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Nano Res
1
Aqueous Self-Assembly and Surface-Functionalized
Nanodots for Live Cell Imaging and Labeling
Mei-Lang Kung,1 Pei-Ying Lin,
1 Chiung-Wen Hsieh,
1 and Shuchen Hsieh
1, 2 ()
Nano Res., Just Accepted Manuscript • DOI: 10.1007/s12274-014-0479-y
http://www.thenanoresearch.com on April 21, 2014
© Tsinghua University Press 2014
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Nano Research
DOI 10.1007/s12274-014-0479-y
1
Aqueous Self-Assembly and Surface-Functionalized
Nanodots for Live Cell Imaging and Labeling
Mei-Lang Kung, Pei-Ying Lin, Chiung-Wen Hsieh, and
Shuchen Hsieh*
“National Sun Yat-sen University”, Kaohsiung
Schematic illustration of a conjugated self-assembled nanodot (SAND)
system for targeted cell imaging
2
Aqueous Self-Assembly and Surface-Functionalized Nanodots for Live Cell Imaging and Labeling
Mei-Lang Kung,
1 Pei-Ying Lin,
1 Chiung-Wen Hsieh,
1 and Shuchen Hsieh
1, 2 ()
1Department of Chemistry and Center for Nanoscience and Nanotechnology, “National Sun Yat-sen University”, Kaohsiung, 80424 2School of Pharmacy, College of Pharmacy, Kaohsiung Medical University, Kaohsiung, 80707
Received: day month year / Revised: day month year / Accepted: day month year (automatically inserted by the publisher)
© Tsinghua University Press and Springer-Verlag Berlin Heidelberg 2011
ABSTRACT Nanoparticles have enormous potential for bioimaging and biolabeling applications, in which conventional
organically based fluorescent labels degrade and fail to provide long-term tracking. Thus, the development of
approaches to make fluorescent probes water soluble and label cells efficiently is desirable for most biological
applications. Here, we report on the fabrication and characterization of self-assembled nanodots (SANDs) from
3-aminopropyltriethoxysilane (APTES) as a probe for protein labeling. We show that fluorescent SAND probes
exhibit both bright photoluminescence and biocompatibility in an aqueous environment. Selective in vitro
imaging using protein and carbohydrate labeling of hepatoma cell lines are demonstrated using biocompatible
SANDs conjugated with avidin and galactose, respectively. Cytotoxicity tests show that conjugated SAND
particles have negligible effects on cell proliferation. Unlike other synthetic systems that require multistep
treatments to achieve robust surface functionalization and to develop flexible bioconjugation strategies, our
results demonstrate the versatility of this one-step SAND fabrication method for creating multicolor fluorescent
probes with tailored functionalities, efficient emission, as well as excellent biocompatibility, requisite for broad
biological use.
KEYWORDS Atomic force microscopy, hydrophilicity, molecular imaging, self-assembly, silane
1. Introduction 1
Fluorescence imaging and biolabeling technologies 2
enable visualization of a vast array of biological 3
moieties in their native environment. Biomolecules, 4
proteins, and ions, for example, may be monitored 5
within cells, tissues, and organisms, in order to better 6
understand their function and role in biological 7
processes [1, 2]. 8
Organic dyes, such as FITC and fluorescent 9
proteins, have been used extensively in cell and 10
tissue bioimaging because they are biodegradable, 11
nontoxic, and target-specific. However, they are 12
vulnerable to chemical and metabolic degradation, 13
and rapid photobleaching, which limits their range 14
of use in bio-applications and biomolecule tracking 15
[3]. 16
Nano Res DOI (automatically inserted by the publisher)
Research Article
————————————
Address correspondence to Shuchen Hsieh, [email protected]
3
Fluorescent nanoparticles such as semiconductor 17
quantum dots (QDs) have been shown to exhibit 18
strong photoluminescence (high quantum yields), 19
multiple fluorescence, and excellent photostability 20
[4]. Thus, QDs have been proposed as promising 21
candidates for molecular labeling as well as in vitro 22
and in vivo imaging. Most importantly for cancer 23
screening, QDs could deliver tens to hundreds of 24
cancer biomarkers simultaneously, making them 25
suitable as a clinical diagnostic tool for the early 26
detection and classification of cancer [5]. 27
However, for most biolabeling and biomedical 28
applications, QDs must undergo secondary chemical 29
processing to modify the outer surface. This serves 30
the dual purpose of providing a mechanism for 31
adding specific chemical functionalities as well as 32
isolating the toxic QD core materials from the 33
environment. For example, it is often desirable to 34
convert hydrophobic characteristics to hydrophilic 35
(water-soluble) [6] so that different functional 36
molecules or biomolecules can be attached. Further, 37
their potential cytotoxicity, caused by the leaching of 38
heavy metal ions, remains a critical concern for in 39
vivo applications [7]. 40
In our previous work, Lin et al. [8] reported on the 41
one-pot self-assembled SiOx nanodots via hydrolysis 42
and condensation reaction using alkylakoxysilanes 43
as molecular precursors. These “self-assembled 44
nanodots (SANDs)” had the benefits of high 45
quantum yield, wide band absorption, multicolor 46
