Biofunctional silicon nanoparticles by means of thiol-ene click chemistry

Article abstract of DOI:10.1002/asia.201100375

The preparation and characterization of butylene-terminated silicon nanoparticles (SiNPs) and their functionalization using thiol-ene chemistry is described, as well as the coupling of DNA strands. Bromide-terminated SiNPs were prepared by means of the oxidation of magnesium silicide and functionalized with butylene chains through treatment with the corresponding Grignard reagent. The successful coupling was confirmed by NMR and FTIR spectroscopy. TEM measurements revealed a silicon-core diameter of (2.4±0.5)nm. The fluorescence emission maximum is at λ_max=525nm when excited at λ_exc=430nm. The conjugation of these alkene-terminated SiNPs by means of thiol-ene chemistry is described for a variety of functional thiols. Efficient coupling was evidenced by NMR and FTIR spectroscopy. Moreover, the characteristic fluorescence properties of the SiNPs remained unaltered, thus demonstrating the value of this approach towards functional oxide-free SiNPs. Activation of the attached carboxylic acid moieties allowed for conjugation of NH₂-terminated oligo-ssDNA (ss=single strand) to the SiNPs. Successful coupling was confirmed by a characteristic new UV absorption band at 260nm, and by the still-present distinctive fluorescence of the SiNPs at 525nm. Gel electrophoresis confirmed coupling of 2 to 3 DNA strands onto the SiNPs, whereas no uncoupled DNA was observed.

Full text of DOI:10.1002/asia.201100375

FULL PAPERS
DOI: 10.1002/asia.201100375
Biofunctional Silicon Nanoparticles by Means of Thiol-Ene Click Chemistry
Loes Ruizendaal,[a] Sidharam P. Pujari,[a] Veronique Gevaerts,[b] Jos M. J. Paulusse,*[a] and
Han Zuilhof*[a]
On the occasion of the 10th anniversary of click chemistry
Abstract: The preparation and charac-
terization of butylene-terminated sili-
con nanoparticles (SiNPs) and their
cence emission maximum is at lmax
=
the value of this approach towards
functional oxide-free SiNPs. Activation
of the attached carboxylic acid moieties
allowed for conjugation of NH2-termi-
nated oligo-ssDNA (ss=single strand)
to the SiNPs. Successful coupling was
confirmed by a characteristic new UV
absorption band at 260 nm, and by the
still-present distinctive fluorescence of
the SiNPs at 525 nm. Gel electrophore-
sis confirmed coupling of 2 to 3 DNA
strands onto the SiNPs, whereas no un-
coupled DNA was observed.
525 nm when excited at lexc =430 nm.
The conjugation of these alkene-termi-
nated SiNPs by means of thiol-ene
chemistry is described for a variety of
functional thiols. Efficient coupling was
evidenced by NMR and FTIR spectros-
copy. Moreover, the characteristic fluo-
rescence properties of the SiNPs re-
mained unaltered, thus demonstrating
functionalization
using
thiol-ene
chemistry is described, as well as the
coupling of DNA strands. Bromide-ter-
minated SiNPs were prepared by
means of the oxidation of magnesium
silicide and functionalized with butyl-
ene chains through treatment with the
corresponding Grignard reagent. The
successful coupling was confirmed by
NMR and FTIR spectroscopy. TEM
measurements revealed a silicon-core
diameter of (2.4Æ0.5) nm. The fluores-
Keywords: bioconjugation
· click
chemistry · fluorescence spectrosco-
py · nanoparticles · silicon
Introduction
the size-dependent bandgap energy, this allows for tuning of
their absorption and fluorescence emission wavelengths.[5]
QDs in biological systems have already found several appli-
cations in bioanalysis[4] and bioimaging.[2,4,6] These QDs are,
however, not tolerant to many common solvents due to the
noncovalent manner in which their surface coating is typi-
cally attached. Moreover, the inherent toxicity of core
atoms like Cd and Pb impedes widespread application in
biological environments.[7] Finally, the total size of typical
core–shell QDs, including the typically essential stabilizing
polymeric coating, is relatively large (20–50 nm). This affects
cell penetration[8] and the residence time within the organ-
ism: only particles smaller than 5 nm in diameter are readily
excreted from the body through urinary excretion; larger
particles accumulate in the body and are disposed of in
other ways.[8]
Silicon nanoparticles (SiNPs) with sizes smaller than the
Bohr exciton radius (ꢀ5 nm) have optical properties com-
parable to conventional QDs.[9] However, extensive studies
have indicated their intrinsic nontoxicity,[10] depending on
the surface charge,[11] thus demonstrating their potential for
imaging in biological systems. The protective organic coating
of the SiNPs may be very thin (ꢀ0.5 nm), thereby maintain-
Bioimaging is essential in gaining a detailed understanding
of cellular processes and disease mechanisms. Organic dyes
have been successfully employed in the labeling of cells,[1]
but are prone to photobleaching. This severely limits the
maximum exposure time,[2] thus rendering multigeneration
staining of cells practically impossible. Quantum dots (QDs)
provide an interesting alternative[3] since they are highly re-
sistant against photobleaching.[2,4] Furthermore, since the
optical properties of QDs are dependent on their size due to
[a] L. Ruizendaal, S. P. Pujari, J. M. J. Paulusse, H. Zuilhof
Laboratory of Organic Chemistry
Wageningen University
Dreijenplein 8, 6703 HB Wageningen (The Netherlands)
E-mail: Jos.Paulusse@wur.nl
Han.Zuilhof@wur.nl
[b] V. Gevaerts
Molecular Science and Technology
Eindhoven University of Technology
Den Dolech 2, 5612 AZ Eindhoven (The Netherlands)
Supporting information for this article is available on the WWW
under http://dx.doi.org/10.1002/asia.201100375.
2776
ꢀ 2011 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
Chem. Asian J. 2011, 6, 2776 – 2786

ing the total diameter at a relatively small size. Several
methods to obtain SiNPs have been developed thus far, such
as electrochemical dispersion of crystalline silicon,[12] ultra-
sonication of silicon wafers,[13] thermal degradation of silanes
in supercritical fluids,[14] and laser-driven pyrolysis of si-
lanes.[15] Furthermore, a number of wet-chemical bottom-up
approaches have been developed, such as the synthesis in
micelles by using organosilanes as silicon source,[16] and
through the reactions with Zintl salts.[17] These wet-chemical
methods typically result in SiNPs that are either hydrogen-
terminated or halogen-terminated, and hence highly prone
to oxidation. Since oxidation of the SiNPs dramatically af-
fects their optical properties,[18] passivation of the SiNPs by
using terminal alkenes[16a] or alkyl-lithium reagents[17a] is es-
rigid, and thus increases the hydrodynamic radius of the par-
ticle accordingly.
The conjugation of biomolecules such as antibodies and
proteins with fluorescent labels is of great importance for
targeted labeling of certain parts of cells or cell-wall recep-
tors. Jaiswal et al. reported a general method for coating
QDs with proteins or antibodies for targeting purposes.[28]
Commercially available QDs were coated with avidin, after
which the biotinylated molecule of interest was coupled.
