Synthesis, characterization, DNA interaction and potential applications of gold nanoparticles functionalized with Acridine Orange fluorophores

Article abstract of DOI:10.1039/c0dt01371d

Two new water-soluble gold nanoparticles (AO-TEG-Au and AO-PEG-Au NPs) are prepared and characterized. They are stabilized by thioalkylated oligoethylene glycols and functionalized with fluorescent Acridine Orange (AO) derivatives. Despite the different core sizes (11.8 and 3.9 nm respectively) and shell composition, they are both well dispersed and are stable in water, even if some self-aggregation is observed in the case of AO-TEG-Au NPs. However, AO-PEG-Au NPs show much lower emission efficiency with respect to AO-TEG-Au NPs. Spectrophotometric and spectrofluorometric experiments indicate that both types of nanoparticle are able to bind to calf thymus DNA, either by external binding or partial intercalation. Preliminary FACS flow cytometry tests seem to indicate that the AO-TEG-Au nanoparticle is able to cross the cell membrane where it is absorbed by Chinese hamster ovary (CHO) cells at the picomolar concentration level.

Full text of DOI:10.1039/c0dt01371d

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PAPER
Synthesis, characterization, DNA interaction and potential applications of
gold nanoparticles functionalized with Acridine Orange fluorophores†
Tarita Biver,*a Nurettin Eltugral,a Andrea Pucci,*a Giacomo Ruggeri,a Alberto Schena,b Fernando Seccoa and
Marcella Venturinia
Received 11th October 2010, Accepted 9th February 2011
DOI: 10.1039/c0dt01371d
Two new water-soluble gold nanoparticles (AO-TEG-Au and AO-PEG-Au NPs) are prepared and
characterized. They are stabilized by thioalkylated oligoethylene glycols and functionalized with
fluorescent Acridine Orange (AO) derivatives. Despite the different core sizes (11.8 and 3.9 nm
respectively) and shell composition, they are both well dispersed and are stable in water, even if some
self-aggregation is observed in the case of AO-TEG-Au NPs. However, AO-PEG-Au NPs show much
lower emission efficiency with respect to AO-TEG-Au NPs. Spectrophotometric and
spectrofluorometric experiments indicate that both types of nanoparticle are able to bind to calf thymus
DNA, either by external binding or partial intercalation. Preliminary FACS flow cytometry tests seem
to indicate that the AO-TEG-Au nanoparticle is able to cross the cell membrane where it is absorbed by
Chinese hamster ovary (CHO) cells at the picomolar concentration level.
thioalkylated oligoethylene glycols and functionalized with two
1. Introduction
similar fluorescent Acridine Orange (AO) derivatives. The AO dye
Nano-sized metal particles show unique properties and growing
has been chosen because of its strong affinity to DNA and its
applications1–4 and several examples of the application of gold
intrinsic sensitivity and unambiguous fluorescence response upon
nanoparticles (Au NPs) have been reported.5–7 Functional groups
on the coating shell can modify the Au NPs properties, e.g. mod-
ulating their solubility8 and self-assembly9 or allowing selective
interactions.10 Different chromophores have been coupled to the
Au NPs for the design of sensors11 and as vehicles for tracers and
drugs.12 In recent years, Au NPs interacting with DNA have been
extensively studiedfor potentialapplications as, for example, smart
bio-based nanoparticle assemblies,9 new intercalating agents13 or
gene delivery vectors.3 Rotello and co-workers used functionalized
Au NPs to carry oligonucleotides inside the cell.5 In that case they
used oligonucleotides labelled with appropriate fluorophores to
detect and monitor the uptake process. Alternatively, Au NPs,
which are highly interacting with nucleic acids and traceable,
can be prepared through the functionalization of the stabilizing
capping layer with fluorescent moieties having a strong affinity for
nucleotides. We have, therefore, synthesized fluorescently labelled
gold nanoparticles showing both notable optical fluorescent
properties and high cellular permeability.
interaction with DNA.14–16 In addition strong gold binding by
thiolic groups is effective in preventing ligand leakage. One of the
binders uses the lipoic acid functionality, as the dithiolane ring is
able to chelate the gold surface with an affinity higher than that
of the single thiols.17,18 The interaction of the nanoparticles with
calf thymus DNA is analyzed and a preliminary test is performed
to verify the ability of the nanoparticles to cross the cellular
membrane in view of possible applications.
2. Materials and methods
2.1. Materials
The reagents used for the synthesis of Acridine Orange
functionalized ligands were 3,6-bis(dimethylamino)acridine hy-
drochloride (C17H19N3·HCl·xH2O, >99% Sigma-Aldrich), lipoic
acid (C8H14O2S2, >98% Sigma-Aldrich), 6-bromo-1-hexanol
(Br(CH2)6OH, >96% Fluka), N,N¢-dicyclohexylcarbodiimide
(DCC, >99% Fluka), 1,6-dibromohexane (Br(CH2)6Br, >97%
Fluka), and thiourea (NH2CSNH2, >99% Sigma-Aldrich). 1,6-
Dibromohexane and toluene (Aldrich) were distilled prior to use.
All the other solvents employed in the synthetic processes were
reagent grade (Carlo Erba or Aldrich) and used as received.
Calf thymus DNA (DNA, lyophilized sodium salt, highly
polymerized – Sigma) was dissolved in water and sonicated in
order to reduce its length. DNA samples (ca. 2 ¥ 10-3 M, 8
mL) were sonicated in a MSE-Sonyprep sonicator by applying
In the present work two new types of water-soluble Au
NPs are prepared and characterized. They are stabilized by
aDepartment of Chemistry and Industrial Chemistry, University of
Pisa, Via Risorgimento, 35-56126, Pisa, Italy. E-mail: tarita@
dcci.unipi.it, apucci@ns.dcci.unipi.it; Fax: 39-0502219260
b
´
Institute of Chemical Science and Engineering, Ecole Polytechnique
Fe´de´rale de Lausanne, Station 6, CH-1015, Lausanne
† Electronic supplementary information (ESI) available. See DOI:
10.1039/c0dt01371d
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twenty consecutive cycles of 10 s sonication and 20 s pause
with an amplitude of 98 mm. The sonicator tip was directly
introduced into the solution which was kept in an ice-bath
to minimize thermal effects caused by sonication. Agarose gel
electrophoresis tests indicated that the polymer length was reduced
to ca. 800 base pairs. Stock solutions of DNA were standardized
spectrophotometrically (e = 13200 M-1 cm-1 at 260 nm, 0.1 M NaCl
and pH = 7.0 for a molar DNA concentration, CDNA, expressed
in base pairs).19 Sodium chloride or sodium nitrate (Fluka) were
used to adjust the ionic strength and 0.01 M sodium cacodylate
(NaCac = (CH3)2AsO2Na, Fluka) was employed to keep the pH
of the solutions at the value of 7.0. The water used to prepare the
solutions and as a reaction medium was purified by a Millipore
Milli-Q water purification system.
anhydrous sodium sulfate. The product was purified by column
chromatography on silica gel, eluting with MeOH–CHCl3 mixture
(0 to 5%) to obtain the product as a bright orange oil (15 mg,
10.5%).