fluorescence, and very low toxicity. This combination 47
of properties were exploited in intravital imaging 48
experiments using zebrafish [8], demonstrating the 49
potential advantages for applications in biological, 50
biomedical, and clinical research. 51
Among the currently available materials from 52
alkylalkoxysilanes, 3-aminopropyltriethoxysilane 53
(APTES) is one an amine containing silane widely 54
used as a coupling agent for the surface modification 55
of silica or mica by self-assembly to enhance the 56
adhesion probability of biomolecules [9]. For 57
instance, Chai et al. [10] have developed an 58
immunosensor, which was fabricated by the 59
cross-linked bond of antibody with the 60
self-assembled monolayer (SAM) of an APTES 61
treated aluminum substrate, for the detection of ricin, 62
a lethal biological toxin. 63
Here, we demonstrate that SANDs from APTES 64
can be formed in aqueous solution and are well 65
suited for biolabeling applications. The APTES 66
derived SAND particles were further amine 67
functionalized, yielding a hydrophilic coating which 68
facilitates optical applications for biological systems. 69
This coating also provides for specific conjugation 70
with biomolecules [11]. Our results demonstrate that 71
the SANDs exhibit low cytotoxicity, are highly 72
photostable, and can be utilized for protein- and 73
carbohydrate-labeling via bioconjugation, producing 74
SAND-avidin and SAND-galactose probes, 75
respectively. Because of their biocompatibility and 76
amine-based labeling in aqueous solution, the 77
functionalized SANDs can be widely used as an 78
imaging probe, biomarker, or drug carrier for the 79
development of nanomedicine. 80
81
2. Experimental 82
2.1 SANDs synthesis 83
3-Aminopropyltriethoxysilane (APTES, >99%) was 84
purchased from (Sigma-Aldrich, USA), and used 85
without further purification. The molar 86
concentration of APTES was 4.3 M and calculated 87
from known densities and molecular weights (0.946 88
g/ml and 221.4 g/mol). To prepare the SAND 89
particles, 2 ml of APTES was placed into a 90
thoroughly cleaned glass vial and heated at a 91
constant temperature of 200°C while stirring (1100 92
rpm) on a hot-plate/stirrer (Suntex Inc. Co., Taiwan, 93
SH-301) for 4 h under an ambient “air” atmosphere 94
and then cooled with continued stirring to room 95
temperature. The SAND liquids were then used 96
directly to prepare sample for analysis by PL, XPS, 97
FTIR, and Raman. Aliquots of these samples were 98
diluted in toluene at a 1:200 V/V ratio to prepare 99
stock solutions for analysis by AFM and TEM. 100
101
2.2 Characterization of SANDs 102
Photoluminescence (PL) spectra measurements 103
were acquired using a HITACHI F-7000 104
Fluorescence Spectrometer. The excitation 105
4
wavelengths were 280 to 520 nm at 20 nm intervals 106
in the excitation domain. Emission spectra were 107
recorded from 200-800 nm. The UV–Vis absorption 108
spectrum was recorded with a UV–Vis 109
spectrophotometer (JASCO V-630, Japan) using a 110
1-cm path length quartz cell. In atomic force 111
microscope (AFM) imaging, 10 μl of SAND/toluene 112
solution was dropcast on a freshly-cleaved mica 113
substrate. An AFM (MFP-3D, Asylum Research, 114
USA), operating in intermittent contact mode under 115
ambient conditions, was used to visualize the 116
resulting surfaces to characterize the Au NP 117
distributions. A silicon cantilever (Olympus, 118
AC240TS) with a nominal spring constant of 2 Nm-1 119
was used for all images at a scan rate of 1.0 Hz and 120
image resolution of 512×512 pixels. Transmission 121
electron microscopy (TEM) images of SAND 122
samples were acquired using a JEOL JEM-2100 123
operated at 200 KV and at a vacuum of 2×10-5 Pa. 124
The TEM sample was prepared by depositing 0.5 μl 125
of SAND solution onto a TEM carbon grid with 126
copper 200 mesh (No. 01801, Ted Pella Inc, CA, USA) 127
and allowed to dry. Transmission Fourier transform 128
infrared (FTIR) spectra were acquired (8 cm-1 129
resolution 256 scans, at a sample compartment 130
vacuum pressure of 0.12 hPa) using a Bruker 66v/s 131
FTIR spectrometer. The sample was prepared by 132
depositing an aliquot of SAND solution to clean 133
double-side polished silicon (100) wafer substrates. 134
Spectra from a freshly plasma-cleaned silicon wafer 135
sample were collected before each measurement to 136
obtain the background spectrum. The X-ray 137
photoelectron spectroscopy (XPS) analysis sample 138
was prepared by placing 0.5 μl of the SAND 139
solution onto clean gold substrates at room 140
temperature. Elemental composition of the SAND 141
particles was determined by XPS using a JEOL JPS 142
9010 MX equipped with a monochromatic Mg Kα 143
X-ray radiation source. 144
145
2.3 Bicinchoninic acid (BCA) assays for the 146
determination of avidin 147
The measurements of protein concentrations by 148
BCA assay were performed with the Pierce® BCA 149
Protein Assay Kit (Thermo Fisher Scientific, 150
Germany). This assay estimates the formation of 151
Cu+ from Cu2+ by the Biuret complex in alkaline 152
solutions of protein (containing cysteine, cystine, 153
tryptophan and tyrosine or peptides containing 154
three or more amino acid residues) [12]. BCA reacts 155
