zero gate voltage.[33,37]
In the present case, the top-gate bias was swept with ≈0.05 V s−1
sweep speed under the source–drain bias of 0.5 V (Figure 2c).
Once the device containing the rGO-NP array was optimized
for biosensing, the selective detection of HER2 and EGFR was
carried out by functionalizing the rGO-NPs with monoclonal
antibodies (mAbs) against HER2 or EGFR. The bioconjugation chemistry is well-established and involved three basic
steps.[2,10] First, the reduced GO surface was functionalized
with 4-(pyren-1-yl)butanal via π–π interactions by incubating
the device in a methanol solution (1:500) of 4-(pyren-1-yl)
butanal (see Supporting Information for synthesis) for 30 min
(Figure 3a). Second, the aldehyde groups were coupled to the
COMMUNICATION
photolithography. The exposed silicon oxide surface and gold surface were functionalized with self assembled monolayers (SAMs)
of positively charged 3-aminopropyltriethoxysilane (APTES)
and cysteamine, respectively. The SAM formation promoted the
assembly of the negatively charged GO-NPs (through electrostatic interactions).[34,35] We then employed a relatively simple
technique involving centrifugation for the uniform assembly of
GO-NPs on the positively charged SAMs. In this technique, the
substrate containing the patterned photoresist along with the
SAMs was centrifuged in a solution of GO-NPs at 2000 rpm
for 3 min in a centrifuge tube. Despite a low concentration of
GO-NPs, we were able to achieve uniform films of NPs with a
high density in a reproducible manner. This is in stark contrast
to the standard methods used for assembling NPs on surfaces.
Other methods generally rely on using larger volumes of the
solution containing higher concentrations of NPs, where contact of the NPs with the surface is mainly made through infrequent Brownian motion, which eventually causes NP assembly.
On the other hand, the centrifugation technique achieved uniform, very dense layers of graphene-encapsulated NPs over a
large area in a short time span (Figure 2a,b). We then generated a uniform NP array by removing the patterned photoresist
using acetone. The removal of photoresist did not disturb the
assembly of GO-NPs. To render the insulating GO electrically
conductive, we reduced the GO through an overnight exposure
to hydrazine vapor. This method of fabricating the device is very
powerful because it can be integrated with conventional microfabrication processes, which makes the device cost effective and
relatively easy to produce on a large scale.
Figure 3. Real-time detection of cancer marker, HER2. a) The preparation of rGO-NP device. b) Surface functionalization of rGO for immobilizing the
antibody. c) Measuring conductance of the devices when the target protein is introduced. d) The sensitivity of the biosensor (relative conductance
change,%) in response to the concentration of HER2 with VDS (voltage drain to source) = 1 V and Vg (gate voltage) = 0 V. e) The selectivity of the
biosensor in response to PBS buffer, BSA with 50 μg mL−1 and HER2 (100 pM and 1 μM). f) Sensor sensitivity (relative conductance change,%) as a
function of the HER2 concentration with VDS = 1 V and Vg = 0 V. All experiments were performed multiple times (sample number, n = 30) to collect
statistical data (with error bars) and confirm the reproducibility and robustness of the biosensing system.
Adv. Mater. 2011, 23, 2221–2225
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2224
amine groups of the monoclonal HER2 or EGFR antibodies
(1:50) through reductive amination in the presence of 4 mM
sodium cyanoborohydride in phosphate buffered saline (PBS;
pH 7.4) for 2 h (Figure 3b). Third, unreacted aldehyde groups
were blocked using 100 mM ethanolamine in a similar manner
to prevent non-specific interactions of proteins. Finally, we
rinsed the device in a continuous flow of PBS (pH 7.4) for
10 min. We show that the surface chemistry used in the device
plays a crucial role in achieving highly selective and sensitive
detection of HER2 or EGFR protein (Figure 3c,d). Furthermore,
due to the large surface-to-volume ratio of the rGO-NPs, the
biosensors were highly efficient as compared to the thin-filmtransistor-based biosensors.
The sensitivity of the rGO-NP devices, functionalized with
HER2 mAbs, was determined by measuring the changes in
conductance as the solution concentration of HER2 was varied
from 10 fM to 1 μM (Figure 3d). In all experiments, only 1 μL of
each solution was added onto the device. Representative timedependent data show that on the addition of a 10 fM solution of
HER2, no change in conductance was observed. However on
increasing the concentration to 1 pM, a decrease in conductance
of the p-type rGO-NP device was observed due to the binding of
HER2 to the mAbs. As the concentrations of the solutions were
subsequently increased, a concentration-dependent decrease
in the conductance of the rGO-NP device was observed. Thus,
the detection limit of the biosensor was observed to be 1 pM
in a solution containing only HER2 protein, which is a significant improvement over thin-film-transistor-type sensors based
on graphene.[7,14,15] The observed change in electrical conductivity can be attributed to the p-type characteristics of the
rGO-NP FET-based sensors because the amine groups on the
protein surface are positively charged. Binding of these positively charged target biomolecules, such as HER2 or EGFR, to
the rGO surface will induce positive potential gating effects that
generate reduced hole density and electrical conductance.
To test the selectivity of the graphene FET-based biosensors,
we further investigated the selective detection of the device in
competitive binding studies with bovine serum albumin (BSA)
(Figure 3e). Time-dependent conductance measurements
recorded on the rGO-NP devices functionalized with HER2
mAbs showed no change in conductance upon addition of PBS
and 50 μg mL−1 BSA. However, upon addition of 1 μL of 100 pM
solution of HER2 to the BSA solution on the device, a rapid and
sharp change in conductance was observed, demonstrating the
high selectivity of the device. Upon adding the 1 μM solution
of HER2, the conductance further decreased rapidly and drastically. In spite of the presence of a solution with a very high concentration of BSA (50 μg mL−1), the detection limit of the target
protein, HER2, was 100 pM, clearly demonstrating the remarkable sensitivity and selectivity of the rGO-NP biosensor.
Figure 3f shows the sensitivity (relative conductance change)
of the biosensor as a function of the HER2 concentration. The
lowest HER2 concentration level that could be detected is 1 pM,
which shows a decrease in conductance (3.9%). Similar to the
non-linear behavior of CNT FET-based sensors,[38–40] the sensor
responses increase non-linearly with the increase in the HER2
concentration from 1 pM to 1 μM, which clearly shows that the
sensor response is due to the binding of HER2 to the HER2
mAbs.
In addition to HER2, we similarly investigated the sensitivity and selectivity of the device for detecting EGFR. We functionalized the device with EGFR mAbs and observed the change
in conductance upon addition of EGFR solution. The trend in
conductance change was similar to that observed with HER2,
with the detection limit being 100 pM for EGFR and 10 nM in
the presence of BSA (50 μg mL−1; see Supporting Information, Figure S2). We believe the slight decrease in sensitivity for
detecting EGFR (relative to HER2) might be due to the difference
in binding affinities of the two mAbs to their respective proteins.
However, the result demonstrates the capacity of the biosensor to
detect different biomarkers in a sensitive and selective manner.