1H NMR (300 MHz, CDCl3) d (ppm): 8.69 (s, 1H), 7.90 (d,
2H, J = 9.29 Hz), 7.03 (dd, 2H, J = 1.48, 9.27 Hz), 6.55 (s, 2H),
4.73 (t, 2H, J = 7.55 Hz), 4.04 (t, 2H, J = 6.37 Hz), 3.51 (m, 1H),
3.29 (s, 12H), 3.07 (m, 2H), 2.40 (m, 1H), 2.27 (t, 2H, J = 7.18
Hz), range 1–2 (several m, 15H). 13C NMR (300 MHz, CDCl3) d
(ppm): 173.7, 155.7, 143.5, 142.7, 133.6, 177.2, 114.2, 92.7, 64.2,
56.5, 47.8, 41.0, 40.3, 38.6, 34.6, 34.1, 28.8, 28.6, 26.9, 26.3, 26.0,
24.7.
AO-TEG-Au nanoparticles. A solution of tetrachloroauric
acid (HAuCl4·3H2O, Sigma; 37 mg, 0.094 mmol) dissolved in
isopropanol (40 mL) was added to a 100 mL Erlenmeyer flask,
equipped with a magnetic stirrer. A solution of monohydroxy-(1¢-
mercaptoundec-11-yl)tetraethylene glycol (12 mg, 0.033 mmol)
dissolved in isopropanol (2 mL) and a solution of AHLB
acridinium ligand (7.5 mg, 0.012 mmol) also in isopropanol (1 mL)
were added. A freshly prepared solution of sodium borohydride
(117 mg, 3.1 mmol) in methanol (5 mL) was added dropwise over
1 min. The cloudy suspension became dark green–brown and was
left to react at room temperature for 4.5 h. The suspension was
poured into n-hexane (200 mL) and a dark brown precipitate
formed, that was cooled at -20 ◦C overnight. The supernatant
hexane was removed by centrifugation and the precipitate was
washed with diethyl ether and ethyl acetate. The precipitate
turned black and was dissolved in a few millilitres of water. The
suspension was then dialyzed through a membrane (Spectra/por
Floatalyzer MWCO = 3500) under continuous stirring at room
temperature for 20 days (deionized water in the flask was changed
daily). The disappearance of the free ligand was monitored by
UV-vis spectrophotometry. The dialyzed nanoparticle solution in
water was assumed to be the stock solution to be employed in
subsequent measurements. A schematic drawing of the obtained
nanoparticle (AO-TEG-Au) is shown in Fig. 1A.
6-(3¢,6¢-Bisdimethylaminoacridinium)hexan-1-ol
bromide
(AHB). The AO base was prepared by the addition of
ammonium hydroxide to a methanol solution of the AO·HCl
salt (99%, Sigma-Aldrich) and subsequent AO base extraction
with benzene. The organic phases were collected and dried over
anhydrous sodium sulfate and concentrated in vacuo. The AO
base (0.17 g, 0.64 mmol) in 8 mL anhydrous toluene was heated
to 80 ◦C under stirring for 30 min to obtain a homogeneous
solution to which 6-bromo-1-hexanol (1.0 g, 5.5 mmol) was
added drop◦wise over 5 min. The orange solution was heated to
reflux (110 C) and after 10 min a bright red precipitate started
to form. The suspension was refluxed overnight, and then filtered
over a 200 mm Teflon filter. The precipitate was washed with hot
toluene and dried in vacuo to obtain a bright red crude solid.
The crude product was purified by column chromatography on
silica gel, eluting with CH2Cl2–MeOH–AcOH = 95 : 5 : 1 to yield
6-(3¢,6¢-bisdimethylaminoacridinium)hexan-1-ol bromide (AHB)
as a bright red solid (100 mg, 35%).
1H NMR (300 MHz, CDCl3) d (ppm): 8.67 (s, 1H), 7.90 (d, 2H,
J = 9.29 Hz), 7.03 (dd, 2H, J = 1.48, 9.27 Hz), 6.63 (s, 2H), 4.79
(t, 2H, J = 7.55 Hz), 3.75 (t, 2H, J = 6.37 Hz), 3.33 (s, 12H), range
1–2 (several m, 8H).
6-(3¢,6¢-Bisdimethylaminoacridinium)-1-hexyl lipoate bromide
(AHLB).
A
three-necked round-bottomed flask equipped
with
a
magnetic stirrer, reflux condenser and pres-
a
a
10-(6-Bromohexyl)-3¢,6¢-bisdimethylaminoacridinium bromide
(BAHB). The AO base (◦0.41 g, 1.6 mmol) in 40 mL anhydrous
toluene was heated to 80 C under stirring for 30 min to obtain
a homogeneous solution. 1,6-Dibromohexane (3.80 g, 16 mmol)
was added to give an orange solution that was heated to reflux
(110 ◦C). After 20 min a bright red precipitate formed and the
suspension was allowed to reflux overnight. The supernatant
toluene was removed by centrifugation and the precipitate was
washed with toluene, which was then removed under reduced
pressure to give a bright red crude solid. The crude product
was purified by column chromatography on silica gel, eluting
with CH2Cl2–MeOH = 7 : 1 to yield 10-(6-bromohexyl)-3,6-
bisdimethylaminoacridinium bromide (BAHB) as a bright red
solid (0.58 g, 73%).
1H NMR (300 MHz, CDCl3) d (ppm): 8.75 (s, 1H), 7.91 (d, 2H,
J = 9.20 Hz), 7.00 (dd, 2H, J = 2.20, 9.40 Hz), 6.61 (s, 2H), 4.89
(t, 2H, J = 7.00 Hz), 3.45 (t, 2H, J = 6.60 Hz), 3.31 (s, 12H), 1.6–
2.0 (several m, 8H). 13C NMR (300 MHz, CDCl3) d (ppm): 155.6,
143.4, 142.7, 133.5, 177.2, 114.0, 93.0, 48.1, 41.2, 34.4, 32.5, 28.1,
26.3, 26.1.
sure equalizing dropping funnel were loaded with 6-(3¢,6¢-
bisdimethylaminoacridinium)hexan-1-ol bromide (AHB, 100 mg,
0.22 mmol), dimethylaminopyridine (DMAP, 25 mg, 0.21 mmol)
and lipoic acid (100 mg, 0.49 mmol) dissolved in 6 ml of
anhydrous dichloromethane. The mixture was stirred and, after
complete dissolution of the solid, the flask was cooled in an
ice-water bath. N,N¢-Dicyclohexylcarbodiimide (DCC, 150 mg,
0.73 mmol) dissolved in 2 mL of anhydrous dichloromethane
was added dropwise to the cooled solution over 10 min. The
solution was then allowed to reach room temperature and, after
10 min, a white solid precipitated. The mixture was stirred at room
temperature overnight (the temperature was maintained below
40 ◦C to prevent polymerization of the lipoic acid derivative). The
precipitate was filtered with a 200 mm Teflon filter and then washed
with dichloromethane (the lipoic acid derivative was dissolved in
dichloromethane to prevent polymerization). The organic phase
was washed first with 0.5 M HCl, followed by saturated sodium
bicarbonate solution and brine. After the extraction, the organic
phase was filtered to remove solid impurities and dried over
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Fig. 1 Schematic drawing of the AO-TEG-Au and AO-PEG-Au nanoparticles used in the present work.