with cuprous ion forming purple-colored reaction 156
product and measured at 562 nm in the UV–Vis 157
Spectrophotometer JASCO V-630. In this method, 10 158
μl of SAND-avidin solution (with different avidin 159
concentrations for 50, 25, 12.5, 6.5, 3.125, and 1.5 μg) 160
or 10 μl of SAND stock solution mixed with 190 μl 161
of deionized water were added to microtube. Then, 162
adding 0.8 ml of BCA in microtube at 65°C for 15 163
minutes and measuring at 562 nm in the UV/Vis 164
Spectrophotometer JASCO V-630. Blanks were 165
prepared by substituting SAND reacted with BCA. 166
167
2.4 The phenol-sulfuric acid spectrophotometric 168
method 169
The phenol-sulfuric acid spectrophotometric 170
method developed by Dubois [13] was adapted for 171
the determination of galactose binding on SAND. In 172
the method, 20 μl of SAND-galactose solution (with 173
different galactose concentrations for 2.5, 5, 10, 20, 174
40 and 80 mM) or 20 μl of SAND stock solution 175
mixed with 480 μl of 5% phenol solution were 176
added to glass vials, which were capped and 177
vortex-stirred. Then, using the concentrated sulfuric 178
acid (0.5 ml) to break down galactose (at room 179
temperature for 10 minutes) and to produce hexose 180
(6-carbon compounds) converting to 181
hydroxymethyl furfural. These compounds then 182
reacting with phenol to produce a yelloworange 183
color, which was measured at 490 nm in the UV/Vis 184
Spectrophotometer JASCO V-630. Blanks were 185
prepared by substituting SAND react with 186
phenol-sulfuric acid. The sulfuric acid (95-97%, ACS 187
reagent grade) was obtained from Scharlau 188
chemical and phenol (>98%) was purchased from 189
Shimakyu's Pure Chemical (Osaka, Japan). 190
191
2.5 Preparation of SAND solution and 192
functionalized SAND bioconjugates 193
We prepared a SAND solution by adding original 194
5
SAND liquid (1 ml) into DMEM medium (7 ml; 195
Thermo Scientific) and adjusted the pH value to 196
7.0-7.3 by 20% acetic acid (2 ml). For preparation of 197
avidin-SAND conjugates and galactose-SAND 198
conjugates, 1.0 mg of avidin (Sigma) and galactose 199
(Sigma) were added respectively into 100 μl of 200
SAND solution (4.3 M), mixed on a Vortex-Mixer 201
(Scientific Industries, Inc.), and then left to stand for 202
15 min at room temperature. Subsequently, 300 μl 203
of 5% acetic acid was added to these two mixtures 204
to adjust the pH to 7.0-7.3 to obtain the stock 205
avidin-SAND (1.07 M) and galactose-SAND (1.07 M) 206
conjugates, respectively. The biomolecules avidin or 207
galactose conjugated with SANDs through either 208
physical adsorption, hydrogen bonding, or 209
electrostatic interactions, between amine groups of 210
the SAND and carboxyl groups of avidin [14] or 211
hydroxyl group of galactose [15] in aqueous 212
solution. 213
214
2.6 Pull-down analysis and biotin competition 215
assay 216
For isolation of avidin-SAND conjugates, a mixture 217
of avidin (Sigma) and SAND solution was pH 218
adjusted to 7.0-7.3 by 20% acetic acid. The 219
avidin-SAND conjugates solution was then filtered, 220
using a cut-off filtration, centricon centrifugation 221
and wash, whereby unconjugated SANDs and 222
avidin-SAND conjugates were separated. The 223
bright avidin-SAND conjugates which remained on 224
the upper column were collected. UV images and 225
protein absorbance (2 µ l conjugates or avidin 226
protein) demonstrated that the unconjugated SAND 227
particles were removed from the avidin-SAND 228
conjugates solution. For a biotin competition assay, 229
avidin-SAND conjugates (100 µ l) were co-incubated 230
with 2-iminobiotin agarose (100 µ l) for 10 min at 231
4˚C. After a series of centrifugation and basic buffer 232
rinsing, excess avidin-SAND conjugates were 233
removed and the 2-iminobiotin agarose pull-down 234
avidin-SAND conjugates were collected. Protein 235
quantification (2 µ l pull-down beads conjugates) 236
were recorded and analyzed. SK-Hep-1 cells were 237
incubated with the 2-iminobiotin-avidin-SAND 238
complex (50 μg/ml) for 48 h, and the uptake of 239
avidin-SAND conjugates and cell morphology were 240
analyzed by fluorescence microscopy. 241
242
2.7 Cell culture and in vitro cytotoxicity assay 243
SK-Hep-1 cells, HepG2 cells and HeLa cells were 244
maintained in DMEM medium containing 10% calf 245
serum (Hyclone) and supplemented with 100 IU/ml 246
penicillin and 100 g/ml streptomycin (Invitrogen) 247
in 5% CO2 at 37°C incubator temperature. For the 248
cytotoxicity assay, SK-Hep-1 cells were harvested 249
and seeded in a 96 well culture plate (1×104 cells/100 250
μl/well) and kept overnight at 37°C and 5% CO2 in a 251
humidified incubator. Cells were then treated with 252
various concentrations (0, 0.25, 1, 2, 5, 10, and 20 253
mM) of SAND for 48 h. Subsequently, the surviving 254
cells were analyzed by MTT 255
(3-(4,5-dimethy-thiazol-2-yl)-2,5-diphenyl 256
tetrazolium bromide) assay. 257
258
2.8 Fluorescence microscopy 259
SK-Hep-1 cells, HepG2 cells, and HeLa cells (4×104) 260
were seeded onto coverslips in 12-well culture plates 261
and were cultured at 37°C overnight. For protein 262
biolabling experiments, SK-Hep-1 cells were treated 263
with the SAND (20 mM), avidin (50 μg/ml), 264
avidin-FITC (50 μg/ml; Sigma) and avidin-SAND (20 265