In conclusion, we have demonstrated the application of a
graphene-encapsulated NP-based biosensor for highly selective and sensitive detection of cancer biomarkers by using surface chemistry principles combined with nanomaterials and
micro- and nanofabrication techniques. The novel 3D structure of graphene-encapsulated NPs significantly increases the
surface-to-volume ratio in FET-type biosensors, thereby
improving the detection limits (1 pM for HER2 and 100 pM for
EGFR) for the target cancer biomarkers. In addition, we demonstrated the highly selective nature of the biosensor as we
detected low concentrations of the target cancer biomarkers in
the presence of a highly concentrated BSA solution. The ease
of fabrication and biocompatibility, along with excellent electrochemical and electrical properties of graphene nanocomposites,
makes the graphene-encapsulated NP-based biosensor an ideal
candidate for future biosensing applications in a clinical setting.
Preparation of Reduced Graphene Oxide and SiO2 NPs: GO was
obtained from SP-1 graphite utilizing the modified Hummer method.[27]
For the GO assembly on the surface of NPs, the GO suspension was
injected into the nanoparticle solution for 10 min, and GO-NPs were
separated from GO solution using a centrifuge. SiO2 NP (100 nm)
solution was purchased from Corpuscular Inc. For GO-NP assembly,
the photoresist-patterned substrate was placed in the NP solution
and GO-NPs were assembled on the substrate by applying centrifugal
force. After the deposition of GO-NPs on the substrate, the GO on the
NP surface, having the low conductance, was reduced to graphene by
exposure to hydrazine vapor overnight.
Surface Molecular Pattering: 3-Aminopropyltriethoxysilane (APTES)
and cysteamine molecules, used to form SAMs, and the solvents were
purchased from Sigma-Aldrich. For the patterning of APTES SAM on SiO2,
the photoresist (AZ5214) was first patterned by photolithography using a
short baking time (<10 min at 95 °C). The patterned substrate was placed
in the APTES solution (1:500 (v:v) in anhydrous hexane) for 7 min. For the
patterning of APTES on the on SiO2 layer, the substrate with photoresist
patterns was placed in an APTES solution (1:500 (v:v) in anhydrous
hexane) for 10 min. The photoresist was then removed with acetone.
Metal Deposition and Measurement of Graphene Devices: For the
electrode fabrication, the photoresist was first patterned on the
substrate. Ti/Au (10/30 nm) was then deposited on the substrate and
the remaining photoresist was then removed with acetone for the lift-off
process. A Keithley-4200 semiconductor parameter analyzer was used
for measurement and data collection.
Supporting Information is available from the Wiley Online Library or
from the author.
Adv. Mater. 2011, 23, 2221–2225
www.advmat.de
This work was supported by the NIH Director’s Innovator Award
[(1DP20D006462–01), K.-B. L.] and the N.J. Commission on Spinal Cord
Injury grant [(09–3085-SCRE-0), K.-B. L.]. K.S.K. acknowledges support
from NRF (National Honor Scientist Program: 2010-0020414).
Received: January 4, 2011
Revised: February 16, 2011
Published online: April 5, 2011
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COMMUNICATION
Acknowledgements
2225
Multifunctional Nanomaterials
Graphite-Coated Magnetic Nanoparticles as Multimodal
Imaging Probes and Cooperative Therapeutic Agents
for Tumor Cells
Joung Kyu Park, Jongjin Jung, Prasad Subramaniam, Birju P. Shah, Cheoljin Kim,
Jong Kyo Lee, Jee-Hyun Cho, Chulhyun Lee, and Ki-Bum Lee*
An effective therapeutic approach against cancer typically
requires the combination of several modalities, such as chemotherapy, radiation, and hyperthermia. In this regard, the
development of multifunctional nanomaterial-based systems
with combined therapeutic and molecular imaging capabilities has shown great potential but has not been fully explored.
In particular, magnetic nanomaterials have been at the forefront of cancer research as noninvasive imaging probes as
well as multifunctional therapeutics.[1] For example, magnetic
nanoparticles (MNPs) with appropriate surface modifications
have been successfully applied to deliver therapeutic biomolecules, such as anticancer drugs, antibodies, and siRNAs, to
target tumor cells or tissues.[2] Moreover, the unique physical
and chemical properties of these magnetic nanostructures
have enabled their wide applications in cancer imaging and
therapy, including magnetic resonance imaging (MRI) and
hyperthermia.[3] Promising advances have been made in
synthesizing multifunctional MNPs from various materials,
including metals,[4] metal oxides,[5] metal alloys,[6] and metal–
graphitic-shell nanomaterials,[7] with different properties.
However, current studies are mostly focused on the synthesis
and characterization of materials with limited demonstration of their biomedical applications, like molecular imaging
and therapy. As a result, research efforts towards developing
MNP-based multimodal therapeutics to control the tumor
microenvironment are highly limited and have not been
fully explored. Therefore, in order to address the challenges
Dr. J. K. Park, J. Jung, P. Subramaniam, B. P. Shah, Dr. C. Kim,
Prof. K.-B. Lee
Dr. J. K. Park, Dr. J. K. Lee
Center for Nano-Biofusion Research
Korea Research
Institute of Chemical Technology
Daejon 305–600, Korea
Dr. J.-H. Cho, Dr. C. Lee
Division of Magnetic Resonance Research
Korea Basic Science Institute
Ochang 363–883, Korea
DOI: 10.1002/smll.201100012
small 2011, 7, No. 12, 1647–1652
of MNP-based therapeutics, as well as to narrow the gap
between current nanoparticle-based multimodal imaging
approaches and their clinical applications, there is a clear
need to synthesize effective chemotherapeutic MNPs and
to develop multimodal therapies for targeting specific oncogenes, thereby activating/deactivating corresponding key signaling pathways.
In this Communication, we describe the novel synthesis
and a systematic in vitro evaluation and application of multifunctional magnetic nanoparticles (MNPs) with an iron cobalt
core and a graphitic carbon shell (FeCo/C) for the targeted
delivery of small interfering RNA (siRNA) to tumor cells with
a concomitant hyperthermia-based therapy, thereby cooperatively inhibiting proliferation of and inducing apoptosis in
tumor cells (Figure 1). In parallel, we also demonstrate that
our MNPs can be used as highly sensitive magnetic resonance
and Raman imaging probes. As a model study, we used glioblastoma multiforme (GBM) cell lines, the most malignant
and difficult-to-treat brain tumor cells. We hypothesized that
the targeted delivery of our siRNA–MNP constructs against
the oncogenic receptor (EGFRvIII) and subsequent hyperthermal treatment would selectively, as well as cooperatively,
damage the tumor cells, resulting in the synergistic inhibition
of tumor-cell proliferation and the induction of apoptosis via
the deactivation of the PI3K/AKT signaling pathway. Hence,
these MNP-based therapeutics could potentially be used for
the simultaneous imaging and therapy of malignant tumors
both in vitro and in vivo.
Recent efforts in cancer therapy have demonstrated the
application of hyperthermia, which involves localized heating
of cancerous cells or tissues, as an adjuvant to chemotherapy
and radiation to improve their efficacy.[8] Hyperthermia typically involves increasing the local temperature of the tumor
region to 42–46 °C over a given time period, ultimately
resulting in apoptosis of the heat-sensitized cancer cells.[9]
One of the best methods of achieving a localized hyperthermal effect is to deliver MNPs to the target cells and
subsequently apply electromagnetic fields following their cellular uptake/localization.[10] Furthermore, hyperthermia and
its downstream effects can be significantly enhanced by the
concomitant use of other cancer therapies, including radiation and drug/gene delivery and vice versa.