Bis(6-(3¢,6¢-bisdimethylaminoacridinium)hexyl bromide) disul-
fide (BAHBS). Thiourea (45 mg, 0.36 mmol) was dissolved in
35 mL dry ethanol under an inert atmosphere. After complete
dissolution of thiourea under constant stirring, BAHB (0.25 g,
0.50 mmol) was added to the solution. The resulting mixture was
refluxed at 90 ◦C for 22 h. 15 mL aqueous sodium hydroxide
solution (34 mg, 0.85 mmol) was then added and the hydration
was carried out for an additional 4 h. The reaction mixture
was then poured into 150 mL aqueous HCl solution (pH =
4) and extracted with dichloromethane. The collected organic
phase was dried over Na2SO4 to remove any trace of water.
The dichloromethane phase was filtered off and the solvent
was evaporated under reduced pressure to obtain a crude oil.
The crude product was purified by column chromatography on
silica gel, eluting with CH2Cl2–MeOH = 7 : 1 to yield bis(6-(3¢,6¢-
bisdimethylaminoacridinium)hexyl bromide) disulfide (BAHBS)
(0.100 g, 22%).
brown precipitate was washed with diethyl ether. The precipitate
was then dissolved in 5 mL doubly distilled water and loaded
into a dialysis membrane (Spectra/por Floatalyzer MWCO =
3500). The suspension was dialyzed with continuous stirring at
room temperature for three days (deionized water in the flask
was changed daily). The disappearance of the free ligand was
monitored by UV-vis spectrophotometry. The final solution
of nanoparticles in water was assigned as the stock and kept
at 4 ◦C. A schematic drawing of the obtained nanoparticle
(AO-PEG-Au) is shown in Fig. 1B.
2.2. Apparatus and methods
◦
1H- and 13C NMR spectra were recorded at 20 C with a Varian
Gemini 300 MHz spectrometer using 5–10% analyte concentration
in CDCl3. Chemical shifts were assigned in ppm using the solvent
1
signal as a reference. H NMR was used to evaluate the ligand
1H NMR (300 MHz, CDCl3) d (ppm): 8.66 (s, 2H), 7.90 (d, 4H,
J = 9.30 Hz), 7.13 (d, 4H, J = 9.30 Hz), 6.80 (s, 4H), 4.89 (t, 4H,
composition of the prepared gold nanoparticles. In particular,
the fraction of the two different ligands that constitute the coating
layer was determined by comparing the integrated area of acridine
signals (6.5–9 ppm range) to the integrated area of ethylene glycol
protons (3.5–4 ppm range). This allows the molar fraction of the
acridinium derivative with respect to the total ligand amount to
be determined.
Fourier transform infrared (FT-IR) spectroscopy spectra were
recorded at room temperature with a Perkin-Elmer Spectrum One
spectrometer.
Transmission electron microscopy (TEM) experiments were car-
ried out either in a JEOL-1200 transmission electron microscope
using an acceleration voltage of 120 kV or in a Philips CM
200 electron microscope working at 200 kV. Scanning electron
microscopy (SEM) experiments were carried out by means of a
Field Emission ZEISS SUPRA 40 scanning electron microscope.
A single drop (10 mL) of a dilute aqueous solution of Au NPs
(ca. 0.1 mg mL-1) was placed on a copper grid coated with
a carbon film. The grid was allowed to dry in a desiccator
for several hours at room temperature. Size distributions of the
nanoparticle cores were measured from enlarged TEM images for
at least 200 individual cluster core images using the Image Tool
3.00 image analyzer software developed at the University of Texas
J = 7.50 Hz), 3.34 (s, 24H), 2.72 (t, 4H, J = 6.90 Hz), 1.6–2.0 ppm
-1
¯
(m, 16H). FTIR n/cm : 2918, 2852, 1640, 1602, 1523, 1503, 1364,
1166, 746, 518.
AO-PEG-Au
0.089 mmol) was dissolved in 50 mL methanol and
0.5 mL glacial acetic acid. Then, solution of O-(2-
nanoparticles. HAuCl4·3H2O
(35
mg,
a
mercaptoethyl)-O¢-methylhexa(ethylene glycol) (20 mg, 0.056
mmol) in methanol (2 mL) and a solution of bis(6-(3¢,6¢-
bisdimethylaminoacridinium)hexyl bromide) disulfide (BAHBS)
(8.3 mg, 0.009 mmol) also in methanol (4 mL) were added to
the gold solution and mixed under constant stirring. A freshly
prepared sodium borohydride (112 mg, 3.0 mmol) solution in
methanol (5 mL) was added dropwise to the resulting cloudy
suspension. The solution immediately turned from orange to
dark green–brown, revealing nanoparticle formation. After a
further 4 h of stirring, the solution was concentrated under
reduced pressure and re-dispersed in isopropanol (50 mL).
The particles were precipitated by pouring the mixture into
n-hexane. The solution mixture was left at -20 ◦C overnight.
The supernatant solution was centrifuged and the collected dark
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Table 1 Characteristic parameters of the AO-TEG-Au and AO-PEG-Au
nanoparticles synthesized in the present work
Health Science Center in San Antonio and available online at
http://ddsdx.uthscsa.edu/dig/itdesc.html.
X-Ray diffraction (XRD) patterns were obtained in Bragg-
Brentano geometry with a Siemens D500 KRISTALLOFLEX
810 (count time per step: 1.0 s; step size: 0.050◦ and Cu-Ka, l =
AO-TEG-Au
AO-PEG-Au
Average diameter from TEM (nm)
Av. no. Au in a NP core
Average MW (Da)
11.8 3.4
5.1 ¥ 104
1.2 ¥ 107
5
3.9 1.0
1.8 ¥ 103
5.2 ¥ 105
16
˚
1.541 A) diffractometer. Data were acquired at room temperature.
To evaluate the gold content (mg L-1) of the stock nanoparticle
solutions, atomic absorption (AA) measurements were made
on a Perkin Elmer Analyst 100 spectrophotometer. The stock
nanoparticle solution was dried under reduced pressure at 35 ◦C
to constant weight; then the recovered nanoparticles were re-
dispersed in precisely known volumes of water. To overcome
matrix effects of the nanoparticle solutions, the concentrations
were evaluated by the method of standard additions using special
grade standards (Aldrich).