mM) for 48 h. For saccharide sugar biolabling 266
experiments, HepG2 cells and HeLa cells were 267
treated with galactose-SAND conjugates (80 mM) for 268
2 h. Subsequently, these cell samples were washed 269
with PBS, fixed with 4% paraformaldehyde, and 270
perfused with 0.2% Triton X-100 for the inverted 271
fluorescence microscope (Leica, DMIL, Germany), 272
though the filters: DAPI, FITC and R-DIL. Image 273
processing and projections were performed using 274
Photoshop 5.5 (Adobe Systems, Mountain View, CA). 275
276
2.9 Flow cytometry 277
SK-Hep-1 cells (3×105) were seeded into 6 cm 278
culture plates overnight and then treated with 279
different concentrations (0, 2, 5, 10, and 20 mM) of 280
SAND for 48 h at 37°C. The cells were harvested, 281
washed twice with PBS, and fixed with 4% 282
paraformaldehyde, and perfused with 0.2% Triton 283
6
X-100. Cells were suspended in the PBS and the 284
SAND uptake was analyzed by detection of the 285
fluorescence intensity in single-cell using flow 286
cytometry analysis (Beckman Coulter) with a 488 287
nm laser. At least 10,000 cells were counted. The 288
data is presented as mean fluorescence index (MFI) 289
and calculated as the ratio of the mean fluorescence 290
intensity of the sample and the control. Green and 291
red fluorescence were monitored using FITC and 292
PE channels, respectively. 293
294
3. Results and discussion 295
3.1 Structural and optical analysis to characterize 296
SANDs 297
Nanoparticles (NPs) such as inorganic 298
nanoparticles, quantum dots (QDs), magnetic NPs 299
(MPs), and multifunctional NPs, often have a 300
hydrophobic surfactant coating that limits their use 301
in biomedical applications. Therefore, methods to 302
functionalize NPs to increase water solubility prior 303
to use are desired [16, 17]. 304
The focus of this work is the fabrication and 305
characterization of a new type of SAND particle 306
derived from 3-aminopropyltriethoxysilanes 307
(APTES). The APTES organofunctional alkoxysilane 308
molecules self-assemble via attack and 309
displacement of the alkoxy groups, (usually the 310
ethoxy (–OCH2CH3) groups), forming a covalent 311
Si–O–Si bond [18, 19]. During SAND formation, the 312
hydrophobic alkoxysilane backbones are most 313
likely embedded inside the SAND particles while 314
the hydrophilic amine groups extend outside into 315
the aqueous environment. Therefore, APTES 316
SANDs may be highly water soluble and thus ideal 317
for a variety of biomedical applications. 318
Additionally, the terminal amine groups at the 319
outer surface of the SANDs would have a positive 320
surface potential [20] and could easily be 321
covalently-linked to other interesting chemicals or 322
biomolecules using established protocols. 323
Structural characterization of the SAND particles 324
was carried out by AFM and TEM. Fig. 1(a) shows 325
AFM topographical images of SANDs deposited on 326
a clean mica substrate. 327
Figure 1 Structural Characteristics of SAND particles. (a) AFM 328 topographic image of SANDs deposited on mica (1×1 μm2 329 scan). The actual AFM tip apex radius of curvature was on the 330 order of ~10-25 nm, so the apparent width of the particles is 331 larger due to the tip broadening effect. The measured height of 332 the particles, was used to determine the size, because the AFM 333 height measurement is not subject to this lateral tip-broadening 334 effect [8]. (b) Particle size distribution histogram of APTES 335 deposited on mica from (a). (c) TEM image with the lattice 336 fringes from HRTEM of the SAND particles. (d) PL spectra of 337 SAND solution recorded for progressively longer excitation 338 wavelengths from 280 to 520 nm in 20 nm increments. The 339 inset photographs show SAND solutions irradiated by visible 340 light and by an ultraviolet lamp (365 nm), respectively. 341
The SAND particles are spherical, monodisperse, 342
and well-separated on the mica surface. The 343
corresponding particle size histograms in Fig. 1(b) 344
show that the SAND particles have a narrow size 345
distribution with an average NP diameter of 2.5 ± 346
0.5 nm as determined from statistical analysis of 178 347
particles. The SAND particle size distribution was 348
further confirmed to be 2.4 ± 0.6 nm by the 349
high-resolution TEM image in Fig. 1(c). These 350
results show a homogeneous distribution and are 351
consistent with AFM results. The inset in Fig. 1(c) 352
shows characteristic lattice fringes of the SAND 353
particles measured at ~0.21 nm. Although we do not 354
know the exact structure of the carbon around the 355
SiOx core, this value is comparable with the (110) 356
planes of graphene [21]. 357
Surface chemistry and elemental composition of 358
the SAND particles was characterized by FTIR, 359
Raman and XPS. In FTIR spectra (Fig. S-1 in the 360
Electronic Supplementary Material (ESM)) we 361
observed peaks at 2935 and 2885 cm-1 that are 362
assigned to the symmetric and anti-symmetric 363
7
–CH2– stretch, and indicate the presence of 364
anchored propyl groups. The vibrational mode 365
peaks at 1570 and 1485 cm-1 arise primarily from 366
surface amine groups [22]. Two peaks attributed to 367
the C–N stretching vibration were observed at 1125 368