To
develop
cooperative
(hyperthermia
and
siRNA delivery) therapeutic systems based upon MNPs, we
1647
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J. K. Park et al.
11-nm FeCo ≈235 emu g−1), has a high
Curie temperature, and has high magnetic
anisotropy energies, all of which are critical in order to enhance their potential for
biomedical applications, such as MRI and
hyperthermia; ii) the thickness of the outer
graphite-shell layers can be controlled by
our synthetic method, which would lead
to an improved Raman signal intensity for
detecting cancerous cells; and iii) the nanoparticles are chemically inert due to the
presence of a graphitic carbon shell[11] and
can be made biocompatible by appropriate
surface modifications (e.g., dextran-ligand
coating). Moreover, compared to conventional methods[1b,12] for the synthesis
of core–shell metal-alloy magnetic nano[13]
Figure 1. Magnetic FeCo–graphite nanoparticles for multimodal imaging and targeted materials, such as electric-arc discharge,
tumor therapy. a) Detailed structure of the MNPs depicting the highly magnetic FeCo core, high-temperature thermal decomposiprotective Raman-active graphite shell, and the biocompatible dextran coating. b) Inhibition tion,[14] and chemical vapor deposition
of proliferation and induction of apoptosis via combined siRNA delivery and hyperthermia (CVD),[11] our novel hydrothermal synusing siRNA–FeCo/C NP constructs.
thetic approach has several advantages,
such as relatively milder synthetic condisynthesized graphitic-carbon-protected iron cobalt (FeCo/C) tions, low environmental impact, cost effectiveness, ease of
nanoparticles (7 and 11 nm in diameter) with a body-centered scalability (see Figure S3 in the SI), and the exclusion of toxic
cubic (bcc) crystalline structure using hydrothermal synthetic solvents and size-separation techniques.
methods followed by an annealing process at 1000 °C. HighAnother critical step to realize the full potential of our
resolution transmission electron microscopy (HR-TEM) and multimodal FeCo/C NPs for in vitro/in vivo biomedical appliX-ray diffraction (XRD) confirmed the excellent chemical/ cations (e.g., targeted drug/gene delivery, MRI, or hyperphysical properties of our FeCo/C NPs, such as monodisper- thermal therapy) is to make the nanoparticles biocompatible
sity, narrow size distribution of the nanoparticles, and the presence of a crystalline
bcc FeCo core (Figure 2). The graphiticcarbon shells surrounding the FeCo core
were confirmed by Raman spectroscopy
analysis and HR-TEM. Furthermore, the
thickness of the graphitic-carbon coating
could be monitored by the intensity of
the Raman signal (marked by arrows in
Figure 2d). We also characterized the magnetic properties of our FeCo/C NPs using
a superconducting quantum interference
device (SQUID). Our FeCo/C NPs were
found to display remarkable superparamagnetic properties at room temperature,
as suggested by the significantly higher
value of the saturation magnetization
(Ms) for our FeCo/C NPs as compared to
that of the commercially available Fe3O4
(Figure 2e). This was attributed to the
higher magnetic moments of the FeCo/C
NPs as a result of the high-temperature Figure 2. Structural and magnetic properties of the FeCo/C nanoparticles. a) TEM image
annealing, as shown in Figure S2 in the of the 11-nm nanoparticles (scale bar = 20 nm). b) HR-TEM image of the nanoparticles
showing the crystalline lattice structure of the FeCo core. c) Powder XRD for the 7- and 11-nm
Supporting Information (SI).
In this work, our FeCo/C MNPs have nanoparticles showing the presence of a bcc crystalline core and graphitic shell. d) Raman
spectrum (excitation at 632.8 nm) of the 11-nm nanoparticles with single and multiple carbon
several advantages over other conven- shells (marked by arrows) showing the D and G bands of graphitic carbon (scale bar = 2 nm).
tional magnetic nanoparticles, such as e) Room-temperature magnetization versus applied magnetic field for 11-nm FeCo/C NPs
FePt, Fe2O3, and Fe3O4: i) FeCo exhibits (red symbols) and Fe3O4 (black symbols). Note that no hysteresis loop exists owing to the
an excellent magnetization value (e.g., superparamagnetism of the nanoparticles.
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small 2011, 7, No. 12, 1647–1652
Graphite-Coated Magnetic Nanoparticles as Multimodal Imaging Probes
and modify the surface for achieving targetspecific intracellular delivery.[15] Several
strategies, including coating with molecules
such as dextran, have been successfully
used for functionalizing magnetic nanoparticles and rendering them biocompatible.[16] Hence, we synthesized a series of
dextran derivatives (see Figure S4a in the
SI) and used them to coat our FeCo/C NPs
(see Figure S4b in the SI). The dextrancoated FeCo/C NPs were found to have
excellent colloidal stability and the hydrodynamic diameter was found to be ≈150 nm
(see Figures S5 and S6 in the SI). These
dextran-modified FeCo/C NPs were then
conjugated to specific cancer-targeting
biomolecules, such as epidermal growth
factor receptor (EGFR) antibodies and
cyclic-RGD (c-RGD) peptide, to target
glioblastoma cells and to improve their
intracellular uptake. The conjugation of
biomolecules such as EGFR antibodies
and c-RGD to our FeCo/C NPs not only
increases transfection of our nanoparticles
via receptor-mediated endocytosis but can
also selectively target the glioblastoma
cells by binding to receptors (EGFR and
integrins) known to be overexpressed in
glioblastoma cells.[17]
Highly sensitive multimodal imaging
tools like MRI, computed tomography
(CT),
positron-emission
tomography
(PET), and Raman imaging, which rely on
nanomaterials as molecular probes, have
gained much attention as diagnostic tools
for specific cancers, such as breast and
brain cancers.[18] In order to investigate
the capability of our FeCo/C NPs as multimodal imaging nanoprobes, we focused on
two important imaging methods for cancer,
in vivo MRI and in vitro Raman imaging,
both of which provide complementary
information about tumor microenvironments at the tissue level and/or cellular Figure 3. MR measurements and imaging of FeCo/C NPs and Resovist. a) Concentrationdependent T2 measurements of FeCo/C NPs and Resovist solutions. b) T2-weighted MR images
level.[3b,18b] In order to test the capability
of various nanoparticle solutions. The FeCo/C NPs show higher MR image contrast (several
of our MNPs as MRI contrast agents, the fold) as compared to Resovist, a traditional MRI contrast agent. c) T -weighted MR images of a
2
transverse relaxation times (T2) of the rat before (t = 0 min) and 30 min after (t = 30 min) injection of FeCo/C NPs (left) and Resovist
water protons were measured in a specific (right) into a rat’s tail vein. The nanoparticles were seen to localize in the liver (green arrow),
magnetic field and their values were com- spleen (blue arrow), and kidneys (red arrow) of the animal. Also, FeCo/C NPs show a higher
pared to those of the commercially avail- imaging contrast at a lower concentration (0.25 mg of Fe) as compared to Resovist (2.5 mg
able MRI contrast agent Resovist (Fe3O4, of Fe). d) Raman spectra of U87-EGFP cells (marked by arrows) treated with 11-nm FeCo/C
NPs. The spectra shows the capability of the FeCo/C NPs to be used as imaging agents at the
from Roche). Both the 7- and 11-nm single-cell level (see Figure S8 in the SI for detailed Raman imaging).