Average TEG (PEG)/AO ratio
(from 1H NMR)
Av. no. AO in a NP
700
28
Av. no. TEG (PEG) in a NP
Av. no. total stabilizer tails in a NP
Au/total stabilizer ratio in a NP
3.7 ¥ 103
4.4 ¥ 103
12
4.5 ¥ 102
4.8 ¥ 102
4
subtracting the blank signal of the cells treated with pure PBS
buffer. Statistics were performed on 10000 cells. In the FACS
apparatus the wavelength of the exciting laser was 480 nm, while
the emission intensity was 520 nm. The high sensitivity required for
the low concentrations detected in our FACS experiments could
be attained due to the fact that the standard excitation wavelength
is very close to the absorption maximum of the AHLB ligand, and
that the maximum emission intensity of the acridinium moiety
almost coincides with the maximum intensity of the detector.
Spectrophotometric measurements were carried out on a Perkin
Elmer Lambda 35 spectrophotometer, while fluorescence measure-
ments wereperformedon aPerkin Elmer LS55spectrofluorometer,
both with temperature control to within 0.1 ◦C. The stability of
nanoparticles was monitored by UV-visible spectrophotometry,
using the surface plasmon resonance band.20 Absorbance and
fluorescence titrations of DNA/nanoparticle systems were done
by adding increasing amounts of the polynucleotide directly into
the cell containing the nanoparticles solution by means of a syringe
connected to a micrometric screw. In the case of the absorbance
measurements data have been collected in a differential manner;
namely, the same amount of DNA was placed in both the sample
and reference cuvettes, to compensate for the DNA contribution
to the measured signal in the UV range. AO-TEG-Au and AO-
PEG-Au NPs concentrations are expressed in AO molarity and
labelled as CAO. In this way a direct comparison can be made
between binding constants evaluated for the NPs/DNA systems
and for the AO derivative/DNA system.
3. Results
3.1. Nanoparticles synthesis
In the first synthetic approach (preparation of AO-TEG-Au
NPs), the AO was connected to the nanoparticle surface through
a linker between the 9 position of the acridine ring and
a terminal lipoic acid moiety.21,23 This functional ligand was
used together with the main stabilizing ligand monohydroxy(1-
mercaptoundec-11-yl)tetraethylene glycol (TEG-SH). The second
approach (preparation of AO-PEG-Au NPs) is based on the use
as stabilizing ligand of a mixture of the synthesized AO-hexyl
disulfide derivative (BAHBS)22,23 and O-(2-mercaptoethyl)-O¢-
methylhexa(ethylene glycol) (m-PEG-SH). Both TEG-SH and m-
PEG-SH provide highly hydrophilic shells.8 AHLB was prepared
following the synthetic procedure described in Scheme 1, whereas
BAHBS was synthesized according to Scheme 2.
Cellular uptake tests were made by means of fluorescence
activated cell sorting (FACS) flow cytometry analysis on Chinese
hamster ovary (CHO) immortalized cells. CHO cells were grown in
10 cm plates to confluency in F12 Ham’s media (10% FBS and 1¥
Pen/Strep). After one day the cells were left with 1 mL of trypsin
◦
and incubated for 5 min at 37 C. Then the cells were passaged:
14 well plates (1 for blank, 6 for free ligand, 7 for Au NPs) were
prepared with 25 mL of a diluted (1 : 40) tryptinized suspension
into 1 mL of media each. After three days the cells were confluent
and ready for the FACS analysis. The cellular uptake was evaluated
on the functionalized gold nanoparticles. Working solutions of the
Au NPs (0.14 to 9.6 nM) were prepared by dilution with phosphate
saline buffer (PBS) of a 50 nM stock, to reach the final volume
of 1 mL. For treatment of the cells each sample was prepared as
follows: the media were removed from the cell plate and the cells
were washed with 3¥ PBS; the washing buffer was removed and
the plate was treated with the buffered sample solution. Pure PBS
Au NPs were prepared following a single-phase reduction
method in alcoholic solution,8 as reported in detail above. The
disulfide is expected to behave in the same way as the correspond-
ing thiol, since it is well established that the chemisorption of an
n-alkyl disulfide on a Au NPs surface occurs in a similar manner
to that of the corresponding alkanethiol.24
3.2. Nanoparticles characterization
The dispersity of the particle distribution and their sizes was
determined by TEM analysis (Fig. 1S and 2S of ESI† for AO-TEG-
Au NPs and AO-PEG-Au NPs, respectively). The nanoparticles
were found to be spherical to a good approximation with mean
core diameters of 11.8 3.4 nm for AO-TEG-Au NPs and 3.9
1.0 nm for AO-PEG-Au NPs (Table 1). For the AO-PEG-Au
nanoparticles SEM and XRD experiments were also performed
(Fig. 3S and 4S of ESI†) that yielded a mean nanoparticle diameter
of 4.6 0.8 nm (SEM) and 5.8 nm (XRD, on applying Scherrer’s
equation on the Au(111) peak).25 The results obtained by the
different techniques are in fairly good agreement.
◦
was added to the blank plate. The cells were incubated at 37 C
for 1 h; then the solutions were removed and the cells were washed
several times with 5¥ PBS and 1 mL of 1¥ trypsin was added.
After 10 min incubation at 37 ◦C the suspension was transferred
into FACS tubes and spun down by 10 min centrifugation at 1000
rpm. The supernatant media was removed from the pellets and
500 mL of BSA medium (0.1% Bovine Serum Albumin + 5 mM
EDTA in PBS) were added, shaking to re-suspend the cells. Each
solution was analyzed with a BD FACS Calibur flow cytometer,
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Scheme 1 Synthesis of acridinium lipoate (AHLB). Reagents and conditions: (a) 9 eq. 6-bromo-1-hexanol, toluene, 110 ◦C, 24 h; (b) 2.2 eq. lipoic acid,
3.3 eq. DCC, 1 eq. DMAP, CH2Cl2, r.t., 18 h.
Scheme 2 Synthesis of bis(6-(3¢,6¢-bisdimethylaminoacridinium)hexyl bromide) disulfide (BAHBS). Reagents and conditions: (a) 10 eq. 1,6-dibromohex-
ane in toluene, reflux at 110 ◦C, overnight, (b) 0.72 eq. thiourea in ethanol, reflux at 85 ◦C for 22 h; then 1.7 eq. NaOH/H2O, reflux for 5 h.
The mean number of gold atoms in a particle (n) was calculated
Therefore, the NPs absorbance data have been corrected for a light
scattering (LS) contribution, following the procedure described
by Leach and Scheraga that uses double logarithmic plots.30 One
example of spectrum correction is provided in Fig. 8S of ESI†. The
two systems display analogous absorption characteristics, with
two maxima in the visible region, one centered at around 475 nm
independently on the nature of the NPs, and the other at 496 nm
for AO-TEG-Au and at 507 nm for AO-PEG-Au NPs (Fig. 2). The
presence of two maxima is observed for similar AO derivatives31
and for AHLB itself (Fig. 9S of ESI†), whereas plasmon bands
centered around 500 nm are typical of gold nanoparticles of
diameter close to that measured in this work.29
As regards the AO-PEG-Au system, the absorption ratio of the
two maxima at different NP concentrations is constant, indicating
that NPs do not self-aggregate in the investigated concentration
range (inset in Fig. 2B). On the other hand, both the AO-TEG-
Au NPs and the AHLB molecule undergo some self-aggregation
(dimerisation) at higher dye concentrations, as confirmed by the
increase of the absorbance ratio A475/A496 (insets of Fig. 2A
and 9S (ESI†) respectively). The quantitative analysis of the
self-aggregation process, described by the simplified dimerisation
process shown in reaction (2), can be done by fitting of the data to
eqn (3) and (4)
using eqn (1)26,27
3
0.5pN Adm
n =
(1)
3Vm
where a spherical particle shape is assumed, NA is Avogadro’s
number, dm is the diameter of the nanoparticle and Vm is the molar
volume of gold(10.2 cm3). The gold content (mg L-1) and the mixed
monolayer composition were determined by the combined use of
1
AA and H NMR measurements (see the experimental section
and Fig. 5S and 6S of ESI† for AO-TEG-Au NPs and AO-PEG-
Au NPs, respectively). These data allowed the average molecular
weight and composition of clusters to be estimated (Table 1).