and 1041 cm-1 [23]. Raman spectra were acquired 369
(Fig. S-2 in the ESM) with peaks at 2892 cm-1, 2936 370
cm-1 and 2980 cm-1, corresponding to the ν(CH2)sym, 371
ν(CH2)asym, and ν(CH3) vibrational modes. The 372
presence of these peaks confirm that alkyl groups 373
were still present in the SAND particles [24]. 374
Elemental analysis by XPS (Fig. S-3 in the ESM) 375
showed that SANDs were mainly composed of Si, O, 376
C, and N. The Si 2p peaks at 99.4 and 103.4 eV arise 377
from silicon and SiO2, respectively. The O 1s 378
spectrum at 533.2 eV indicates a Si–O–Si bonding 379
environment (Fig. S-3(a-b) in the ESM) [25]. The C 380
1s spectrum had one peak at 286.0 eV which was 381
attributed to C–N bonds [26], and the N 1s 382
spectrum showed a binding energy peak at 400.2 eV, 383
which was attributed to NH2 groups [19] (Fig. 384
S-3(c-d) in the ESM). These results confirm that bare 385
amine groups were present in the SAND and that 386
all expected vibrational modes of the coupled 387
ligand shell were present. Our elemental analysis 388
revealed that the SANDs were composed of SiO2 389
clusters with amine-alkyl groups. The sensitivity 390
factors for Si, O, C, and N, are 0.654, 2.29, 1, and 391
1.61, respectively [27]. The integrated peak areas 392
normalized with the sensitivity factors for the Si 2p : 393
O 1s : C 1s : N 1s ratios are 2.1 : 3.3 : 6.0 : 1.0. 394
Multicolor imaging is ideal for simultaneous 395
evaluation of multiple biomarkers within a complex 396
tissue for medical applications. Since cancer and 397
many diseases involve a large number of genes and 398
proteins, multicolor imaging could provide an 399
efficient and quantitative way to distinguish each 400
biomarker accurately. Nanoparticles exhibit bright 401
and multicolor photoluminescence [28], and are 402
thus an important tool for biomedical applications 403
such as biolabeling, biosensing and bioimaging [29, 404
30]. To characterize the optical properties of the 405
SAND particles, we performed absorption and 406
emission measurements. As demonstrated in Fig. 407
S-4 (in the ESM), the as-obtained SANDs show a 408
clear absorption feature centered at 365 nm which is 409
comparable to those previously reported for 410
fluorescent CDs [31, 32]. The PL spectra (Fig. 1(d)) 411
revealed that SANDs exhibit wideband and 412
multicolor fluorescence emission, the intensity of 413
which depends on the excitation wavelength. The 414
fluorescence emission spectra at several excitation 415
wavelengths varied from 280 to 520 nm. The PL 416
peak exhibited a fluorescence emission maximum 417
of λem = 502 nm at an excitation wavelength of 440 418
nm, above which the PL peak shifted to longer 419
wavelengths and its intensity decreased rapidly. 420
Further, it exhibited strong bright blue fluorescence 421
under excitation by an ultraviolet lamp at 365 nm 422
(Fig. 1(d) inset). 423
The quantum yield (QY) was measured using the 424
F-7000 fluorescence spectrophotometer combined 425
with a Hitachi 5J0-0148, 60-mm integrating sphere 426
with a coating of aluminum oxide (Hitachi Inc., 427
Japan). The QY was calculated by the built-in 428
quantum yield measurement system in the Hitachi 429
software using direct and indirect excitation. Direct 430
excitation is when the sample is directly facing the 431
excitation beam. Indirect excitation uses an 432
integrating sphere, where a portion of the emitted 433
photons from the sample are reflected from within 434
the integrating sphere and lead to further sample 435
excitation. The QY was obtained from direct and 436
indirect excitation, and the final QY values were 437
corrected by the following equation: 438
439
ϕ_QY is the corrected SAND quantum yield. ϕ_P1 440
is the internal QY using direct excitation, abs_P1 is 441
absorptance using direct excitation, and ϕ_P2 is the 442
internal QY using indirect excitation. The 443
measurement of QY for the progressive excitation 444
wavelengths from 320 to 520 nm in 20 nm 445
increments is shown in Table S-1 (in the ESM). The 446
SANDs had a maximum quantum yield of 29.5% at 447
an excitation wavelength of 440 nm, which may be 448
compared to the measured QY values for 449
silica-coated carbon dots (11-14.7%) [33], 450
laser-ablated graphite (4-10%) [34], and the thermal 451
oxidation of carbon dots (<5%) [35]. 452
8
Figure 2 Dose-dependent cellular toxicity of SANDs to 453 SK-Hep-1 cells. Relative cell viabilities after 48 h treatment 454 with SANDs from MTT assay were plotted against SAND 455 concentration. No obvious toxicity was observed. Values are 456 expressed as mean ± S.D. of three individual experiments. 457
458
This result shows that SANDs have a high quantum 459
yield compared to the other types of NPs 460
mentioned above. 461
462
3.2 The biocompatibility and multifluorescence 463
of SANDs 464
To validate the performance of SANDs as 465
cell-imaging agents, cytotoxicity was evaluated 466
using human hepatoma SK-Hep-1 cells through the 467
MTT assay (Fig. 2). SK-Hep-1 cells were cultured in 468
a medium containing different concentrations of 469
SANDs (0, 0.25, 1, 2, 5, 10, and 20 mM) for 48 h and 470
resulted in cell viability values of 100 ± 2.3, 95 ± 3.24, 471