FeCo/C NPs exhibited a higher relaxivity
coefficient (r2, 252 and 392 mm−1 s−1, respectively) and enhanced T2-weighted MR contrast (Figure 3a,b) nanoparticles accumulated in the liver, spleen, and kidneys
as compared to Resovist (r2 = 140 mm−1 s−1). Characteriza- and their imaging contrast was significantly higher (about
tion of our biocompatible dextran-coated FeCo/C NPs as in 10 times) than that of Resovist (Figure 3c). We also monitored
vivo MRI agents was performed by injecting the MNPs into the in vivo biodistribution and MRI contrast of these FeCo/C
a rat’s tail vein. Preliminary MRI studies showed that our NPs over 10 days. It was found that the NPs were retained
small 2011, 7, No. 12, 1647–1652
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communications
J. K. Park et al.
in the liver and spleen and that there was
no significant loss of imaging contrast
over this time period (see Figure S7 in the
SI). Furthermore, the analysis of in vitro
Raman imaging results, wherein confocal
microscopy was used to observe the NPinternalized U87 cell lines, confirmed that
the intensity of the D and G bands of the
graphitic-carbon shells was indeed proportional to the thickness of the carbon
shells on the FeCo/C NPs (Figure 3d). This
data strongly suggests that our FeCo/C
NPs have a well defined correlation and
sensitive response when used as Raman
imaging probes at the single-cancer-cell
level. These preliminary in vitro and in
vivo studies showed no visible cytotoxicity due to the degradation of the FeCo/C
NPs. It has been reported that FeCo/C
NPs are safely excreted over time through
the biliary system after being taken up by
the liver and spleen.[19] Hence, it is highly Figure 4. In vitro hyperthermia and siRNA-delivery studies of the FeCo/C NPs. a,b) U87–EGFP
unlikely that these FeCo/C NPs would cell death induced by hyperthermia in cocultures of the highly tumorigenic U87–EGFP cells
pose any serious toxicity issues when used (marked by arrows) and the less-tumorigenic PC-12 cells (marked by arrows) via the targeted
in vivo. A detailed toxicological evaluation delivery of FeCo/C NPs to the U87 cells. Fluorescence images (a) and quantitative analysis
of these MNPs in animal models would be (b) show that significant hyperthermia-induced cell death is observed in U87 cells, while the
PC-12 cells keep proliferating with time. (Note that the number of cells at 0 h was taken to
forthcoming in the future.
be 100% and the cell counts at other time points were normalized to this value.) Annexin V
Having demonstrated the potential of assays for detection of early apoptosis proved that the cell death was caused by localized
our FeCo/C NPs as multimodal imaging hyperthermia-induced apoptosis rather than necrosis (see Figure S12 in the SI). c) MTS assay
probes, we then focused on evaluating demonstrating the synergistic inhibition of proliferation and induction of cell death by the
their ability to be used as targeted hyper- combined siRNA and hyperthermia treatment using siRNA-FeCo/C NPs in U87–EGFRvIII cells
thermia agents for tumor therapy in vitro. as compared to individual treatments and nontreated controls. The scale bar in all images
We hypothesized that our FeCo/C NPs is 100 μm.
would be more efficient hyperthermia
agents than Fe3O4 NPs of similar size due to their high mag- incubated in cocultures of glioblastoma cells (U87–EGFP),
netization and therefore would be more effective in inhibiting which had been genetically labeled with enhanced green fluproliferation and inducing apoptosis of target brain tumor orescent protein (EGFP) and present EGFRs on their memcells (bTCs). To test their efficacy as targeted hyperthermia branes and several other less tumorigenic cells, such as PC-12
agents, we first measured the minimum time required to and astrocytes, which tend to have low expression levels of
attain the therapeutic temperature (≈43 °C) when placed in integrins and EGFRs. Our data clearly indicates that the sura homogeneous magnetic field. The specific absorption rates face modification of our FeCo/C NPs with c-RGD peptide or
(SARs) of FeCo/C and Fe3O4 NPs (≈69 W g−1 for FeCo/C EGFR antibodies resulted in their selective cellular uptake
and 13 W g−1 for Fe3O4), as derived from the plots of temper- by the target bTCs, as compared to PC-12 or astrocytes (see
ature versus time in aqueous solutions under a 334 kHz mag- Figure S11 in the SI). Following intracellular uptake of the
netic field (which is the optimum frequency range estimated aforementioned NP constructs, the cells were exposed to
by the Neel and Brownian relaxation-time simulations[20]), an alternating current (AC) magnetic field for 15 min. Sigindicated that the time required for our FeCo/C NPs to reach nificant inhibition of proliferation and hyperthermia-induced
the therapeutic temperature was ten times shorter than that cell death was observed mainly in the U87 cells while the
of Fe3O4 NPs (see Figure S9 in the SI). We next evaluated less-tumorigenic PC-12 cells largely continued proliferating
the concentration-dependent cytotoxicity of our FeCo/C NPs with time (Figure 4a,b).
by serial-dilution investigations. From this study, the range of
In the past decade, there has been considerable interest
concentrations inducing negligible cytotoxic effects on cells in the development of nanoparticle-based siRNA-delivery
was identified and the concentrations (95% cell viability at methods for cancer therapy. RNA interference (RNAi)
300 μg mL−1 after ≈24 h post transfection) within this range involves the use of siRNAs to selectively mediate the cleavage
were used for our subsequent experiments (see Figure S10 in of complementary mRNA sequences and thus regulate target
the SI). We then sought to precisely increase the temperature gene expression.[21] In combination with other chemotheraof tumor cells while minimizing the exposure of other cells peutic methods like hyperthermia, RNAi could prove to be
to hyperthermic temperatures. For this purpose, FeCo/C NPs a powerful tool to manipulate the tumor microenvironment.
functionalized with c-RGD peptide or EGFR antibodies were In order to deliver siRNA to the target cells, the FeCo/C
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small 2011, 7, No. 12, 1647–1652
Graphite-Coated Magnetic Nanoparticles as Multimodal Imaging Probes
NPs were conjugated to the siRNA using polyethyleneimine
(PEI) as a cationic polymer via a layer-by-layer approach[22]
(see Figure S13 in the SI). We initially optimized the knockdown efficiency of our siRNA–FeCo/C NP construct by the
suppression of EGFP in U87–EGFP cells. The decrease in
green fluorescence intensity due to siRNA-mediated knockdown of EGFP was monitored to assess the knockdown efficiency (≈80% after 3 days of transfection) of our siRNA–NP
constructs (see Figure S14 in the SI). Once the conditions
for the siRNA delivery and knockdown were optimized, we
focused on inhibiting the proliferation and inducing apoptosis of U87–EGFRvIII cell lines overexpressing the oncogenic EGFRvIII gene. EGFRvIII is a mutant type of EGFR
as well as an oncogenic receptor that is highly expressed only
in tumor cells, and not in normal cells.[23] The effect of the
knockdown of the EGFRvIII oncogene using our siRNA–
FeCo/C NP constructs was assessed using a microscope at
96 h post transfection (see Figure S14 in the SI). We hypothesized that the knockdown of the target oncogene, EGFRvIII,
with our multifunctional siRNA–FeCo/C NP constructs, combined with hyperthermia, would lead to a cooperative inhibition of tumor-cell proliferation and increase in cell death.