The diameter of the AO-TEG-Au NPs with [Au]/[total stabilizer]
ratio equal to 12 is about three times larger than the diameter
of the AO-PEG-Au NPs with [Au]/[total stabilizer] ratio equal
to 4 (Table 1). This behavior is in agreement with similar results
reported in the literature for thiol-stabilized Au NPs, which shows
that NPs increase significantly their diameter when the [Au]/[total
stabilizer] ratio becomes higher than 3.28
The Au NPs are found to be well dispersed in water and
their stability against both decomposition and/or aggregation was
checked using optical spectroscopy. AO-TEG-Au NPs solutions
are stable in pure water, but precipitate on addition of salt (0.1 M
NaCl or 0.1 M NaNO3). On the other hand, AO-PEG-Au NPs are
stable both in pure water and 0.1 M NaCl solutions. Absorbance
spectra reveal that both NPs do significantly scatter light (Fig. 7S
of ESI†), in agreement with previous findings on similar systems.29
K
D
(2)
ꢀꢀꢀꢁ
2 M
D
ꢂꢀꢀꢀ
A = eM[M] + eD[D]
(3)
(4)
C
AO/(A - eMCAO) = 2/De + (1/KDDe)(1/[M])
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Fig. 2 Absorbance spectra of AO-TEG-Au NPs (A) and of AO-PEG-Au NPs (B) once corrected for light scattering contribution at various NP
concentrations, T = 25 ◦C. (A) CAO = 3.6 ¥ 10-7–2.9 ¥ 10-6 M, pH = 7.0, I = 0.01 M (NaCac); (B) CAO = 3.4 ¥ 10-7–5.2 ¥ 10-6 M, pH = 7.0, I = 0.1 M
(NaCl). The insets are plots of the ratio of the two maxima with respect to NP concentration.
where M is the nanoparticle monomer, D its dimer form, A is
the experimental absorbance (corrected for LS in the case of AO-
TEG-Au NPs), eM and eD are the molar extinction coefficients of
monomer and dimer respectively, De = eD - 2eM and CAO is the total
nanoparticle concentration (expressed in AO concentration). For
the derivation of eqn (4) see ref. 32. First, a rough estimate of eM was
obtained by the spectra and used to plot CAO/(A - eMCAO) vs. 1/[M]
that, in this first estimation, was set equal to 1/CAO. A linear fit of
the data plot according to eqn (4) enabled KD and De be evaluated.
Then, KD was used to calculate [M] and [D] values for a 3D fit
according to eqn (3) that yielded better eM and eD estimates, and the
emission of the AO-TEG-Au NP/DNA complex (520 nm) is the
same as exhibited by the AO/DNA complex,35 confirming that
the acridine moiety is the main DNA binder.
It can be calculated that only 55% of AO-TEG-Au NPs are
present as monomer under the conditions of DNA binding
experiments; therefore, the interaction between nanoparticles and
DNA is expressed by the system of reactions (2), (5) and (6)
M + S ꢀ MS
(5)
D + S ꢀ DS
(6)
procedure is repeated until convergence is reached. For the AO-
TEG-Au NPs it is found that eM
eD
for AHLB eM
= (2.80 0.05)¥105 M-1 cm-1,
475 nm
where the assumption is made that the nanoparticle could react
with a DNA site (S) both in the form of the monomer (M) and
in the form of dimer (D), to give the bound species MS and DS
respectively. The equilibrium constants of (5) and (6) are defined as
Kmon = [MS]/([M][S]) and Kdim = [DS]/([D][S]). The titration data
were analyzed using eqn (7), that can be obtained on the basis of
reactions (2), (5) and (6) (for its derivation see ESI†).
= (9.47 0.09) ¥ 105 M-1 cm-1 and KD = (6.3 0.3) ¥ 105 M-1;
475 nm 475 nm
475 nm
= (2.80 0.06) ¥ 104 M-1 cm-1, eD
= (1.82
0.07) ¥ 105 M-1 cm-1 and KD = (2.7 0.2) ¥ 103 M-1. The features of
the AHLB self-aggregation process here observed are in agreement
with those displayed by AO, proflavine and their alkyl-derivated
compounds.16,33,34 The extinction coefficient of AO-PEG-Au NPs
in 0.1 M NaCl (Fig. 2B) at 507 nm is e = 4.2 ¥ 105 M-1 cm-1 (not
corrected for LS) or e = 1.9 ¥ 105 M-1 cm-1 (after correction for
LS).
A
eD
2
De1 + De2Kmon[S]+ De3KDKdim[M][S]
1+ 2KD[M]+ Kmon[S]+ 2KDKdim[M][S]
=
+
(7)
CAO
where [S] is the free DNA concentration and [M] the free monomer
concentration, De1 = eM – (eD/2), De2 = eMS – (eD/2) and De3 = eDS
– eD. The 3D data fit to eqn (7) requires an iterative procedure as
[S] and [M] have to be calculated using the values of KD, Kmon and
3.3. DNA interaction
AO-TEG-Au NPs/DNA. The binding of AO-TEG-Au NPs
to DNA was investigated by spectrophotometric and spectrofluo-
rometric titrations. The interaction is revealed by changes in the
absorbance and fluorescence spectra: NP absorbance decreases
and fluorescence increases upon addition of increasing amounts
of DNA to the NP sample present in the measuring cell (Fig. 3A
and 4A). As regards absorbance data, the experimental spectra
are shown in Fig. 10S of ESI†, whereas Fig. 3A show the results
of the LS correction. The behavior below 300 nm and the binding
isotherm at 270 nm (Fig. 3) reveals that the binding process is
biphasic, thus suggesting the occurrence of two interaction modes
(let’s define them as binding mode (1) and binding mode (2)). On
the other hand, the isotherm at 475 nm (Fig. 3C) is monophasic
and provides information on binding mode (1). Binding mode (2)
was investigated by spectrofluorometry (Fig. 4). The maximum
K
dim, where KD only is known. The same procedure was used for
fluorescence titrations, by replacing A, eD and Dei by F, jD and
Dji respectively. Both analyses evidenced that the De3 (Dj3) and
Kdim contributions can be neglected, indicating NP binds to DNA
as a monomer only. Under such circumstances eqn (7) reduces to
eqn (8)
A
eD
2
De1 + De2Kmon[S]
1+ 2KD[M]+ Kmon[S]
=
+
(8)
CAO
The obtained values of Kmon = K are collected in Table 2. Note
that, as the number of Acridine Orange molecules contained in
each nanoparticle cluster is known (nAO, Table 1), the numerical
value of K given in Table 2 can be converted to the binding affinity
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Fig. 3 (A) Absorbance spectra of the AO-TEG-Au NPs/DNA system (corrected for LS), CAO = 1.2 ¥ 10-6 M, pH = 7.0, T = 25 ◦C, (a) CDNA = 0, (b)
CDNA = 6.8 ¥ 10-5 M; (B) binding isotherm at 270 nm; (C) binding isotherm at 475 nm.