97 ± 3.76, 94 ± 3.11, 96 ± 3.18, 99 ± 1.56, and 98 ± 3.07 472
(% of control) respectively. This result indicated that 473
the SANDs had no significant cytotoxic effect on 474
SK-Hep-1 cells, and thus may be a good candidate 475
for in vitro labeling or cellular imaging. 476
We further studied the uptake efficiency and 477
multicolor fluorescence of SANDs by flow 478
cytometry (using FITC and PE filters to detect the 479
green and red fluorescence, respectively), because 480
cell fluorescence intensity correlates with the 481
number of internalized nanoparticles. In flow 482
cytometry dot plots, each dot represents an 483
individual cell. Cells appearing in the lower left 484
quadrant (I) exhibited no fluorescence. Cells 485
appearing in the upper left (II) and lower right 486
quadrant (IV) correspond to cells emitting either 487
red or green fluorescence, respectively. And finally, 488
cells in the upper right quadrant (III) are cells 489
emitting both green and red fluorescence. 490
After cells were treated with various 491
concentrations of SAND (0, 2, 5, 10, and 20 mM; Fig. 492
3(a-e)) for 48 h, the cells emitting both red and 493
green fluorescence were detected using two 494
fluorescent filters: FITC and PE, and revealed an 495
increasing SAND uptake efficiency (Fig. 3(a-e), 496
quadrant III). These results are shown in the 497
histogram plot (Fig. 3(f)) and indicate that ~19% of 498
detected cells took up SANDs at lower dose (2 mM) 499
and more than 95% of cells were observed with 500
high SANDs uptake at higher dose (20 mM). 501
Further, the separate detection of cells exhibiting 502
red (Fig. 3(a-e), quadrant II and quadrant III) or 503
green fluorescence (Fig. 3(a-e), quadrant III and 504
quadrant IV) were also analyzed, and histograms 505
plots (Figs. S-5 and S-6 in the ESM) showed that 506
these cells were elevated in a dose-dependent 507
manner. Conversely, the control cells, which 508
exhibited no fluorescence, were decreasing (Fig. 509
3(a-e), lower left quadrant I). 510
Figure 3 Fluorescence intensity monitored by flow cytometry. 511 SK-Hep-1 cells were treated with various concentrations of 512 SAND for 48 h. (a-e) Flow cytometry dot plot (Fluorescence of 513 x-axis and y-axis is FITC and PE, respectively). The number of 514 dots in each quadrant represents the fluorescence intensity of 515 the cells. The lower left-hand (I), upper left-hand (II), lower 516 right-hand (IV), and upper right-hand (III) quadrants indicate 517 non-fluorescent, red fluorescence, green fluorescence, and both 518 green and red fluorescence, respectively. (f) Bar graph 519 representation of the fluorescence intensity of cells from the dot 520 plot at various concentrations of SAND. Increased cell 521 fluorescence due to accumulation of fluorescent nanoparticles is 522 presented as mean fluorescence index (MFI). The results were 523 obtained from three separate experiments and the bar represents 524 mean ± S.D. of three individual experiments. *p < 0.01. 525
9
Here, auto-fluorescence (2-4%) was occurring in 526
the control cells (Fig. 3(a), quadrant III and IV), but 527
it did not interfere with the multi-fluorescence 528
emission of SAND in SK-Hep-1 cells. This result 529
demonstrated the excellent biocompatible nature 530
and multi-fluorescence potential of SANDs. 531
Next, we explored in vitro internalization of 532
SANDs in SK-Hep-1 cells. Cells were cultured and 533
maintained in DMEM medium containing 20 mM of 534
SANDs for 48 h, with a detected concentration of 535
95% cellular SAND-internalization efficiency. The 536
cell fluorescence images (Fig. S-7 in the ESM) shows 537
bright field (a), blue (b), green (c), and red (d) 538
emission, respectively under fluorescence 539
microscopy. The multi-photoluminescence images 540
of SK-Hep-1 cells revealed that SANDs were 541
internalized and accumulated into the cytoplasm 542
region and that they surrounded, but did not enter 543
into the nucleus (Fig. S-7 in the ESM). Endocytosis 544
is the likely internalization mechanism of SANDs 545
from outside the cell membrane into the cytoplasm. 546
The cell membrane is a negatively charged 547
phospholipid bilayer which separates the 548
intracellular substance and organelles from the 549
extracellular space. SANDs have abundant surface 550
functional amine groups, which imparts to them 551
more cationic characteristics. They thereby 552
interacted electrostatically with the anionic cell 553
membrane and were further taken up through an 554
adsorptive endocytosis mechanism and internalized 555
by cells [36-38]. More detailed internalization 556
mechanisms of SANDs will require additional 557
experiments. 558
559
3.3 Biomolecule labeling and imaging with 560
SANDs 561
Luminescent NPs are very attractive because of 562
their potential for applications in biolabeling, 563
bioimaging [6] and biomedicine [4]. However, these 564
hydrophobic core NPs frequently needs complex 565
grafting to provide the necessary surface chemistry 566
[6]. Our SANDs are derived from APTES, which is 567
commonly used for surface functionalization to 568
prepare amine-terminated (–NH2) silica-based 569
materials, or to convert nanoparticles from 570
hydrophobic to hydrophilic character [39, 40]. The 571