Nonetheless, it remains a challenge to prepare QDs that do
not remain in clusters but move independently and are effi-
ciently internalized by cells. Due to the relatively large di-
mensions of QDs (20–50 nm), regular uptake by cells would
entail endocytosis. This leads to encapsulated clusters of
QDs, which are only barely released from the capsules into
the cell.[29] However, QDs functionalized with targeting pep-
tides have been successfully applied in the specific labeling
of cell nuclei,[6b,30] whereas proteins located in the cytoplasm
were specifically labeled by making use of His-tag modified
QDs.[31] Specific labeling of a certain cell type was also ac-
complished with antibody-coated QDs, which may then bind
to membrane receptors. In this way, Nie and co-workers
achieved the specific targeting of human prostate tumor
cells in mice by coupling prostate-specific membrane anti-
gen monoclonal antibodies onto QDs.[32] Other examples are
the labeling of tumorous cells with peptide-conjugated QDs,
as well as the monitoring of diffusion dynamics of single re-
ceptors on cell membranes with antibody-coated QDs.[33]
DNA-conjugated QDs are commonly applied in fluores-
cence in situ hybridization (FISH)[23] that allows certain
parts of chromosomes to be labeled to detect chromosomal
defects.[34]
Direct coupling of functional groups or biomolecules onto
SiNPs is not trivial due to the high reactivity of the passiva-
tion agent (alkyl lithium reagents or Grignard reagents for
bromide-terminated SiNPs), as well as the susceptibility of
hydrogen-terminated or halogen-terminated SiNPs towards
nucleophilic attack.[35] Several approaches have been pur-
sued, for example, the use of protective group chemistry[36]
or the use of innocent reactive groups, such as epoxides,[37]
azides,[10] amines,[38] or terminal alkenes.[37] Wang et al. re-
ported on the conjugation of DNA strands onto alkyl-func-
tionalized SiNPs by using a photochemical reaction with an
aryldiazirine to activate the alkyl moiety by means of car-
bene-chemistry.[39] NHS groups attached in this way were
subsequently substituted by amine-terminated single-strand
(ss) DNA strands. Gel electrophoresis confirmed attachment
of the DNA strands. An alternative promising approach for
the functionalization of SiNPs is the use of “click chemistry.”
Click reactions are characterized by their mild reaction con-
ditions, the use of benign solvents, high regio- and chemose-
lectivity, and high yields.[40] Thiol-ene chemistry, which in-
volves a radical-initiated coupling of a thiol to an alkene,
has been known for over a century.[41] Recently, it has at-
tracted great interest as a click reaction, since it does not re-
quire a metal catalyst, proceeds under very mild conditions,
and is insensitive to water and oxygen.[42] The ready availa-
À
sential. The resulting stable Si C bonds form a stable shell
around the Si core and prevent oxidation. However, for ap-
plication of such SiNPs in selective labeling, functional moi-
eties on the particle are desired.
The development of new and accurate biodiagnostic tools
makes the identification of infectious diseases faster and
thereby prevents unnecessary illness and loss of lives. Cur-
rent methods to detect pathogenic bacteria and viruses—
such as tuberculosis and influenza—are laborious and time-
consuming, since typically an amplification step is involved,
which may take several days.[19] The search for faster tests
with even lower detection limits therefore remains impor-
tant. For DNA and RNA sensing in particular, hybridization
of the target sequence with a probe is a commonly em-
ployed approach because of its simplicity and effective-
ness.[20] Recently developed nanomaterials for the detection
of DNA and RNA—which are based, among others, on
magnetic sensing, electronic detection, surface plasmon res-
onance, or fluorescence spectroscopy[21]—are highly promis-
ing, as they combine fast analysis with high specificity and
low detection limits. For these detection systems, the conju-
gation of biomolecules is crucial. Fluorescence spectrosco-
py-based detection methods are increasingly employing
QDs as the source of fluorescence instead of organic dyes.[22]
This is due to their strongly diminished photobleaching[2,4]
and size-tunable emission wavelength.[5] The detection of
biomolecules in solution with QDs may be achieved in sev-
eral ways: by making use of Fçrster resonance energy trans-
fer (FRET), fluorescence quenching, and by changes in the
fluorescence anisotropy.[16a,23] In FRET, the QD acts as a
FRET donor onto which a capture molecule is attached;
upon binding of the target molecule—that is, DNA or pro-
teins—a FRET acceptor is brought close enough to the
donor to display FRET emission.[24] With this method, sensi-
tive detection of DNA target strands has been achieved.[25]
Analogously, binding of target DNA may bring a fluores-
cence quencher into the close proximity of a QD,[26] thereby
resulting in loss of fluorescence. Recently, changes in the
fluorescence anisotropy of QDs were used to detect binding
of a complementary DNA strand to a single-strand DNA-
functionalized QD.[27] The rotation correlation time in-
creased drastically upon hybridization of the complementary
DNA strand, since a double-stranded DNA molecule is very
Chem. Asian J. 2011, 6, 2776 – 2786
ꢀ 2011 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
www.chemasianj.org
2777

J. M. J. Paulusse, H. Zuilhof et al.
FULL PAPERS
bility of thiol-functional biomolecules is an additional ad-
vantage. Several excellent reviews exemplify the many ap-
plications of the versatile thiol-ene reaction[40c,41b,42b,43] Ex-
amples of successful application are the modification of sur-
faces,[44] polymers,[45] and the preparation of dendrimeric
structures[46] and soft polymeric stamps;[47] this reaction was
even demonstrated to proceed in sunlight without extra irra-
diation with UV light.[45a] The versatility and mild conditions
of this reaction prompted us to employ thiol-ene click
chemistry in the modification of alkene-terminated SiNPs.
In this paper we present the development of alkene-termi-
nated SiNPs, their detailed characterization, and their subse-
quent functionalization with a variety of functional groups
using thiol-ene click chemistry (Scheme 1). Furthermore, the
functionalization of carboxylic acid-terminated SiNPs with
amine-functional DNA strands is described, and the detailed
characterization of the resulting bioconjugates is provided.
Results and Discussion
Alkene-Terminated SiNPs
Figure 1. TEM image of alkene-terminated SiNPs (1), with corresponding
size-distribution histogram (inset).
Bromine-terminated SiNPs were synthesized following an
adapted procedure by Kauzlarich and co-workers by means
of oxidation of Mg2Si with bromine.[17a] Capping these bro-
mine-terminated SiNPs with 3-butenylmagnesium bromide
yielded alkene-terminated SiNPs (SiNP-ene, 1). The result-
ing SiNPs were purified by size-exclusion chromatography
(SEC) with ethyl acetate as eluens to yield per reaction
batch SiNPs in 30 mg quantities as an orange waxy material.
The size of the SiNP core was determined by TEM.