The U87–EGFRvIII cells were incubated with our FeCo/C
NPs modified with siRNA against EGFRvIII, followed by
hyperthermia treatment for 5 min at 72, 96, and 120 h post
siRNA treatment. Quantitative analysis based upon the
3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)2-(4-sulfophenyl)-2H-tetrazolium assay, commonly known
as the MTS assay (Figure 4c) showed that treatment of cells
with our siRNA–MNPs against EGFRvIII followed by hyperthermia induced significantly more cell death as compared to
the controls. This could be attributed to the fact that silencing
of the EGFRvIII oncogene results in a decrease in expression
of the focal adhesion proteins that make the cells more susceptible to heat, thereby leading to a synergistic increase in
cell death.[24] Our study clearly demonstrates that the appropriate combination of various therapeutic modalities using
our FeCo/C NPs can significantly enhance the therapeutic
efficacy relative to the individual components.
In summary, this work provides an early demonstration of
integrating multimodal imaging with combined hyperthermia
and siRNA-based therapy in malignant tumor cells using
highly efficient FeCo/C NPs. Our FeCo/C NPs have successfully been demonstrated to have a well defined correlation
and a fast and sensitive thermal response to the strength of
the applied magnetic field. At the same time, our FeCo/C NPs
could be developed as novel therapeutic and diagnostic tools
for cancer research. The ability to functionalize the graphitic
surface of the FeCo/C NPs with targeting ligands and biomolecules would be critical to realize the potential of
nanoparticle-based diagnosis and therapy of various cancers.
It should be noted that our FeCo/C NPs showed excellent
MRI contrast results compared to the conventional MRI
contrast agents as well as enabling us to collect the Raman
spectral information at the single-cell level. More importantly, the use of our FeCo/C NPs for site-specific and localized hyperthermia in conjunction with siRNA therapy against
oncogenes would greatly complement and enhance the effects
of other therapeutic modalities, including gene therapy and
small 2011, 7, No. 12, 1647–1652
chemotherapy, thereby reducing the dose of anticancer drugs,
mitigating their toxic side effects, and effectively circumventing drug resistance in cancers.
or from the author.
Acknowledgements
We thank Prof. Huixin He for helping us with the hyperthermia
studies, Prof. Sang-Wook Cheong and Prof. Martha Greenblatt for
their support for the SQUID measurements and the IAMDN center
for allowing us to use their high-resolution TEM facility. We are
grateful to the KBLEE group members (Aniruddh Solanki, Shreyas
P. Shah, and Michael Koucky) for their valuable suggestions for
the manuscript. J.K.P. acknowledges the research program at
KRICT and Dr. Young-Duk Suh and Dr. Kee-Suk Jun for the Raman
imaging. K.-B.L. acknowledges the NIH Director’s Innovator Award
(1DP20D006462–01) and is also grateful to the N.J. Commission
on Spinal Cord Research grant (09–3085-SCR-E-0).
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a) H. Lee, E. Lee, D. K. Kim, N. K. Jang, Y. Y. Jeong, S. Jon, J. Am.
Chem. Soc. 2006, 128, 7383–7389; b) X. M. Qian, X. H. Peng,
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Received: January 4, 2011
Revised: March 1, 2011
Published online: May 11, 2011
small 2011, 7, No. 12, 1647–1652
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Generation of a Library of Non-Toxic Quantum Dots for
Cellular Imaging and siRNA Delivery
Prasad Subramaniam, Seung Jae Lee, Shreyas Shah, Sahishnu Patel,
Valentin Starovoytov, and Ki-Bum Lee*
Dissecting the spatio-temporal interaction of biomolecules
inside cells at the subcellular level is an important facet of
molecular cell biology and chemical biology. Over the last few
decades, a variety of fluorescent molecular probes have been
developed to investigate these complex bio-interactions for both
in vitro and in vivo cellular imaging and subcellular detection. In particular, semiconductor nanoparticles like quantum
dots (QDs) have shown great potential as nanoparticle-based
fluorescent probes due to their excellent physiochemical properties, which allow them to overcome the limitations of conventional fluorescent probes such as organic dyes and fluorescent
proteins.[1] These unique attributes of QDs have proven to be
crucial in elucidating the intricate interactions through which
small molecules and biomolecules (e.g., proteins, peptides and
nucleic acids) bind to their targets in specific signaling cascades.[2] While QDs are excellent molecular probes, they also
can be used as effective delivery vehicles for several therapeutic
biomolecules. For example, the use of QDs for the simultaneous imaging and delivery of small interfering RNA (siRNA)
for selectively knocking-down target oncogenes in tumor cells
has been successfully demonstrated.[2c,3]
However, the major limiting factor in harnessing the maximum potential of QDs as multifunctional imaging probes and
delivery systems is their inherent cytotoxicity; most of the wellestablished QDs are composed of highly toxic elements, such
as cadmium (Cd), selenium (Se) or tellurium (Te).[2b,4] Owing
P. Subramaniam, S. J. Lee, S. Shah, S. Patel,
Prof. K.-B. Lee
Institute for Advanced Materials
Devices and Nanotechnology (IAMDN), Rutgers
E-mail: kblee@rutgers.edu; http://rutchem.rutgers.edu/∼kbleeweb/
S. J. Lee
Center for Nano-Bio Fusion Research
Korea Research Institute of Chemical Technology
Daejeon 305-600, Korea
V. Starovoytov
Department of Cell Biology and Neuroscience, Rutgers
The State University of New Jersey,
Prof. K.-B. Lee
School of Medicine
Kyung Hee University
Seoul, South Korea
DOI: 10.1002/adma.201201019
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to this obstacle, the wide applications of QDs are currently
delayed and the main focus of QD imaging has been limited
to the cell and small animal studies. In response to the above
issues, the recent development of I-III-VI2 type QDs[5] like
AgInS2,[5c] CuInS2[5b,5d] and ZnS-AgInS2[5e] offers better control
of band-gap energies and demonstrates the great potential of
these QDs as non-toxic molecular probes. For instance, several
research groups have successfully synthesized ZnS-AgInS2
(ZAIS) QDs through the decomposition of single source precursors using thermal,[5e] hydrothermal,[6] photothermal[7] and
microwave-assisted methods.[8] Yet, these conventional synthetic
methodologies for preparing these I-III-VI2 type QDs have several shortcomings such as high reaction temperatures, poor
control of growth rates, long reaction times, difficulty of high
throughput synthesis, and the need for complicated synthetic
procedures to prepare QDs with different emissions profiles,
all of which would be critical in investigating the diversity and
dynamic processes of multiple biomarkers in cancer and stem
cells. Thus, in order to harness the full potential of QDs as biological imaging probes as well as drug delivery platforms for
clinical and translational research, there is an urgent need to
develop a simple and straightforward methodology that affords
both the synthesis of non-toxic QDs and the versatility of generating a library of QDs with tunable properties favoring their use
as imaging agents in biology.