presence of amine groups on the surface of targets 572
is important for biological applications, such as 573
biolabeling [20] and clinical immunosensor [41], as 574
it allows proteins or other biomolecules to be 575
attached in a simple manner [14, 42]. Thereby, 576
SANDs prepared using APTES as the precursor 577
possess the characteristics of APTES and can be 578
effective tools in the biological sciences for such 579
applications as biolabeling, cell imaging, biosensors, 580
and biomimetic membranes. 581
Before the cell-binding and cell-labeling 582
experiments, we used bioconjugate techniques for 583
the determination of whether a particular protein or 584
sugar is located on the SANDs. The bicinchoninic 585
acid protein assay was performed to measure the 586
total avidin content on SANDs (Fig. S-8 in the ESM), 587
and the phenol-sulfuric acid spectrophotometric 588
method was performed to monitor the 589
incorporation of galactose on SANDs (Fig. S-9 in the 590
ESM). Repeatability and reproducibility were 591
evaluated from the results obtained by two analysts 592
working on the same sample, which exhibit a linear 593
relationship between the absorbance and the 594
varying concentrations. 595
Avidin is a tetramic protein [43] and is bound to 596
biotin with high affinity and specificity [44]. It is 597
used in many biochemical assays, including 598
western blot, ELISA, and pull-down assays. To 599
demonstrate the feasibility of SANDs as a protein 600
labeling probe, we conjugated SANDs with avidin 601
(Fig. S-10 and S-11 in the ESM). To validate the 602
conjugated SAND interaction with biotin, a pull 603
down assay was executed using a commercial 604
2-iminobiotin agarose (Fig. S-12 in the ESM). Our 605
data showed that SANDs are successfully 606
conjugated to avidin and implied that avidin-SAND 607
conjugates could be taken up via a biomolecule 608
interaction-mediated uptake within the cell. We 609
chose SK-Hep-1 cells, which are a good source of 610
biotin [45, 46], for the cell targeting experiment. 611
SK-Hep-1 cells were cultured in a medium 612
containing SANDs, avidin, avidin-FITC, and 613
SAND-avidin conjugates for 48 h, and the particle 614
uptake efficiency was then analyzed using 615
10
fluorescence microscopy. Fig. 4 shows that the cells 616
internalized the avidin-SAND conjugates over the 617
SANDs and avidin-FITC, thus demonstrating their 618
potential for protein labeling. 619
The SAND uptake was monitored using bright 620
field (Fig. 4(a)) and DAPI, FITC, and R-DIL filters 621
under fluorescence microscopy. SANDs were 622
internalized in the cytoplasm and surrounded the 623
cell nucleus, as shown in the blue, green and red 624
fluorescence data shown in Fig. 4(b-d). This result is 625
consistent with the data of Fig. S-7 in the ESM. Cells 626
that were incubated with avidin (Fig. 4(e)), as a 627
fluorescence negative control exhibited no 628
fluorescence signal (Fig. 4 (f-h)). 629
SK-Hep-1 cells treated with the avidin-FITC (Fig. 630
4(i)), which is a well-known organic dye and used 631
as a fluorescence positive control, showed bright 632
green fluorescence. Here, avidin-FITC internalized 633
in the cytoplasm (Fig. 4(k)), but the fluorescence 634
quickly decayed (Fig. S-13(d-f) in the ESM) using 635
excitation comparable to the SAND treated group 636
(Fig. S-13(a-c) in the ESM). 637
Figure 4 Comparison of the uptake of SANDs, avidin, 638 avidin-FITC conjugates, and SAND-avidin conjugates by 639 human hepatoma cell line. SK-Hep-1 cells were treated with 640 SAND (20 mM) (a-d), avidin (50 μg/ml) (e-h), avidin-FITC (50 641 μg/ml) (i-l) and SAND-avidin conjugates (20 mM) (m-p), 642 respectively for 48 h. Avidin was used as a fluorescent negative 643 control and avidin-FITC as a fluorescent internalized positive 644 control. The uptake of nanodots and cell morphology were 645 analyzed by fluorescence microscopy. The surface NH2 groups 646 of the SAND particles interacted with avidin and thus the 647 avidin-SAND conjugates were more internalized into the 648 cytoplasm than SANDs or avidin-FITC treated cells. The 649 cellular multi-photoluminescence images were monitored using 650 DAPI, FITC and R-DIL channels, respectively. Original 651 magnification was 400X. 652
Not surprisingly, avidin-FITC treatment showed 653
no blue or red signals (Fig. 4(j-l)). These results 654
show that SANDs were more photostable than the 655
organic dye-FITC, which suffered from significant 656
photobleaching. Further, we tested the SAND 657
solution stability at different pH values (0, 2, 4, 6, 8, 658
10, 12, 14) and in phosphate buffered saline (PBS) 659
and sodium chloride (NaCl) to mimic various 660
normal and extreme physiological conditions 661
(Figure S-14 and S-15 in the ESM). The PL results 662
from these experiments show that the SANDs were 663
stable under a variety of harsh solution conditions 664
(Table S-2 in the ESM). 665
Finally, we treated cells with avidin-SAND 666
conjugates (Fig. 4(n-p)) and the results revealed not 667
only the characteristic multi-color fluorescence 668
signals consistent with the SAND treated group 669
(Fig. 4(b-d)), but also showed agglomerated 670
fluorescent particles unlike in the SANDs and 671