Figure 1 shows a typical TEM image of 1; the observed par-
ticle size is (2.4Æ0.5) nm, with a radius-based polydispersity
(PDI) of 1.12. (see the Supporting Information) The alkene-
terminated SiNPs have a slightly smaller radius than ob-
served earlier for butyl-terminated SiNPs.[48] This is likely
due to the use of SEC instead of silica chromatography as
part of the purification procedure. SEC is based on hydrody-
namic radius, and may result in a different size distribution
than that obtained by a predominantly polarity-based sepa-
ration.
whereas the double bond protons are observed at d=4.96
(Figure 2c) and d=5.79 ppm (Figure 2d). The observed
signal broadening is most likely due to the many different
chemical environments of the protons, caused by attachment
to different surface sites of the butylene chains. Moreover,
due to substantial bromination of the reaction solvent, the
attachment of linear and branched octyl chains as well as
octyl bromides is also observed (d=3.5–4.5 ppm). Diffusion-
ordered NMR spectroscopy (DOSY; see the Supporting In-
formation) revealed that the alkene moieties as well as
these brominated species are attached to the SiNPs, since
they have the same relatively low diffusion coefficient. The
low diffusion coefficient (as compared to solvent) indicates
that the signal stems from a relatively large moiety, that is,
an SiNP. The amount of (brominated) octyl chains was de-
1
termined by H NMR spectroscopy to be butene/octyl=2.78
(see the Supporting Information). The 13C NMR spectrum
(Figure 2, bottom) shows signals that correspond to alkene
moieties at d=112 ppm (Figure 2b) and d=136 ppm (Fig-
ure 2c), whereas the alkyl chain appears as multiple signals
SiNP-ene (1) was characterized by NMR spectroscopy.
1
À
Figure 2 (top) shows an H NMR spectrum in which the Si
CH2 protons result in a signal at d=0.87 ppm (Figure 2a),
Scheme 1. Functionalized fluorescent SiNPs obtained by using thiol-ene click chemistry.
2778
www.chemasianj.org
ꢀ 2011 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
Chem. Asian J. 2011, 6, 2776 – 2786

Biofunctional Silicon Nanoparticles by Means of Thiol-Ene Click Chemistry
À
Furthermore, the antisymmetric C H stretch of the alkene
moiety is visible at 3077 cmÀ1. Silicon oxide stretch frequen-
cies are normally observed at approximately 1000–
1100 cmÀ1, and characterized by strong signals due to the
À
high polarity of the Si O bond. The silicon oxide stretch
signal in the IR spectrum of the SiNP-ene (1) is, however,
weak, which indicates that only minor oxidation of the
SiNPs took place during synthesis. Since no increases of the
À
Si O signals were observed for aged samples (several
months), the Si core of such butylene-coated SiNPs (1) is
presumably well protected against further oxidation.
SiNP-ene (1) was further characterized by X-ray photo-
electron spectroscopy (XPS) to obtain information about
the elemental composition. The wide-scan XPS spectrum re-
vealed the presence of silicon (21.1%), carbon (61.7%),
bromine (0.4%), and oxygen (16.9%). The oxygen in the
sample is most likely largely due to environmental entrap-
ment in the deposited SiNPs within the XPS and not as sili-
con oxide, since the silicon narrow-scan spectrum displays
only a single signal at 101.8 eV, with a full width at half-
maximum (FWHM) of 1.3 eV (Figure 4, right), which is
near-identical to what is observed under similar instrumental
Figure 2. 1H NMR (top) and 13C NMR (bottom) spectra of SiNP-ene (1).
in the d=21–31 ppm region
(Figure 2a). The assignment of
these signals was confirmed by
2D COSY and 2D heteronu-
clear single quantum coher-
ence (HSQC) spectra (see the
Supporting Information). In
the COSY spectrum, protons b
and c were confirmed to be
positioned on neighboring
carbon atoms, furthermore, the
1H–13C HSQC confirms the as-
signment of carbon atoms a, b,
Figure 4. XPS spectra of SiNP-ene (1). Left: C1s narrow scan; right: Si2p narrow scan.
and c in the 13C NMR spec-
trum, due to coupling with the
corresponding signals in the 1H NMR spectrum. A broad
signal at d=130 ppm (Figure 2d) in the 13C NMR spectrum
indicates the presence of internal alkenes, most likely result-
ing from elimination reactions that take place on the afore-
mentioned brominated octyl chains due to the Grignard re-
agent acting as a base.
conditions for the base peak of Si at 99.4 eV with a FWHM
of 0.5 eV in a siliconACTHNUTRGNEUNG(111) wafer. The shift in binding energy
from 99.4 to 101.8 eV is thus at least partially caused by
charging effects, which would indeed be expected for a
small object that is well surrounded by an electrically insu-
lating organic shell. Moreover, XPS analysis of samples of
intentionally oxidized SiNPs resulted in two signals in the
silicon region, the first at 101.8 eV corresponding to silicon,
and a second signal at 104.9 eV for silicon oxide (see the
Supporting Information). The peak shift is in line with the
observation that the binding energy shifts to higher levels
with decreasing nanoparticle size relative to bulk silicon.[49]
The narrow scan of the C1s region (Figure 4, left) reveals
two types of carbon: carbon-bound carbon at 285.0 eV, and
a minor fraction (14%) of carbon atoms bound to electro-
negative elements such as Br or O (broad shoulder around
287 eV). The precise identity of this peak can, however, not
be deduced from XPS analysis alone.
IR spectroscopy (Figure 3) shows characteristic signals
that correspond to the CH2 groups at 2871 and 2959 cmÀ1.
Figure 3. IR spectrum of SiNP-ene (1).
Chem. Asian J. 2011, 6, 2776 – 2786
ꢀ 2011 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
www.chemasianj.org
2779

J. M. J. Paulusse, H. Zuilhof et al.
FULL PAPERS
Figure 5. UV/Vis absorption (left) and fluorescence emission map (right) of SiNP-ene (1). Irel =relative intensity.
The UV/Vis absorption spectrum of 1 (Figure 5, left) was
recorded. However, the spectrum does not display a maxi-
mum, it only shows a gradually higher absorption for de-
creasing wavelengths, as was observed before for this type
of SiNPs.[17a,48] From these samples, the absorption coeffi-
cient was determined to be 0.14 (mgmLÀ1)À1 cmÀ1 at 300 nm
and 0.035 (mgmLÀ1)À1 cmÀ1 at 350 nm. The extinction coeffi-
cient was determined in (mgmLÀ1)À1 cmÀ1, since the exact
molecular mass of the SiNP is unknown. Based on a molecu-
lar weight of 10000 gmolÀ1—estimated from TEM measure-
ments in combination with a dense packing of butylene
chains—approximate molar extinction coefficients of 3.0ꢁ
102 mÀ1 cmÀ1 at 350 nm and 1.4ꢁ103 mÀ1 cmÀ1 at 300 nm were
calculated. These values are lower than observed for the
1.5 nm SiNPs prepared by other wet-chemical methods
(9.4ꢁ103 mÀ1 cmÀ1; 1.1 (mgmLÀ1)À1 cmÀ1)[16a] or for those re-
ported for 1.5 nm QDs of II–VI semiconductors, such as
CdSe and CdS (7ꢁ104 mÀ1 cmÀ1).[50] However, bulk silicon
has an indirect bandgap, which makes optical transitions
without the assistance of a phonon inefficient. The optical
transition in an SiNP is very efficient in comparison to the
transitions in bulk silicon. In the nanometer size regime, the
distinction between indirect and direct bandgaps is blur-
red.[51]
The fluorescence emission spectrum is strongly dependent
on the excitation wavelength. Figure 5 (right) depicts fluo-
rescence emission spectra of 1 at several excitation wave-
lengths that range from 350 to 450 nm. The spectra are cor-
rected for the UV/Vis absorption at each excitation wave-
length. The relatively highest fluorescence emission is ob-
served at 525 nm at an excitation wavelength of 430 nm.
This is distinctly redshifted as compared to butyl-terminated
SiNPs described previously,[17a] which showed an emission
maximum at 390 nm (obtained for irradiation at 340 nm).