To address the aforementioned issues, we developed a novel
sonochemical approach for the high throughput synthesis of
a library of biocompatible ZnxS-AgyIn1-yS2 (ZAIS) quantum
dots with tunable physical (photoluminescence, PL) properties,
thereby allowing them to be used as multifunctional nanoparticles for the simultaneous imaging and effective delivery of
siRNA to brain tumor cells with negligible cytotoxicity. These
ZAIS QDs also proved to be useful for imaging stem cells,
which are otherwise quite sensitive towards nanomaterial-based
molecular imaging probes (Figure 1). A sonochemical synthetic
method uses ultrasound irradiation to drive the main synthetic
reaction process. With the rapid growth of its use for applications in material science, the sonochemical synthetic approach
is particularly attractive for the preparation of novel nanomaterials. Its advantages include a fast reaction rate (for e.g., its possible to generate a whole library of nanoparticles of several compositions in a span of few hours), controllable reaction conditions and the ability to form nanoparticles with uniform shapes,
narrow size-distributions and high purity in relatively less time
and at ambient conditions. Specifically, the synthetic methodology used for the preparation of our ZAIS QDs involved the
sonochemical decomposition of the organometallic precursor
(AgyIn1-y)Znx(S2CN(C2H5)2)4 at ambient conditions. Compared
Adv. Mater. 2012, 24, 4014–4019
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Figure 1. Synthesis of a library of ZnS-AgInS2 QDs for simultaneous imaging and delivery of siRNA. A) Synthetic procedure to obtain a library of
dodecylamine-capped hydrophobic ZAIS QDs. B) Conversion of the the hydrophobic ZAIS QDs into water-soluble ones via ligand exchange with
3-mercaptopropionic acid (MPA) followed by attachment of siRNA using a layer-by-layer approach. C) Delivery of the siRNA against EGFP using ZAIS
QD-siRNA conjugates into brain cancer cells overexpressing EGFP.
QD) to 700 nm (Red QD) by just varying the mole ratios of
zinc (Zn), silver (Ag), and indium (In) in the precursor solutions (Figure 2A). It is worthwhile to note that as compared to
conventional CdSe or CdTe QDs, which show size-dependent
fluorescence properties, our composition-dependent tunable
fluorescence of the ZAIS QDs presents a unique advantage
for obtaining QDs with emission in the near-UV range (blue
or blue-green emission). This observation can be explained
by the fact that a blue-colored CdSe QD needs to be less than
2.0 nm in size, which is practically impossible to obtain using traditional thermal
decomposition techniques. After assessing
the optimum conditions for obtaining the
desired emission profiles, we observed that
for a given concentration of precursor ions,
ultrasonication for 5 minutes at 20 kHz and
200W output power, gave the highest emission intensity (data not shown). Figure 2B
depicts a 3D heat map which summarizes
the physical and chemical properties of the
partial ZAIS QD library. For the different
combinations of the precursor elements (i.e.
Zn, Ag, and In), the 3D heat map shows the
Figure 2. Photoluminescent properties of the ZAIS QD library. A) Representative fluorescent
emission wavelength and the corresponding
image of the entire library of ZAIS QDs with varying compositions (ZnxS-AgyIn1-yS2) synthesized
photoluminescence (PL) peak intensities of
via the sonochemical approach. B) Heat map depicting the PL intensity (z axis) vs. Zn and
Ag/In concentrations (x and y axis) for select compositions of the ZAIS nanoparticle library. selected ZAIS QDs under UV irradiation
(λex = 365 nm). This data analysis not only
Column color indicates the maximum emission wavelength.
to the traditional synthetic methods for preparing QDs, this
approach does not require high-temperature and high-pressure
conditions (See Experimental Section for the detailed synthetic
information). The resulting ZAIS QDs exhibited intense emission at room temperature, regardless of the size of the particles
(Figure 2). More interestingly, the energy gap of the ZAIS QDs
and their emission wavelength can be controlled by varying
the concentration of each precursor element. For instance, the
emission of the ZAIS QDs could be tuned from 480 nm (Blue
Adv. Mater. 2012, 24, 4014–4019
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provides the key characteristic properties for the various compositions, but it can also facilitate the selection of ZAIS QDs
with appropriate physicochemical properties for further studies
and applications. For example, examining the 3D heat map, it
is evident that ZAIS QDs without zinc (i.e. AgyIn1-yS2) had the
longest emission wavelengths (Figure 2B). We also observed
that the PL peak wavelength was blue-shifted on increasing the
Zn concentration. Furthermore, QDs obtained with 0.5 mol Ag
and 0.5 mol In, had the longest emission wavelength (λem =
697nm), with an observed blue-shift on increasing or decreasing
the Ag concentration. In conclusion, the emission profiles of
the ZAIS QDs could be easily tuned by varying the Zn or Ag
(In = 1-Ag) concentrations and hence the reported method used
to synthesize our QD library can allow scientists to investigate
the composition-dependent physicochemical properties of QDs
of interest. (See Supporting Information, Figure S1 for the
detailed information about absorption and PL spectra).
Comprehensive characterization of the resulting ZAIS QDs
was performed using several different methods. The elemental
composition of one representative ZAIS QD (Zn0S-Ag0.2In0.8S2)
was analyzed using X-Ray fluorescence spectroscopy (XRF).
It was observed that the relative atomic ratios of the constituent elements was consistent with the calculated mole ratios
(Figure 3 A), thus confirming the efficiency of the sonochemical
synthetic methodology in obtaining QDs with desired chemical
compositions. We also analyzed the crystal structures of several
compositions of the ZAIS QDs using powder X-ray diffraction
(XRD). The particles prepared with varying mole ratios of Zn,
Ag and In exhibited three broad peaks which lie in between
the diffraction patterns of the tetragonal AgInS2 and bulk cubic
ZnS crystals (Figure 3B). Furthermore, there was a clear peak
shift towards a higher angle with an increase in the amount
of Zn2+, thus indicating that the QDs obtained were a not a
just mixture of bulk ZnS and AgInS2 but a ZnS-AgInS2 solid
solution.[5e] Transmission electron microscopy (TEM) analysis
clearly revealed the spherical shape and monodispersity of our
ZAIS QDs (Figure 3C). The average size of the QDs as determined by TEM was found to be 12 ± 1.3 nm. This was further
confirmed by measuring the hydrodynamic size and the polydispersity index (PDI) of the ZAIS QDs using dynamic light
scattering (Figure S3). Additionally, the size of the QDs did
not significantly change when their composition was altered
(data not shown). We also determined the PL quantum yields
of the ZAIS QDs and compared them to the quantum yield
of the CdSe/ZnS QDs (∼0.4). It was found that the relative
quantum yields of the ZAIS QDs depended on their composition and were comparable or in some cases higher than that
of the CdSe/ZnS QDs (Figure S4). Finally, in terms of stability,
the physical properties of the ZAIS QDs such as absorption and
photoluminescence remained unchanged for two months when
stored at ambient conditions (see Figure S2).
Hence, these ZAIS QDs could be potentially
used for long term cellular labeling without
any loss of photoluminescence.