Fig. 4 (A) Fluorescence spectra collected for AO-TEG-Au NPs/DNA, CAO = 1.2 ¥ 10-6 M; pH = 7.0, T = 25 ◦C. (a) CDNA = 0, (b) CDNA = 6.8 ¥ 10-5 M;
(B) binding isotherm at lex = 480 nm, lem = 520 nm.
of one DNA base pair to one cluster (K¢) simply by using the
relationship K¢ = nAOK.
In the case of the AHLB/DNA system less than 5% dimer is
present under conditions of DNA binding experiments and the
KD term in eqn (8) can be neglected; eqn (8), once rearranged,
becomes eqn (9).
Control experiments have been done on the interaction with
DNA of the free AO derivative (AHLB) in the absence of gold
nanoparticles, to be compared with the behavior of the AO-TEG-
Au/DNA system. The absorbance spectra of the AHLB/DNA
system at different polymer to dye ratios show two different
behaviors indicating that two modes of binding are operative
(Fig. 5). In particular, upon addition of DNA, an absorbance
decrease is first observed till a threshold AHLB/DNA ratio is
attained, and then an absorbance recovery occurs with a red-shift
of the peak centered at 500 nm.
DA
KDe[S]
=
(9)
CAO (1+ K[S])
where DA = A – A0 and A0 = eMCAO is the absorbance of the NP
solution in the absence of DNA. De = eMS – eM is the amplitude of
binding isotherm. Initially, the total DNA concentration, CDNA, is
introduced in eqn (9) in place of [S] to obtain a first estimation of
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Table 2 Binding constants to DNA for the two analyzed nanoparticles and for the free AHLB derivative at pH = 7.0, T = 25 ◦
K/M-1
C
System
Spectrophotometry
Spectrofluorimetry
a
AO-TEG-Au NPs/DNA
AO-PEG-Au NPs/DNAc
AHLB/DNA
5.4 ¥ 106 (K1)
—
b
3.3 ¥ 104 (K2)
3.8 ¥ 104
—
>106 (K1)
<105 (K2)
2.0 ¥ 104
—
>1.0 ¥ 106 (K1)
3.5 ¥ 104 (K2)
6.3 ¥ 104
a Fit to eqn (8) of the data at 475 nm. b Fit to eqn (8) of the second part of the data at 270 nm. c I = 0.1 M (NaCl).
Fig. 5 (A) Absorbance spectra of the AHLB/DNA system, CAO = 1.3 ¥ 10-5 M, pH = 7.0, T = 25 ◦C, (a) CDNA = 0 M, (b) CDNA = 5.9 ¥ 10-6 M, (c)
CDNA = 1.5 ¥ 10-4 M. (B) Corresponding binding isotherm revealing the biphasic behavior of the binding process at 496 nm (the inset is the expansion of
the descending portion of the plot). The continuous line represents the trend calculated according to eqn (9).
K that can be used to calculate [MS] and [M] = CDNA – [MS].
Then, K and De are re-evaluated by introducing [M] in eqn
(9) and the procedure is repeated until convergence is reached.
The two branches of the binding isotherm were analyzed with
the help of eqn (9) and the equilibrium constants obtained are
shown in Table 2. Fluorescence titrations, done with much more
diluted amounts of AHLB (1.9 ¥ 10-7 M) are, on the other hand,
monophasic (Fig. 11S of ESI†). The dye fluorescence increases
upon DNA addition, as is also observed in the case of the
AO/DNA system.36 The value of K, obtained by data fit to eqn
(9), is collected in Table 2.
3.4. Cellular uptake tests of the AO-TEG-Au NPs
Preliminary cellular uptake tests were carried out by means of the
FACS flow cytometry technique, in order to check if the ability
of the AO-TEG-Au NPs to bind nucleic acids could make them
suitable for potential applications in biology. It has to be noted
that the AO-TEG-Au NPs are self-emissive and, furthermore, the
interaction with the nucleic acids can be real-time monitored
because of the spectroscopic features widely discussed in the
previous sections. A preliminary study was performed on the
free ligand AHLB and even at very low concentrations (10 nM
AHLB in PBS) saturation conditions were reached; actually,
all the Chinese hamster ovary (CHO) cells incubated in the
presence of the dye have been shown to incorporate the fluorescent
molecule. Then the uptake of the AO-TEG-Au NPs was tested.
The FACS analysis was performed on several identical samples
of CHO immortalized cells incubated in solutions at variable
concentrations of the AO-TEG-Au NPs. The results are shown in
Fig. 6. Both the shape of the histograms and the concentration
dependence reveal nanoparticle uptake. At CAO = 1.4 nM, a
bimodal distribution is observed. A shoulder in the histogram at
higher emission intensity appears, relative to the occurrence of the
uptake of NPs. Below 1.4 nM the uptake is negligible, as indicated
by the strongly reduced level of cell fluorescence. At 2.8 nM almost
all the cells have shown to contain the AO-TEG-Au NPs and above
7 nM saturation of the system is reached, such that the maximum
number of functionalized nanoparticles has entered living cells
AO-PEG-Au NPs/DNA. Several spectroscopic titrations
were carried out in 0.1 M NaCl to investigate AO-PEG-Au
NPs binding to DNA. The absorbance titration isotherms reveal
the presence of two interaction modes (Fig. 12S of ESI†) but
the variation of absorbance is too low to enable a quantitative
evaluation of the reaction parameters; only lower and upper limits
for K1 and K2, respectively, could be estimated (Table 2). The
spectrofluorometric method is more sensitive to investigate the
binding, but with this technique one binding mode only can be
observed (Fig. 13S of ESI†). Fit of the data to eqn (9) yields
the binding constants in Table (2). It is worth to emphasize that,
differently from AO-TEG-Au NPs, AO-PEG-Au NPs are almost
not fluorescent when in solution and it is only when DNA is added
that the green–yellowish acridinium fluorescence becomes readily
observable.
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Fig. 6 (A) Statistical distribution of the FACS data at different nanoparticle concentrations (in AO molarity): cell count vs. fluorescence intensity at
520 nm for CAO = 0 (red), 0.14 (orange), 0.7 (yellow), 1.4 (green), 2.8 (cyan), 7.0 (blue) and 9.6 (violet) nM. (B) Emission intensity relative to the maximum
of the distribution of the cell count vs. the AO-TEG-Au NP concentration.
and no significant change is observed if the concentration of the
incubation solution is higher.