For SiNPs with a gradual increase in UV/Vis absorption at
shorter wavelengths, correction for UV/Vis absorption re-
sults in a redshifted fluorescence emission maximum.
The QYs are (1.5Æ0.2)% and (1.8Æ0.3)% for lexc =350 nm
and 366 nm, respectively. The QY increases further with in-
creasing excitation wavelength; the highest QY is observed
at an excitation wavelength of 496 nm, and is (7.1Æ1.2)%.
This is about 2% higher than observed for highly similar
butyl-terminated SiNPs,[48] but still considerably smaller
than the highest QY reported for SiNPs (60%).[53] It has
been suggested that a high oxidation grade of the SiNPs re-
sults in lower quantum yields,[54] however, such high degrees
of oxidation are absent in this case as follows from IR and
XPS data. The larger size (2.5 nm in this case) is therefore a
more likely cause.
Time-resolved fluorescence spectroscopy was performed,
and the amplitude-weighed fluorescence lifetime (see the
Supporting Information) was determined to be 3.40 ns,
which is similar to the lifetimes observed for butyl-terminat-
ed SiNPs.[48] These results indicate that the optical properties
of the SiNPs are not significantly influenced by the change
in coating from butyl to butylene.
Thiol-Ene Functionalization of Alkene-Terminated SiNPs
With the alkene moieties present on the surface of the
SiNPs, functionalization by means of thiol-ene chemistry is
possible. Functional groups may be attached, which make
the SiNPs suitable for applications that require specific
labels. The SiNPs were mixed with an excess amount of the
respective thiol and 0.2 equiv (with respect to the thiol) of
2,2-dimethoxy-2-phenylacetophenone (DMPA) as photoini-
tiator or 4,4’-azobis(4-cyanovaleric acid) (ACVA) as thermal
initiator. The mixture was exposed to UV light (DMPA,
365 nm) or heated to 808C (ACVA) while stirring under am-
1
bient atmosphere. After 1 h, H NMR spectroscopy revealed
complete conversion of the alkene moieties, and the func-
tionalized SiNPs were purified by means of SEC. A number
of thiols with different functionalities was selected: thiolace-
tic acid (SiNP-TAA, 2) and 2-mercaptoethanol (SiNP-OH,
3)—which provide for a functional group that may be used
in further conjugation reactions—as well as thiolated tri-
ethylene glycol monomethyl ether (SiNP-EO3, 4), to render
the SiNPs water-soluble and biocompatible. Furthermore,
The fluorescence quantum yield (QY) was determined
using a comparative method[23,52] at lexc =350, 366, and
496 nm. These dyes have a fluorescence emission that
broadly overlaps the fluorescence emission of the SiNPs.
2780
www.chemasianj.org
ꢀ 2011 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
Chem. Asian J. 2011, 6, 2776 – 2786

Biofunctional Silicon Nanoparticles by Means of Thiol-Ene Click Chemistry
carboxylic acid terminated thiols with three different spacer
lengths (no spacer (6), an EO4 spacer (7), and a PEG3000
spacer (8)) were coupled to the SiNPs to obtain functional
groups available for bioconjugation on the surface of the
SiNPs (Scheme 1).
at d=2.62 ppm (Figure 7d) (CH2-CH2-S) and d=3.19 ppm
(Figure 7e) (S-CH2-COOEt). The signals of the ethyl ester
moiety were observed at d=4.17 ppm (Figure 7 f) and d=
1.26 ppm (Figure 7g), which correspond to the CH2 and
CH3, respectively. After hydrolysis with KOtBu, these sig-
nals are no longer present. The NMR spectra of the thiol-
ene-modified SiNPs were assigned with 2D COSY NMR
spectroscopy, and in all cases confirm quantitative coupling
of the indicated thiols to the SiNPs, since no terminal al-
kenes are observed after modification, whereas the intensi-
The functionalized SiNPs 2, 3, 4, 6, 7, were purified by
means of SEC, whereas pegylated SiNPs (8) were purified
with a Amicon Ultra 3K nominal weight cutoff (NWCO)
centrifugal units. To obtain SiNP-TGA (6), direct coupling
of thioglycolic acid is possible and results in full conversion
by means of the thiol-ene reaction. However, it appeared
that the SiNP-TGA (6) obtained in this manner could not
be purified. Therefore, the ethyl ester-protected SiNP-
TGAEE (5) was hydrolyzed with potassium tert-butoxide
(KOtBu) to obtain thioglycolic acid-modified SiNPs (SiNP-
TGA, 6). The sample was neutralized and purified by evap-
oration of the solvents.
1
ties of the new signals fit accordingly. H NMR spectra of
other modified SiNPs can be found in the Supporting Infor-
mation.
Infrared spectroscopy was performed on the SiNPs to
confirm the presence of the characteristic groups and to de-
termine whether the functionalized SiNPs retained their low
oxidation grade (Figure 8). The C=CH2 antisymmetric
In the 1H NMR spectrum of purified SiNP-TAA (2)
(Figure 6), no terminal alkenes are observed. Furthermore,
Figure 8. FTIR spectra of SiNPs 2, 4, and 7. FTIR spectra of SiNP 3, 5, 6,
and 8 can be found in the Supporting Information.
Figure 6. 1H NMR spectrum of SiNP-TAA (2).
stretch signal, which appeared at 3077 cmÀ1 in the case of
SiNP-ene, was absent in the spectra of the functionalized
SiNPs. All functionalized SiNPs display the typical symmet-
ric and antisymmetric stretch vibrations for CH2 bonds. Sig-
nals for the attached functional groups are readily observed.
SiNP-TAA (2), for example, displays a distinct carbonyl
stretch signal at 1690 cmÀ1, which confirms the presence of a
thioester. SiNP-OH (3) displays a broad OH stretch vibra-
tion at 3354 cmÀ1, whereas SiNP-EO3 (4) displays character-
istic ether bonds at 1108 cmÀ1. The spectra of SiNP-TGAEE
(5), SiNP-TGA (6), and SiNP-EO4-COOH (7) display dis-
tinct carbonyl stretch signals at 1732 cmÀ1. The signal is,
however, hardly observed in the case of SiNP-PEG3000-
COOH (8), most likely due to the large amount of ether
groups (approximately 170 per chain, at 1103 cmÀ1) relative
to other characteristic groups present in the sample. The
presence of ether groups also results in broadening of the
signals for the CH2 symmetric and antisymmetric stretch
(they appear as a single peak at 2880 cmÀ1). In the IR spec-
trum of SiNP-EO4-COOH (7) the characteristic ether bonds
a new signal at d=2.86 ppm (Figure 6d) corresponds to the
newly formed thioester; signals that correspond to the CH2
next to the thioester appear at d=1.57 ppm (Figure 6c),
whereas the signal of the methyl group next to the carbonyl
appears at d=2.32 ppm (Figure 6e).
The 1H NMR spectrum of SiNP-TGAEE (5) (Figure 7)
displays characteristic signals of the newly formed thioether
are observed at 1108 cmÀ1. Importantly, the signal that cor-
À1
À
responds to Si O (1000–1100 cm ) continues to be weak in
Figure 7. 1H NMR spectra of SiNP-TGAEE (5).
all spectra, despite the high polarity of the Si O bond. This
À
Chem. Asian J. 2011, 6, 2776 – 2786
ꢀ 2011 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
www.chemasianj.org
2781

J. M. J. Paulusse, H. Zuilhof et al.
FULL PAPERS
indicates that even upon per-
forming
the
radical-based
thiol-ene coupling the low
degree of oxidation of the Si
core is maintained, thus pro-
viding further proof of the pas-
sivating nature of the organic
coating.