The solubility and stability of QDs in an
aqueous solution is essential for their wide
application as molecular probes in molecular
cell biology. The ZAIS QDs reported here
were functionalized with 3-mercaptopropionic acid (MPA) to render them soluble in
physiological conditions (See Experimental
section for the details regarding the surface
modification).[9] These water-soluble ZAIS
QDs were found to be extremely stable at
physiological conditions without any signs
of aggregation, even when stored for several months (Figure S5). This was further
confirmed by monitoring the PL intensity
of one of the ZAIS QDs (x = 0, y = 0.2) in
phosphate-buffered saline (PBS, pH = 7.4) at
37 ºC over a period of 6 days. The ZAIS QDs
were found to be quite stable without any significant loss of photoluminescence over the
test period (Figure S6). Another critical factor
limiting the use of conventional QDs for varFigure 3. Physical characterization of the ZnxS-AgyIn(1-y)S2 QDs. A) X-ray Fluorescence analysis ious biological applications is their inherent
[10]
This could be partially
of one of the ZAIS QD (x = 0, y = 0.2) showing the elemental composition and relative atomic cellular toxicity.
mole ratios of the constituent elements. The composition agrees to that calculated theoreti- attributed to the lack of suitable methods to
cally. B) X-ray diffraction patterns of the ZAIS QDs prepared using sonochemistry. The values of generate QDs with different compositions in
x and y are indicated in the figure. Reference patterns of bulk ZnS and AgInS2 are also shown. a high throughput manner for subsequent
Broad peak width is attributed to the amorphous nature of the ZAIS QDs. (C) TEM images of
toxicological screening. Hence, the generaone of the ZAIS QDs (x = 0, y = 0.2) clearly shows the monodispersity and narrow size distrition of a QD library and assaying their cytobution (n = 100). Inset shows the high resolution image of a single nanoparticle (Scale bar =
10 nm). (D) Photoluminescence spectra of select water-soluble ZAIS QDs (a: x = 0.6, y = 0.5.; toxicity in a simple and quick way can facilitate the selection of appropriate QDs eliciting
b: x = 0.3, y = 0.4; c: x = 0, y = 0.2; d: x = 0, y = 0.5)
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COMMUNICATION
Figure 4. Biocompatibility and cellular imaging studies of ZAIS-QDs in mammalian cells. A) Comparison of the cellular cytotoxicity of the water soluble
ZAIS QD (Zn0S-Ag0.2In0.8S2) and CdSe/ZnS QDs at different concentrations in U87 cells (A1) and hMSCs (A2). The results are presented as means ±
SD from three independent experiments. Student’s unpaired t-test was used for evaluating the statistical significance of the cytotoxicity of ZAIS QDs
(* = P < 0.001, ** = P < 0.01, *** = P < 0.05) as compared to the CdSe/ZnS QDs. B) Transmission electron microscopy of the ZAIS QD in hMSCs.
The image clearly shows the presence of the QDs (marked by blue arrows) in the cytoplasm and the nucleus (marked in red). The inset depicting the
magnified image of the QD cluster, confirms the monodisperse nature of the QDs inside the cell. C,D) Fluorescence microscopy imaging demonstrating
the uptake of the water soluble ZAIS-QD (λem = 606 nm) in U87 cells (C) and hMSCs (D).
minimum toxic effects and thus enabling the long term cellular
imaging of cancer and stem cells in vitro and in vivo. To test the
biocompatibility of our ZAIS QDs for use as imaging probes
in vitro, a cytotoxicity assay was carried out in both cancer and
stem cells (Figure 4A). The concentration-dependent cytotoxicity of the water-soluble ZAIS QDs was assessed in human
bone marrow-derived mesenchymal stem cells (hMSCs) and
human brain tumor cells (U87 glioblastoma cell line) for two
days using a cell proliferation assay (MTS). The toxicity of the
ZAIS QDs was also compared to the water-soluble CdSe/ZnS
QDs (control sample). The ZAIS QDs showed significantly
improved biocompatibility (less cytotoxic, 95% cell viability) in
both U87 cells and hMSCs even at high concentrations (upto
100 μg/mL) in comparison to the control sample (CdSe/ZnS
QDs), which were found to noticeably cytotoxic at even low concentrations. The cytotoxicity of other compositions of the ZAIS
QDs is presented in the supporting information (Figures S7
and S8). In addition, we carried out the cytotoxicity assay of
the ZAIS QDs in normal healthy cells (NIH-3T3 mouse fibroblasts). The ZAIS QDs were found to be extremely biocompatible as compared to the CdSe/ZnS QDs (Figure S9). The prolonged exposure of nanoparticles, especially QDs, to an oxidative environment (aerial oxidation or UV-induced oxidation) has
been known to catalyze their decomposition, thereby leading
to the leaching of the metal ions, which is known to induce
cytotoxic effects in cells. In order to test the cytotoxicity of our
ZAIS QDs under an oxidative environment, we subjected the
ZAIS QDs to high-intensity UV light to catalyze the oxidation
Adv. Mater. 2012, 24, 4014–4019
process. The water-soluble ZAIS QD solutions were exposed to
a UV-light source (λem = 365 nm, power density of 12 mW/cm2)
for 1–4 hours after which they were incubated with U87 cells.
It was found that even after 4h of UV-induced photo-oxidation,
the ZAIS QDs were not toxic to the cells (>85% viability, Figure
S10) which is in stark contrast to the photo-toxicity of CdSe/ZnS
QDs reported previously. For our cellular imaging experiment,
Figure 4B shows the internalization of the water-soluble ZAIS
QDs (with 606 nm emission) in human MSCs. The QDs were
primarily found to be present in the cytoplasm of the hMSCs
with a few QDs being found in the nucleus of the cells. Furthermore, fluorescent microscopy images of U87 cells (Figure 4C)
and hMSCs (Figure 4D) incubated with the water soluble
ZAIS-QDs proved that the QDs showed strong fluorescence
even when inside the cells. Collectively, our ZAIS-QDs showed
excellent biocompatibility (non-toxic) and these results can be
extended to their wide use in cellular imaging and delivery
applications, especially for stem cells, which are known to be
extremely sensitive towards nanomaterials.
Having demonstrated the potential of our ZAIS QDs as nontoxic fluorescent imaging probes, we evaluated their ability
to be used as vehicles for the efficient delivery and tracking
of siRNA in vitro. In the past decade, there has been a considerable interest in the development of nanomaterial-based
siRNA and gene delivery methods for controlling cell fate and
behavior.[11] RNA interference (RNAi) involves the use of small
interfering RNAs (siRNAs) to selectively mediate the cleavage
of complementary mRNA sequences and thus regulate target
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Figure 5. In vitro testing of the ZAIS-QD-siEGFP cell uptake and silencing efficiency in stably transfected U87-EGFP glioblastoma cells. A) Control U87EGFP cells with PEI-coated ZAIS-QD; A1) represents the phase contrast image, and A2) is the corresponding fluorescence image. B) EGFP knockdown
using the ZAIS QD-siRNA constructs; B1) Phase contrast image showing the the viability of U87-EGFP cells has not changed appreciably after the
transfection of the ZAIS QD-siRNA constructs as compared to the control cells in (A). B2) Fluorescence image clearly shows the knockdown of EGFP
in cells which have internalized the siRNA-QDs (red) after 72 h. The red fluorescence from the ZAIS QDs correlates well with the loss of the green
fluorescence in cells (indicated by yellow arrows). Scale bar is 50 μm.