2.7, whereas for DNA/AO-TEG-Au NPs and DNA/AO-PEG-
Au NPs absorbance titrations n = 1.7–2.0 and n = 1.5, respectively.
The decrease of the n value in the latter cases indicates that the
size of the DNA/NPs systems is larger than that of the NPs alone.
The fluorescence characteristics of the two nanoparticles differ
from each other. The fluorescence of both AO moieties attached
to the NPs decreases considerably compared to the fluorescence
intensity of free AHLB. However, although AO-TEG-Au NPs
still moderately fluoresce, AO-PEG-Au NPs emit a negligible
luminescence. This might be a consequence of some fluorescence
quenching by the gold core which depends on the chain length of
the AO linker.13,37 Actually, previous studies showed that the effi-
ciency of the fluorescence emission of the chromophore strongly
depends on the metal surface–fluorophore distance (d).38,39 It has
been observed that strong quenching of fluorescence emission and
a dramatic reduction of the lifetimes of the excited states occur
when d < 5 nm, whereas these effects are not observed for d >
10 nm.
As regards the interaction with natural DNA, the results show
that this does take place. Indeed, it is suggested that the length of
the ligand was adequately chosen to avoid steric hindrance effects
by the TEG-SH tethers on the AO intercalating effectiveness. The
discussion on the behavior of the nanoparticle/DNA systems is
made easier by comparison with the AHLB/DNA system. The
acridine derivatives are known to differently bind to DNA giving
rise either to external or intercalated complexes, depending on the
dye/DNA ratio.16,35,40–43 Our data on the AHLB/DNA system also
reveal the occurrence of two binding modes, the first one (K1 >
106 M-1) being likely related to external binding under dye excess
conditions, the second one (K2 = 3.5 ¥ 104 M-1) corresponding
to AO intercalation between DNA base pairs. Concerning the
intercalative mode of binding, our K values (Table 1) are in good
agreement with literature data on the AO/DNA system under
similar conditions.16,44,45
4. Discussion
Acridine orange functionalized water-soluble gold nanoparticles
have been successfully synthesized using different ethylene glycol
based ligands and acridine derivatives. They have different core
sizes (11.8 nm for AO-TEG-Au NPs and 3.9 nm for AO-PEG-
Au NPs) and shell composition, but are both stable against
precipitation in water over a period of months when kept at
+4 ◦C. AO-PEG-Au NPs are stable also in the presence of added
salts (0.1 M NaCl or NaNO3), whereas AO-TEG-Au NPs stability
is sensitive to electrolyte concentration (regardless of salt type). As
Acridine Orange self-association is promoted when salt is added
to aqueous dye solutions, this effect could induce nanoparticle
precipitation. In particular, in the case of the AO-TEG-Au NPs,
the higher number of AO residues in the shell (700 against 28
for the AO-PEG-Au system) could be the cause of the higher
self-aggregation tendency of this (bigger) nanoparticle; the same
effect is actually less probable in case of the smaller AO-PEG-Au
NPs where, moreover, AO is more buried into the PEG ligand. The
above qualitative findings agree with the results of the quantitative
analysis of the absorbance features of the two nanoparticles at
different NP concentrations. Indeed, it is found that AO-PEG-Au
NPs do not self-aggregate, whereas some aggregation phenomena
occur in the case of the AO-TEG-Au NPs. The dimerisation
constant obtained for the latter NP system is significantly high
(KD = 6.3 ¥ 105 M-1) with respect to the values generally found for
acridine dyes (for AHLB alone KD = 2.7 ¥ 103 M-1). This finding
might be explained by taking into account that the presence of the
nanoparticles produces a non-homogeneous distribution of the
AO dye in the reaction medium that is locally concentrated on the
NP.
For both nanoparticles it is found that, even if light absorption
gives the main contribution to the observed spectral behavior,
the light scattering (LS) component cannot be neglected. The
LS contribution is proportional to l-n. It is found that for AO-
TEG-Au NPs n = 2.3–2.4 and for AO-PEG-Au NPs n = 2.3–
Absorbance titration results obtained for the AO-TEG-Au
NPs/DNA system (K = K1 = 5.4 ¥ 106 M-1), compared with
results obtained for the free AO derivative, suggest that AO-TEG-
Au NPs are able to interact with DNA by external binding. On
the other hand, the relatively low value of the binding constant
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(K = K2 = 3.8 ¥ 104 M-1) obtained from fluorescence data seems to
indicate that some intercalation of the AO residue attached to the
nanoparticle is also occurring.45,46 Fluorescence data of the AO-
PEG-Au NPs/DNA system show similarities with AO-TEG-Au
NPs/DNA which suggest a dual mode of binding for the former
system as well. The results show that the Acridine Orange residue,
even if linked to a bulky substituent (nanoparticle), is able to
intercalate into DNA, although the AO intercalation should be
only partial. It was, indeed, found that the binding constant for
intercalation into DNA of AO, under conditions similar to that
used in present work, is close to 105 M-1,45 whereas addition of
substituents in the opposite position with respect to the aromatic
nitrogen decreases the binding affinity (K £ 8 ¥ 104 M-1),46 the latter
finding being in agreement with the value obtained for AHLB
intercalation into DNA (Table 2). If one reasonably supposes that
a higher binding affinity reflects a deeper dye insertion, it can be
suggested that dye penetration follows the order AO > AHLB >
AO-Au NP.
Preliminary FACS flow cytometry tests indicate the AO-TEG-
Au NPs are able to cross the cell membrane and are absorbed by
the CHO cells at quite a low concentration. These results for the
AO-TEG-Au NPs agree with previous findings that indicate that
gold nanoparticles undergo endocytosis and that high NP uptake
occurs under experimental conditions similar to those used in the
present work (also important is likely to be the higher adhesion
to cell membranes of cationic Au NPs with respect to anionic Au
NPs).47–49 Our results suggest that the absorption could occur on
a picomolar scale with respect to the whole NP, opening the way
for potential uses in cellular biology.
10 X. Li, J. Wang, L. Sun and Z. Wang, Chem. Commun., 2010, 46, 988–
990.
11 S. Ribrioux, G. Kleymann, W. Haase, K. Heitmann, K. Ostermeier and
H. J. Michel, J. Histochem. Cytochem., 1996, 44, 207–213.
12 U. H. F. Bunz and V. M. Rotello, Angew. Chem., Int. Ed., 2010, 49,
3268–3279.
13 G. Wang, J. I. Zhang and R. W. Murray, Anal. Chem., 2002, 74, 4320–
4327.
14 B. H. Robinson, A. Lo¨ffler and G. Schwarz, J. Chem. Soc., Faraday
Trans. 1, 1973, 69, 56–69.
15 V. V. Tscharner and G. Schwarz, Biophys. Struct. Mech., 1979, 5, 75–90.
16 J. Kapuscinski and Z. Darzynkiewicz, J. Biomol. Struct. Dyn., 1987, 5,
127–143.