The SiNPs were further
characterized by using XPS to
obtain an elemental composi-
tion of the SiNPs, as well as
characterizing the different
functional groups. A thin film
of SiNP-TAA (2) was cast onto
a gold surface and analyzed.
Figure 10. UV/Vis absorption spectra (left) and fluorescence emission (right) of SiNPs 1, 2, and 4. Spectra of
SiNPs 3, 5, 6, 7, and 8 can be found in the Supporting Information.
The XPS narrow scan spectrum of the C1s region of 2 was fit
with four components, of which the main peak at 285.0 eV is
assigned to carbon-bound carbon atoms (Figure 9, left). The
rescence emission spectra were recorded. In Figure 10
(right), the UV/Vis absorption-normalized fluorescence
emission spectra of SiNPs excited at 430 nm are shown, at
which excitation wavelengths
is obtained. The fluorescence
emission maxima do not shift
significantly for the modified
SiNPs and remain at 525 nm.
The signal intensity, however,
does change to some degree.
A slight shift in maximum is
observed when comparing
SiNP-PEG3000-COOH
(8)
and the other functionalized
SiNPs. This is likely as a result
of the size-based purification
process of 8, which may alter
the size distribution of the
SiNPs. Since the fluorescence
Figure 9. XPS spectra of SiNP-TAA (2). Left: C1s narrow scan; right: S2p narrow scan.
intensity of this signal is relatively high (78% of the total
carbon content), due to the earlier mentioned side reactions
that occur, thereby leading to substantial attachment of
alkyl chains (see the Supporting Information). The signals
emission of SiNPs is size-dependent, changes in the size dis-
tribution consequently result in changes in the fluorescence
emission maxima.[15b] This shift is not expected to result
from the thiol-ene conjugation, since fluorescence originates
from the Si core, with only a small electronic coupling to the
covering organic layer, which is unlikely to extend to the
distance at which the conjugation reaction takes place.[55]
Besides steady-state fluorescence spectroscopy, time-re-
solved fluorescence emission was studied as well. The Sup-
porting Information summarizes the amplitude-weighted
averages of the fluorescence lifetimes of the functional
SiNPs excited at 372 nm. The results are all in the same
time range (ꢀ4 ns), thereby further confirming that thiol-
ene modification does not significantly affect the SiNPs.
À
for the C=O (288.0 eV) and C S (286.4 eV) carbon atoms
integrate to a 1:1 ratio. The signal at 289.3 eV (2% of the
C1s total area) may arise from slight oxidation of the C=C
bond. The XPS narrow scan of the Si2p region (Figure 9,
center) reveals only a single signal at 101.8 eV, similar to
that observed for SiNP-ene (1). The sulfur region shows a
clear signal of a single type of sulfur, which was fit with a
single spin–orbital doublet (Figure 9, right), in agreement
with the thioether that results from the attachment step.
XPS spectra of other modified SiNPs can be found in the
Supporting Information.
To determine whether the SiNPs retain their optical prop-
erties after thiol-ene functionalization, the UV/Vis absorp-
tion spectra were recorded (Figure 10, left). As evidenced
by the spectra, with normalized absorption at 320 nm, the
absorption is not significantly affected by the modification
of the SiNPs. This confirms that the Si core remains unal-
tered under thiol-ene coupling conditions. In addition, fluo-
DNA Conjugation
To obtain biofunctional SiNPs with possible applications in
biosensors, single-strand deoxyribonucleic acid (ssDNA)
with a length of 100 bases with a 3’-amino modification was
coupled to the SiNPs by using 1-ethyl-3-(3-dimethylamino-
propyl)carbodiimide and N-hydroxysuccinimide (EDC/
2782
www.chemasianj.org
ꢀ 2011 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
Chem. Asian J. 2011, 6, 2776 – 2786

Biofunctional Silicon Nanoparticles by Means of Thiol-Ene Click Chemistry
Scheme 2. ssDNA attachment onto SiNPs by means of EDC/NHS chemistry (example shown: SiNP-TGA (6)).
NHS) chemistry (Scheme 2). Samples were purified with
50 k NWCO Amicon centrifugal filter units. Control experi-
ments revealed that both uncoupled ssDNA and uncoupled
SiNPs readily pass through 50 k NWCO filters, as evidenced
by UV/Vis absorption spectroscopy.
After coupling of the ssDNA and subsequent purification
to obtain ssDNA-functionalized SiNPs, the amount of DNA
in the NP sample was determined by UV/Vis absorption
measurements to determine its concentration. The comple-
mentary 3’-Atto 488 dye-modified ssDNA strand was added
(1.5 equiv) and incubated at 808C for 1 min, after which the
sample was allowed to cool to room temperature followed
by cooling on ice. Excess amounts of complementary
ssDNA were removed by filtration over a 50 k NWCO
Amicon centrifugal filter unit. This resulted in double-
strand (ds) DNA-modified SiNPs SiNP-TGA-dsDNA (9),
SiNP-EO4-dsDNA (10), and SiNP-PEG-dsDNA (11)
(Figure 11).
Figure 12. Agarose gel with dsDNA-modified SiNPs. Lane 1: 1 kb+
ladder; lane 2: SiNP (9); lane 3: SiNP (10); lane 4: SiNP (11); lane 5: un-
coupled 100 bp dsDNA.
purification of the conjugated SiNPs. The heights of the
bands in lanes 3 and 4 correspond to approximately 300 bp
(Figure 12, arrow II). The SiNPs to which the DNA are cou-
pled are of a specific size, which may influence the height of
the band slightly. As such, the bands are proposed to indi-
cate the presence of two to three DNA strands per NP for
SiNP-EO4-dsDNA (10) and SiNP-PEG-dsDNA (11). A
slightly less distinct band at the same height is observed in
lane 2, which indicates that in the SiNP-TGA-dsDNA (9)
sample, up to three strands are also attached to the SiNP.
The clearer band in lane 2 just above the 100 bp level
(Figure 12, arrow III) most likely corresponds to SiNP-
TGA-dsDNA (9) with only a single dsDNA strand attached.
Tailing of the bands may be due to the range of NP sizes
present in the samples (due to a different number of poly-
meric chains attached), or different numbers or orientations
of DNA strands on the SiNPs, thereby resulting in different-
ly sized DNA-functional particles.
UV/Vis absorption and fluorescence emission data of the
DNA-modified samples with different spacer lengths un-
fortunately do not display characteristics that allow for dif-
ferentiation. The UV/Vis absorption spectrum of the pegy-
lated SiNPs conjugated to ssDNA is shown in Figure 13
(left). However, the main absorption observed originates
from the DNA. This points to attachment of the DNA
strands to the SiNP, since otherwise these would have been
removed in the purification process. The absorption of the
SiNP core is still observed when ssDNA is conjugated to the
particle, albeit only as a minor contribution, since the ex-
Figure 11. Schematic representation of SiNPs conjugated to a dsDNA
strand; complementary strand contains a 3’-Atto 488 dye. (Dimensions
scaled, spacers in blue.)