gene expression. In combination with other modalities like
small molecules and peptides, RNAi could prove to be a powerful tool to manipulate the cellular microenvironments.[11b] In
this context, the use of QDs as a multimodal delivery vehicle
becomes a promising choice, thereby allowing for the efficient
delivery of siRNA and real-time tracking of the siRNA-mediated
gene knockdown.[2c] As a proof-of-concept experiment, we used
a brain tumor cell line (U87) which was genetically labeled
to express the green fluorescent protein (GFP). In order to
deliver the siRNA to target brain tumor cells expressing EGFP
(U87-EGFP), the MPA-coated ZAIS QDs were conjugated
to the siRNA (against the EGFP gene) using polyethyleneimine (PEI) as a cationic polymer via a layer-by-layer approach
(Figure 1B).[12] The efficiency of the PEI coating and siRNA conjugation was monitored using zeta potential (Figure S11). The
decrease in the green fluorescence due to the siRNA-mediated
knockdown of the EGFP gene using our ZAIS QD-siRNA constructs was then monitored (Figure 5) in order to to assess the
transfection efficiency and RNA interference (RNAi) activity. It
was found that the ZAIS QD-siRNA constructs were efficiently
taken up by cells as evident by the intracellular red fluorescence
of the QDs. In addition, the QD localization correlated well
with the decrease of the green fluorescence (∼80% after 3 days
of transfection) resulting from the siRNA-mediated knockdown
of the EGFP mRNA. These results show the ability of our ZAIS
QDs to efficiently translocate siRNA into the cells and achieve
gene knockdown in vitro. With further surface modifications
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using targeting ligands (RGD or TAT peptides and antibodies)
and bifunctional linkers (cleavable linkers or covalent linkers),
the ZAIS QDs could be used for the targeted delivery of siRNA
to cancer/stem cells and for the real-time monitoring of its
delivery in an efficient manner. In addition, the presence of
a library of multicolored QDs obtained via our sonochemical
approach, would allow the for multiplexed imaging of transplanted cell populations in vivo (i.e., tracking different cell
populations with different QDs using different emission wavelengths at the same time).
In summary, we successfully demonstrated the preparation
of ZnxS-AgyIn1-yS2 (ZAIS) QDs using a facile sonochemical synthetic method. The physicochemical and bio-relevant properties
of the resulting QDs can be easily tuned over the entire visible
spectrum by varying the chemical composition of the precursors. We also demonstrated that our ZAIS QDs can exhibit
excellent biocompatibility for the efficient delivery of siRNA and
simultaneous imaging/tracking of the same in cancer cells (and
stem cells) with negligible QD-induced cytotoxicity. While the
ZAIS QDs show great potential for the imaging and delivery of
siRNA in vitro, a more thorough investigation of their long-term
cytotoxicity is needed before they can be used in vivo. Efforts in
this direction are underway. Overall, the ease of the synthesis
of the ZAIS QDs, their excellent cyto-compatibility, and their
versatility as multiplexed imaging agents provides an attractive alternative over conventional QD-based molecular imaging
probes and siRNA delivery vehicles. The above methodology
Adv. Mater. 2012, 24, 4014–4019
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and the IAMDN (Rutgers) for allowing us to use their high-resolution
TEM facility. We are also grateful to Drs. Joung Kyu Park and Jeong Je
Cho for their scientific input and the K.-B.L. group members for their
valuable suggestions for the manuscript. K.-B.L. acknowledges the NIH
Director’s Innovator Award (1DP20D006462-01) and is also grateful to
the N.J. Commission on Spinal Cord grant (09-3085-SCR-E-0). This study
is also funded in part by the US EPA (Grant# 83469302) and the UK
NERC (Grant# NE/H012893).
Synthesis of Precursor Complexes: An aqueous solution of sodium
diethyldithiocarbamate (0.05 M, 5.0 mL) was mixed with an aqueous
solution containing appropriate amounts of AgNO3, In(NO3)3·xH2O
and Zn(NO3)2·6H2O in order to get the required mole ratios (total
concentration of the metal ions was 0.025 M). The solution was allowed
to stir for 5 minutes after which it was filtered using a buchhner funnel,
washed several times with distilled water and MeOH and finally dried
in a convection oven at 60 deg C overnight to obtain the precursor as
a dried powder. Several precursor powders were synthesized by varying
the mole ratios of the metal salts.
Sonochemical Synthesis of Dodecylamine-Capped ZnS-AgInS2 Quantum
Dots: The precursor complex (0.1 g) and dodecylamine (10.0 mL)
were put into a 20 mL vial and sonicated using a tip probe-based high
frequency sonicator (Branson) for 5 minutes in an air-atmosphere.
The resulting suspension was allowed to sit at room temperature for
two minutes after which 5.0 mL of chloroform (5.0 mL) and MeOH
(5.0 mL) were added to it and centrifuged at 4000 rpm. The supernatant
containing the ZAIS QDs was collected and equal amount of MeOH
was added to it in order to isolate the nanoparticles. The obtained
ZAIS QDs were then resuspended in chloroform for absorbance and
photoluminescence measurements.
Surface Modification of ZAIS-QDs with 3-Mercaptopropionic Acid
(MPA): The dodecylamine-capped ZAIS QDs were subjected to a ligand
exchange reaction using 3-mercaptopropionic acid (MPA) according to
a previously reported protocol.[9] Briefly, a 3.0 mL ethanolic solution of
MPA (0.2 M) and KOH (0.3 M) was added dropwise to an equal amount
of the dodecylamine-capped ZAIS QD solution in chloroform. The turbid
solution was stirred for 3h at room temperature followed by centrifugation
at 4000 rpm. The wet precipitate of the MPA-coated ZAIS QDs was
washed with EtOH and redissolved in phosphate-buffered saline (PBS).
The water soluble MPA-coated ZAIS QDs were stable in buffer solution
with no significant change in absorption and photoluminescence for
upto 2 months, when stored at ambient conditions.
Culture of Human U87 Glioblastoma Cells, NIH-3T3 Mouse Fibroblasts
and Human Mesenchymal Stem Cells: The EGFRvIII overexpressed U87
glioblastoma cells (U87) and human mesenchymal stem cells (hMSCs)
were cultured using previously reported methods. For U87-EGFRvIII
cells, DMEM with high glucose, 10% fetal bovine serum (FBS, Gemini
Bioproducts), 1% Streptomycin-penicillin and 1% Glutamax (Invitrogen,
Carlsbad, CA) were used as basic components of growth media including
Hygromycin (30 μg/mL, Invitrogen) as a selection marker. Human
bone marrow-derived MSCs (Lonza, Walkerville) were cultured in the
conditioned media (Lonza, Walkerville) according to manufacturer’s
recommendations. For the NIH-3T3 mouse fibroblasts, DMEM (with
high glucose) supplemented with 10% fetal calf serum (FCS, Gemini
Bioproducts), 1% Streptomycin-penicillin and 1% Glutamax was used as
the growth medium. All cells were maintained at 37 oC in humidified 5%
CO2 atmosphere.
Supporting Information is available from the Wiley Online Library or
from the author.
Acknowledgements
We thank Prof. Gene Hall for help with the XRF measurements,
Dr. Tom Emge, and Vaishali Thakral for their support with the XRD
Adv. Mater. 2012, 24, 4014–4019
Received: March 10, 2012
Revised: April 5, 2012
Published online: June 29, 2012
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could be potentially extended to synthesize libraries of various
types of nanoparticles (magnetic nanoparticles and upconverting near-IR fluorescent nanoparticles), thereby allowing for
rapid screening of the nanomaterials for biomedical applications such as drug delivery and cellular labeling.
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Selective Inhibition of Human Brain Tumor Cells through

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