17 J. Sharma, R. Chhabra, C. S. Andersen, K. V. Gothelf, H. Yan and Y.
Liu, J. Am. Chem. Soc., 2008, 130, 7820–7821.
18 G. Han, T. Mokari, C. Ajo-Franklin and B. E. Cohen, J. Am. Chem.
Soc., 2008, 130, 15811–15813.
19 H. R. Mahler, B. Kline and B. D. Mehrotra, J. Mol. Biol., 1964, 9,
801–11.
20 P. Mulvaney, Langmuir, 1996, 12, 788–800.
21 C. Courvoisier, M. J. Paret, J. Chantepie, J. Gore´, G. Fournet and G.
Quash, Bioorg. Chem., 2006, 34, 49–58.
22 E. Miethke and V. Zanker, Z. Phys. Chem., 1958, 18, 375–390.
23 Y. Kubo, S. Uchida, Y. Kenmochi and T. Okubo, Tetrahedron Lett.,
2005, 46, 4369–4372.
24 L. A. Porter, D. Jr. Ji, S. L. Westcott, M. Graupe, R. S. Czernuszewicz,
N. J. Halas and T. R. Lee, Langmuir, 1998, 14, 7378–7386.
25 H. P. Klug and L. E. Alexander, in X-ray Diffraction Procedures: for
Polycrystalline and Amorphous Materials, 2nd edn, J. Wiley and Sons
Inc., New York, 1974.
26 A. Henglein and M. Giersig, J. Phys. Chem. B, 1999, 103, 9533–
9539.
27 U. Hotje, C. Rose and M. Binnewies, Solid State Sci., 2003, 5, 1259–
1262.
28 A. I. Frenkel, S. Nemzer, I. Pister, L. Soussan and T. Harris, J. Chem.
Phys., 2005, 123, 184701.
29 T. Shimizu, T. Teranishi, S. Hasegawa and M. Miyake, J. Phys. Chem.
B, 2003, 107, 2719–2724.
In summary, a multifunctional probe is able to bind to DNA
with a characteristic photoresponse in both absorbance and
emission and can cross the cellular membrane and be easily
detected inside the cell.
30 S. J. Leach and H. A. Scheraga, J. Am. Chem. Soc., 1960, 82, 4790–4792.
31 B. E. Bowler, K. J. Ahmed, W. I. Sundquist, L. S. Hollis, E. E. Whang
and S. J. Lippard, J. Am. Chem. Soc., 1989, 111, 1299–1306.
32 T. Biver, B. Garc´ıa, J. M. Leal, F. Secco and E. Turriani, Phys. Chem.
Chem. Phys., 2010, 12, 13309–13317.
33 G. G. Hammes and C. D. Hubbard, J. Phys. Chem., 1966, 70, 2889–
2895.
Acknowledgements
34 A. D. James and B. H. Robinson, Adv. Mol. Relax. Processes, 1976, 8,
Prof Yitzhak Tor, Department of Chemistry & Biochemistry,
University of California, San Diego, is kindly acknowledged
for the research stage conducted by A. Schena at the UCSD
laboratory and for helpful discussions. The authors thank Prof.
B. H. Robinson for assistance in the editing of the English
language.
287–304.
35 M. B. Lyles and I. L. Cameron, Biophys. Chem., 2002, 96, 53–76.
36 E. Fredericq and C. Houssier, Biopolymers, 1972, 11, 2281–2308.
37 K. George Thomas and P. V. Kamat, Acc. Chem. Res., 2003, 36, 888–
898.
38 U. H. F. Bunz and V. M. Rotello, Angew. Chem., Int. Ed., 2010, 49,
3268–3279.
39 K. Aslan and V. H. Perez-Luna, J. Fluoresc., 2004, 14, 401–405.
40 C. Ciatto, M. L. D’Amico, G. Natile, F. Secco and M. Venturini,
Biophys. J., 1999, 77, 2717–2724.
References
41 T. Biver, C. Ciatto, F. Secco and M. Venturini, Arch. Biochem. Biophys.,
2006, 452, 93–101.
1 M. Han, X. Gao, J. Z. Su and S. Nie, Nat. Biotechnol., 2001, 19, 631–
635.
42 B. Garcıa, J. M. Leal, R. Ruiz, T. Biver, F. Secco and M. Venturini,
´
2 M. E. Stewart, C. R. Anderton, L. B. Thompson, J. Maria, S. K. Gray,
J. A. Rogers and R. G. Nuzzo, Chem. Rev., 2008, 108, 494–521.
3 A. K. Boal, F. Ilhan, J. E. Derouchey, T. Thurn-Albrecht, T. P. Russell
and V. M. Rotello, Nature, 2000, 404, 746–748.
4 S. K. Ghosh and T. Pal, Chem. Rev., 2007, 107, 4797–4862.
5 P. S. Ghosh, C.-K Kim, G. Han, N. S. Forbes and V. M. Rotello, ACS
Nano, 2008, 2, 2213–2218.
J. Phys. Chem. B, 2010, 114, 8555–8564.
43 C. Bazzicalupi, A. Bencini, A. Bianchi, T. Biver, A. Boggioni, S.
Bonacchi, A. Danesi, C. Giorgi, P. Gratteri, A. Marchal Ingraın, F.
Secco, C. Sissi, B. Valtancoli and M. Venturini, Chem.–Eur. J., 2008,
14, 184–196.
´
44 S. Nafisi, A. A. Saboury, N. Keramat, J. Neault and H. Tajmir-Riahi,
J. Mol. Struct., 2007, 827, 35–43.
6 J. J. Storhoff and C. A. Mirkin, Chem. Rev., 1999, 99, 1849–1862.
7 D. A. Giljohann, D. S. Seferos, W. L. Daniel, M. D. Massich, P. C. Patel
and C. A. Mirkin, Angew. Chem., Int. Ed., 2010, 49, 3280–3294.
8 A. Kanaras, F. S. Kamounah, K. Schaumburg, C. J. Kiely and M.
Brust, Chem. Commun., 2002, 2294–2295.
9 A. P. Alivisatos, K. P. Johnsson, X. Peng, T. E. Wilson, C. J. Loweth,
M. Bruchez and P. J. Schultz, Nature, 1996, 382, 609–611.
45 W. Muller and D. M. Crothers, Eur. J. Biochem., 1975, 54, 267–277.
46 W. Muller, H. Bunemann and N. Dattagupta, Eur. J. Biochem., 1975,
54, 279–291.
47 K. Song, P. Xu, Y. Meng, J. Chen, X. Yang, W. Roa, B. Kong and J.
Xing, IEEE/NIH Life Sci. Syst. Appl. Workshop, 2009, 33–35.
48 K. Y. Win and S.-S. Feng, Biomaterials, 2005, 26, 2713–2722.
49 J. Lin, H. Zhang, Z. Chen and Y. Zheng, ACS Nano, 2010, 4, 5421–5429.
This journal is
The Royal Society of Chemistry 2011
Dalton Trans., 2011, 40, 4190–4199 | 4199
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