To confirm coupling of the DNA strands to the SiNPs, gel
electrophoresis on an agarose gel was performed. Figure 12
shows the agarose gel after a run time of 1 h. Lane 1 con-
tains the reference ladder; the numbers correspond to the
number of base pairs (bp) in the respective band. Lane 2
contains SiNP-TGA-dsDNA (9); lane 3 SiNP-EO4-dsDNA
(10); and lane 4 contains SiNP-PEG-dsDNA (11). Lane 5 is
a reference lane that contains uncoupled dsDNA (100 bp)
with the same sequence as the strands coupled to the NPs.
The band in lane 5 runs slightly lower than the 100 bp
ladder reference (Figure 12, arrow I). No DNA of this
length is observed in lanes 3 and 4, thus indicating successful
Chem. Asian J. 2011, 6, 2776 – 2786
ꢀ 2011 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
www.chemasianj.org
2783

J. M. J. Paulusse, H. Zuilhof et al.
FULL PAPERS
such as those terminated with
thioacetic acid, mercaptoetha-
nol, ethylene glycol, and car-
boxylic acid. The functionaliza-
tion retains the low degree of
oxidation obtained for such
NPs and does not change the
optical properties of the func-
tionalized SiNPs, which is in
line with the Si core being the
origin of the optical properties
of the SiNPs. Coupling of
amine-terminated DNA to the
Figure 13. Left: normalized UV/Vis absorption of SiNP-PEG-ssDNA and dsDNA (11). Right: normalized
fluorescence emission of SiNP-PEG-ssDNA (lexc =430 nm) and SiNP-PEG-dsDNA (11) (lexc =501 nm).
carboxylic
acid-terminated
SiNPs was confirmed by UV/
Vis absorption and fluores-
tinction coefficient of ssDNA (9.8ꢁ105 mÀ1 cmÀ1, for this par-
ticular sequence) is approximately 1000-fold higher than
that of the SiNPs at 350 nm (ꢀ3ꢁ102 mÀ1 cmÀ1). Further-
more, the fluorescence emission of the SiNPs with ssDNA is
still observed, although again as a relatively small signal
(Figure 13, right). Fluorescence emission contributions from
the SiNP core as well as the UV absorption of the ssDNA
moiety, indicate that both the SiNPs and ssDNA are present
in the sample.
The UV absorption spectrum of the SiNPs that contain
dsDNA (9, 10, 11) is shown in Figure 13 (left). The spectrum
is dominated by the absorption of dsDNA and additionally
the Atto 488 dye. This is similar to the ssDNA SiNP samples
due to the high extinction coefficient for dsDNA (1.6ꢁ
106 mÀ1 cmÀ1, for this particular sequence). The fluorescence
emission of the Atto 488 dye fully dominates the fluores-
cence spectra of the samples that contain dsDNA-conjugat-
ed SiNPs (Figure 13, right).
The fluorescence contribution of the Atto dye indicates
that the complementary strand is indeed hybridized to the
DNA strand present on the SiNP. The dominance of the
Atto 488 dye in the fluorescence spectra with dsDNA is due
to the rather low quantum yield (QY) of the SiNPs (7%),
whereas the QY of the Atto 488 dye is 80%. Figure 13
(right) shows the UV/Vis absorption and fluorescence emis-
sion of SiNP-dsDNA. The spectra are dominated by the ab-
sorption and emission of the coupled dsDNA strands and
the Atto 488 dye. This indicates that the fluorescent dye is
indeed coupled to the SiNPs through the complementary
DNA strand, since all uncoupled DNA strands are removed
by purification as observed by gel electrophoresis.
cence spectroscopy, which indicated both the presence of
DNA and SiNPs in the purified samples. Addition of a fluo-
rescently labeled complementary DNA strand yielded SiNPs
with two to three covalently coupled dsDNA strands, as
confirmed by UV/Vis absorption and fluorescence spectros-
copy and by gel electrophoresis. These results demonstrate
the successful bioconjugation of DNA onto SiNPs. The em-
ployed coupling technique is very mild, and the presented
concept is therefore readily extended to proteins or other
biomolecules, thus bringing the application of biofunctional
SiNPs in sensing devices within reach.
Experimental Section
Experimental details can be found in the Supporting Information. In
brief, the SiNPs were synthesized under an argon atmosphere by the oxi-
dation of Mg2Si by Br2 while heating to reflux in octane for 72 h. Subse-
quently, freshly synthesized 4-bromo-1-butene in THF was added to
obtain alkene-terminated SiNPs. The SiNPs were purified by extraction
and size-exclusion chromatography. For the thiol-ene reaction, in general,
an excess amount of the thiol and the radical initiator (0.2 equiv) with re-
spect to the thiol were added to a solution of SiNP-ene (1; 10 mg)
(approx. 0.1 mmol alkene groups as determined by 1H NMR spectrosco-
py with an internal standard) in chlorobenzene or DMF. The reaction
was stirred at room temperature while being exposed to UV light
(365 nm) (DMPA) or stirred at 808C to activate the thermal initiator
(ACVA), under ambient atmosphere. The resulting SiNPs were purified
by size-exclusion chromatography. Coupling of DNA strands was per-
formed under sterile conditions by adding an excess amount of EDC/HCl
to SiNPs terminated with carboxylic acid, followed by the addition of
NH2-terminated ssDNA. Samples were purified with Amicon centrifugal
filters.
Acknowledgements
Conclusion
The authors thank Anke Trilling (Wageningen University) for experimen-
tal assistance and stimulating discussions. This work was funded by the
Dutch Ministry of Economic Affairs within the MicroNed innovation
program (WP 2-C-II).
The synthesis of silicon nanoparticles (SiNPs) by means of
the oxidation of Mg2Si and subsequent capping with 3-bute-
nylbromide resulted in alkene-functionalized SiNPs with a
silicon core size of 2.4 (Æ0.5) nm. Thiol-ene chemistry per-
formed on these SiNPs proved to be a facile, efficient, and
versatile approach to obtain a variety of functional SiNPs,
[1] J. B. Pawley, Handbook of Biological Confocal Microscopy, Spring-
er, 2006.
2784
www.chemasianj.org
ꢀ 2011 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
Chem. Asian J. 2011, 6, 2776 – 2786

Biofunctional Silicon Nanoparticles by Means of Thiol-Ene Click Chemistry
[2] X. Wu, H. Liu, J. Liu, K. N. Haley, J. A. Treadway, J. P. Larson, N.
Ge, F. Peale, M. P. Bruchez, Nat. Biotechnol. 2003, 21, 41–46.
[3] V. Biju, T. Itoh, M. Ishikawa, Chem. Soc. Rev. 2010, 39, 3031–3056.
[4] M. Bruchez, Jr., M. Moronne, P. Gin, S. Weiss, A. P. Alivisatos, Sci-
ence 1998, 281, 2013–2016.
[24] A. M. Smith, H. Duan, A. M. Mohs, S. Nie, Adv. Drug Delivery Rev.
2008, 60, 1226–1240.
[25] a) R. Gill, I. Willner, I. Shweky, U. Banin, J. Phys. Chem. B 2005,
109, 23715–23719; b) C.-Y. Zhang, H.-C. Yeh, M. T. Kuroki, T.-H.
Wang, Nat. Mater. 2005, 4, 826–831; c) H. Zhang, P. G. Stockley, D.
Zhou, Faraday Discuss. 2011, 149, 319–332; d) X. Wang, X. Lou, Y.
Wang, Q. Guo, Z. Fang, X. Zhong, H. Mao, Q. Jin, L. Wu, H. Zhao,
J. Zhao, Biosens. Bioelectron. 2010, 25, 1934–1940; e) G. Jiang, A. S.
Susha, A. A. Lutich, F. D. Stefani, J. Feldmann, A. L. Rogach, ACS
Nano 2009, 3, 4127–4131.
[26] a) J. Lee, Y. Choi, J. Kim, E. Park, R. Song, ChemPhysChem 2009,
10, 806–811; b) X. Li, J. Qian, L. Jiang, S. He, Appl. Phys. Lett.
2009, 94; c) L. Dyadyusha, H. Yin, S. Jaiswal, T. Brown, J. J. Baum-
berg, F. P. Booy, T. Melvin, Chem. Commun. 2005, 3201–3203.
[27] G. Giraud, H. Schulze, T. T. Bachmann, C. J. Campbell, A. R.
Mount, P. Ghazal, M. R. Khondoker, S. W. J. Ember, I. Ciani, C.
Tlili, A. J. Walton, J. G. Terry, J. Crain, Chem. Phys. Lett. 2010, 484,
309–314.
[28] J. K. Jaiswal, E. R. Goldman, H. Mattoussi, S. M. Simon, Nat. Meth-
ods 2004, 1, 73–78.
[29] J. B. Delehanty, H. Mattoussi, I. L. Medintz, Anal. Bioanal. Chem.
2009, 393, 1091–1105.
[30] F. Chen, D. Gerion, Nano Lett. 2004, 4, 1827–1832.
[31] K. Boeneman, J. B. Delehanty, K. Susumu, M. H. Stewart, I. L. Med-
intz, J. Am. Chem. Soc. 2010, 132, 5975–5977.
[32] X. Gao, Y. Cui, R. M. Levenson, L. W. K. Chung, S. Nie, Nat. Bio-
technol. 2004, 22, 969–976.
[5] a) A. P. Alivisatos, Science 1996, 271, 933–937; b) L. Brus, J. Phys.
Chem. 1986, 90, 2555–2560.
[6] a) K.-T. Yong, H. Ding, I. Roy, W.-C. Law, E. J. Bergey, A. Maitra,
P. N. Prasad, ACS Nano 2009, 3, 502–510; b) A. M. Derfus, W. C. W.
Chan, S. N. Bhatia, Adv. Mater. 2004, 16, 961–966; c) D. S. Lidke, P.
Nagy, R. Heintzmann, D. J. Arndt-Jovin, J. N. Post, H. E. Grecco,
E. A. Jares-Erijman, T. M. Jovin, Nat. Biotechnol. 2004, 22, 198–
203; d) P. Mitchell, Nat. Biotechnol. 2001, 19, 1013–1017;
e) W. C. W. Chan, S. M. Nie, Science 1998, 281, 2016–2018.
[7] a) A. M. Derfus, W. C. W. Chan, S. N. Bhatia, Nano Lett. 2004, 4,
11–18; b) N. Lewinski, V. Colvin, R. Drezek, Small 2008, 4, 26–49.
[8] H. Soo Choi, W. Liu, P. Misra, E. Tanaka, J. P. Zimmer, B. Itty Ipe,
M. G. Bawendi, J. V. Frangioni, Nat. Biotechnol. 2007, 25, 1165–
1170.
[9] a) N. Shirahata, Phys. Chem. Chem. Phys. 2011, in press; b) Z.
Kang, Y. Liu, S.-T. Lee, Nanoscale 2011, 3, 777–791.
[10] L. Ruizendaal, S. Bhattacharjee, K. Pournazari, M. Rosso-Vasic,
L. H. J. De Haan, G. M. Alink, A. T. M. Marcelis, H. Zuilhof, Nano-
toxicology 2009, 3, 339–347.
[11] S. Bhattacharjee, L. H. J. de Haan, N. M. Evers, X. Jiang, A. T. M.
Marcelis, H. Zuilhof, I. M. C. M. Rietjens, G. M. Alink, Part. Fibre
Toxicol. 2010, 7, 25.
[12] E. V. Rogozhina, D. A. Eckhoff, E. Gratton, P. V. Braun, J. Mater.
Chem. 2006, 16, 1421–1430.
[33] a) H. M. E. Azzazy, M. M. H. Mansour, S. C. Kazmierczak, Clin.
Biochem. 2007, 40, 917–927; b) A. M. Smith, X. Gao, S. Nie, Photo-
chem. Photobiol. 2004, 80, 377–385.
[34] P. Chan, T. Yuen, F. Ruf, J. Gonzalez-Maeso, S. C. Sealfon, Nucl.
Acids Res. 2005, 33, e161.
[35] C. Chatgilialoglu, D. Griller, M. Lesage, J. Org. Chem. 1989, 54,
2492–2494.
[36] R. S. Tanke, S. M. Kauzlarich, T. E. Patten, K. A. Pettigrew, D. L.
Murphy, M. E. Thompson, H. W. H. Lee, Chem. Mater. 2003, 15,
1682–1689.
[37] A. Shiohara, S. Hanada, S. Prabakar, K. Fujioka, T. H. Lim, K. Ya-
mamoto, P. T. Northcote, R. D. Tilley, J. Am. Chem. Soc. 2009, 132,
248–253.
[38] a) M. Rosso-Vasic, L. De Cola, H. Zuilhof, J. Phys. Chem. C 2009,
113, 2235–2240; b) M. Rosso-Vasic, E. Spruijt, Z. Popovic, K. Over-
gaag, B. van Lagen, B. Grandidier, D. Vanmaekelbergh, D. Domi-
nguez-Gutierrez, L. De Cola, H. Zuilhof, J. Mater. Chem. 2009, 19,
5926–5933.
[39] L. Wang, V. Reipa, J. Blasic, Bioconjugate Chem. 2004, 15, 409–412.
[40] a) H. C. Kolb, M. G. Finn, K. B. Sharpless, Angew. Chem. 2001, 113,
2056–2075; Angew. Chem. Int. Ed. 2001, 40, 2004–2021; b) R. K.
Iha, K. L. Wooley, A. M. Nystrçm, D. J. Burke, M. J. Kade, C. J.
Hawker, Chem. Rev. 2009, 109, 5620–5686; c) C. Remzi Becer, R.
Hoogenboom, U. S. Schubert, Angew. Chem. 2009, 121, 4998–5006;
Angew. Chem. Int. Ed. 2009, 48, 4900–4908.
[41] a) T. Posner, Ber. Dtsch. Chem. Ges. 1905, 38, 646–657; b) C. E.
Hoyle, T. Y. Lee, T. Roper, J. Polym. Sci. Part A 2004, 42, 5301–
5338.
[13] a) Z. Yamani, S. Ashhab, A. Nayfeh, W. H. Thompson, M. Nayfeh,
J. Appl. Phys. 1998, 83, 3929–3931; b) J. L. Heinrich, C. L. Curtis,
G. M. Credo, K. L. Kavanagh, M. J. Sailor, Science 1992, 255, 66–68.
[14] a) J. D. Holmes, K. J. Ziegler, R. C. Doty, L. E. Pell, K. P. Johnston,
B. A. Korgel, J. Am. Chem. Soc. 2001, 123, 3743–3748; b) D. S. Eng-
lish, L. E. Pell, Z. Yu, P. F. Barbara, B. A. Korgel, Nano Lett. 2002,
2, 681–685.
[15] a) X. Li, Y. He, S. S. Talukdar, M. T. Swihart, Langmuir 2003, 19,
8490–8496; b) F. Huisken, G. Ledoux, O. Guillois, C. Reynaud, Adv.
Mater. 2002, 14, 1861–1865.
[16] a) M. Rosso-Vasic, E. Spruijt, B. van Lagen, L. De Cola, H. Zuilhof,
Small 2008, 4, 1835–1841; b) J. H. Warner, A. Hoshino, K. Yamamo-
to, R. D. Tilley, Angew. Chem. 2005, 117, 4626–4630; Angew. Chem.
Int. Ed. 2005, 44, 4550–4554; c) J. P. Wilcoxon, G. A. Samara, Appl.
Phys. Lett. 1999, 74, 3164–3166; d) J. P. Wilcoxon, G. A. Samara,
P. N. Provencio, Phys. Rev. B 1999, 60, 2704–2714; e) R. D. Tilley,
J. H. Warner, K. Yamamoto, I. Matsui, H. Fujimori, Chem.
Commun. 2005, 1833–1835; f) J. Zou, P. Sanelle, K. A. Pettigrew,
S. M. Kauzlarich, J. Cluster Sci. 2006, 17, 565–578.
[17] a) K. A. Pettigrew, Q. Liu, P. P. Power, S. M. Kauzlarich, Chem.
Mater. 2003, 15, 4005–4011; b) Q. Liu, S. M. Kauzlarich, Mater. Sci.
Eng. B 2002, 96, 72–75; c) D. Neiner, H. W. Chiu, S. M. Kauzlarich,
J. Am. Chem. Soc. 2006, 128, 11016–11017; d) D. Mayeri, B. L. Phil-
lips, M. P. Augustine, S. M. Kauzlarich, Chem. Mater. 2001, 13, 765–
770; e) X. M. Zhang, D. Neiner, S. Z. Wang, A. Y. Louie, S. M. Kau-
zlarich, Nanotechnology 2007, 18, 095601.
[18] G. Ledoux, O. Guillois, D. Porterat, C. Reynaud, F. Huisken, B.
Kohn, V. Paillard, Phys. Rev. B 2000, 62, 15942.
[42] a) A. Dondoni, Angew. Chem. 2008, 120, 9133–9135; Angew. Chem.
Int. Ed. 2008, 47, 8995–8997; b) C. E. Hoyle, C. N. Bowman, Angew.
Chem. 2010, 122, 1584–1617; Angew. Chem. Int. Ed. 2010, 49, 1540–
1573.
[43] C. E. Hoyle, A. B. Lowe, C. N. Bowman, Chem. Soc. Rev. 2010, 39,
1355–1387.
[44] a) A. W. Harant, V. S. Khire, M. S. Thibodaux, C. N. Bowman, Mac-
romolecules 2006, 39, 1461–1466; b) M. A. Caipa Campos, J. M. J.
Paulusse, H. Zuilhof, Chem. Commun. 2010, 46, 5512–5514; c) P.
Jonkheijm, D. Weinrich, M. Kohn, H. Engelkamp, P. C. M. Christi-
anen, J. Kuhlmann, J. C. Maan, D. Nusse, H. Schroeder, R. Wacker,
R. Breinbauer, C. M. Niemeyer, H. Waldmann, Angew. Chem. 2008,
120, 4493–4496; Angew. Chem. Int. Ed. 2008, 47, 4421–4424.
[19] a) R. S. Wallis, M. Pai, D. Menzies, T. M. Doherty, G. Walzl, M. D.
Perkins, A. Zumla, Lancet 2010, 375, 1920–1937; b) P. Tallury, A.
Malhotra, L. M. Byrne, S. Santra, Adv. Drug Delivery Rev. 2010, 62,
424–437; c) H. Takahashi, Y. Otsuka, B. Patterson, J. Infect. Chemo-
ther. 2010, 16, 155–161.
[20] A. MerkoÅi, Biosens. Bioelectron. 2010, 26, 1164–1177.
[21] C. Kumar, Nanomaterials for Biosensors, Wiley-VCH, 2006.
[22] U. Resch-Genger, M. Grabolle, S. Cavaliere-Jaricot, R. Nitschke, T.
Nann, Nat. Methods 2008, 5, 763–775.
[23] J. R. Lakowicz, Principles of Fluorescence Spectroscopy, 3rd ed.,
Springer, Singapore, 2006.
Chem. Asian J. 2011, 6, 2776 – 2786
ꢀ 2011 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
www.chemasianj.org
2785

J. M. J. Paulusse, H. Zuilhof et al.
FULL PAPERS
[45] a) N. ten Brummelhuis, C. Diehl, H. Schlaad, Macromolecules 2008,
41, 9946–9947; b) L. M. Campos, K. L. Killops, R. Sakai, J. M. J.
Paulusse, D. Damiron, E. Drockenmuller, B. W. Messmore, C. J.
Hawker, Macromolecules 2008, 41, 7063–7070.
[46] a) K. L. Killops, L. M. Campos, C. J. Hawker, J. Am. Chem. Soc.
2008, 130, 5062–5064; b) T. Kang, R. J. Amir, A. Khan, K. Ohshimi-
zu, J. N. Hunt, K. Sivanandan, M. I. Montanez, M. Malkoch, M.
Ueda, C. J. Hawker, Chem. Commun. 2010, 46, 1556–1558.
[47] a) M. J. Kade, D. J. Burke, C. J. Hawker, J. Polym. Sci. Part A 2010,
48, 743–750; b) L. M. Campos, I. Meinel, R. G. Guino, M. Schier-
horn, N. Gupta, G. D. Stucky, C. J. Hawker, Adv. Mater. 2008, 20,
3728–3733.
[50] W. W. Yu, L. Qu, W. Guo, X. Peng, Chem. Mater. 2003, 15, 2854–
2860.
[51] W. D. A. M. de Boer, D. Timmerman, K. Dohnalova, I. N. Yassie-
vich, H. Zhang, W. J. Buma, T. Gregorkiewicz, Nat. Nanotechnol.
2010, 5, 878–884.
[52] A. T. R. Williams, S. A. Winfield, J. N. Miller, Analyst 1983, 108,
1067–1071.
[53] D. Jurbergs, E. Rogojina, L. Mangolini, U. Kortshagen, Appl. Phys.
Lett. 2006, 88, 233116–233118.
[54] R. Anthony, U. Kortshagen, Phys. Rev. B 2009, 80, 115407–115412.
[55] F. A. Reboredo, G. Galli, J. Phys. Chem. B 2005, 109, 1072–1078.
[48] L. Ruizendaal, U. Chinnaswamy, L. Voorhaar, J. M. J. Paulusse, H.
Zuilhof, 2011, manuscript in preparation.
Received: April 13, 2011
Published online: August 24, 2011
[49] J. A. Kelly, E. J. Henderson, R. J. Clark, C. M. Hessel, R. G. Cavell,
J. G. C. Veinot, J. Phys. Chem. C 2010, 114, 22519–22525.
2786
www.chemasianj.org
ꢀ 2011 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
Chem. Asian J. 2011, 6, 2776 – 2786