avidin-FITC treated groups (Fig. 4(k)). In addition, 672
the agglomerated avidin-SAND conjugates showed 673
good distribution in the cytoplasm and around the 674
nucleus. The avidin-SAND conjugates (Fig. 4(n-p)) 675
were more accessible and accumulated in the cells 676
more efficiently than SAND (Fig. 4(a-d)) or 677
avidin-FITC (Fig. 4(k)) during an equivalent 678
incubation time. Moreover, a competition assay was 679
also executed. After avidin-SANDs conjugates were 680
pre-precipitated by 2-iminobiotin agarose and then 681
treated on SK-Hep-1 cells for 48 h. The cells 682
revealed that no fluorescence could be observed 683
(Fig. S-16 in the ESM) when compared to 684
avidin-SANDs conjugates and SANDs groups. This 685
result suggests that exogenous biotin interacts with 686
avidin-SAND conjugates and further interfered 687
with the internalization of avidin-SAND conjugates. 688
The bright fluorescence from avidin-SAND 689
conjugates revealed the high degree of avidin-biotin 690
interaction in SK-Hep-1 cells. Although the 691
avidin-FITC group was expected to exhibit a high 692
degree of avidin-biotin interaction, as in the 693
avidin-SAND conjugate group, both SANDs and 694
the avidin-SAND conjugates exhibited much more 695
stable fluorescence than the avidin-FITC group 696
when cells were exposed to continuous excitation 697
11
(Fig. S-13 in the ESM). This indicates that SANDs 698
are more photostable and photobleaching-resistant 699
than avidin-FITC. We note also that the rapid 700
photobleaching of organic dyes has been reported 701
in the literature and is a significant limitation on 702
their use in bio-applications [3]. 703
Another example demonstrates targeted cell 704
imaging through SANDs conjugated with 705
carbohydrate-galactose to examine their feasibility 706
as a biomolecule probe. Carbohydrate 707
functionalized nanoparticles have been successfully 708
administered as nanoprobes for cellular targeting or 709
bioimaging applications and drug delivery in vivo 710
and in vitro [47, 48]. For example, an anti-cancer 711
drug-paclitaxel was loaded into a galactose 712
conjugated nanoparticle (NP-Gal-PTX) and 713
significantly inhibited HepG2 cell viability through 714
the interaction between galactose and 715
asialoglycoprotein receptor (ASGP-R) [47]. Here, 716
galactose-SAND conjugates were formed through 717
hydrogen bonding between the hydroxyl end of the 718
galactose and the primary amine group of the 719
nanoparticles [49]. Two cell lines, human 720
hepatocellular carcinoma cells (HepG2) and human 721
cervical cancer cell line (HeLa) were specifically 722
chosen for cell targeting experiments because 723
HepG2 cells express ASGP-R on the cell surface (the 724
galactose binding protein), whereas HeLa cells 725
without ASGP-R expression were chosen as the 726
negative control [50] (Fig. 5). 727
Both HepG2 and HeLa cells were incubated in 728
medium containing galactose-SAND conjugates (80 729
mM) for 2 h and then the cell targeting efficiency 730
was analyzed using fluorescence microscopy. Fig. 731
5(a) shows a healthy HepG2 cell without survival 732
stress from the higher concentration SAND 733
treatment. The strong signals of blue, green, and red 734
multicolor fluorescence demonstrate that the 735
galactose-SAND conjugate interacts strongly with 736
ASGP-R-positive HepG2 cells (Fig. 5(b-d)). Our 737
galactose-SAND conjugate can specifically target 738
the cell surface and then is internalized by HepG2 739
cells through the receptor-mediated endocytosis 740
process [51]. 741
742
Figure 5 The receptor mediated uptake of SAND-galactose 743 conjugates in hepatoma cell line-HepG2 cells. Images of 744 HepG2 cells (a-d) and HeLa cells (e-h) cells treated with 745 SAND-galactose conjugates (80 mM) for 2 h and their 746 corresponding optical images. The SAND-galactose conjugates 747 exhibited specific binding and uptake by HepG2 cells but not 748 by HeLa cells. The cellular multi-photoluminescence images 749 were monitored using DAPI, FITC and R-DIL channels, 750 respectively. Original magnification was 400X. 751
752
On the other hand, the ASGP-R-negative HeLa cells 753
(Fig. 5(e)) revealed that the high dose of SAND had 754
an insignificant effect on cell survival. As expected, 755
the negative ASGP-R expressed-HeLa cells showed 756
very weak fluorescence (Fig. 5(f-h)). This 757
observation demonstrates that functionalized 758
SANDs can be modified with specific ligands for 759
targeting biomolecule detection, such as 760
carbohydrates, and have potential for further 761
biomedical and biosensing applications. 762
In this present study, we have demonstrated that 763
bright and substrate-independent 764
functionalized-SANDs are an effective labeling 765
probe for proteins and carbohydrates. Further, 766
peptides, drugs, or biomolecules (i.e. DNA) can be 767
directly modified or conjugated with the APTES 768
derived SANDs via the amine-terminated (–NH2) 769
functional groups. This study demonstrates a new 770
and novel application for alkoxysilane 771
nanoparticles in bioimaging and biolabeling. 772
773
12
4. Conclusion 774
In summary, our study offers a novel systematic 775
study of a new nanomaterial for bioimaging and 776
biolabeling. The APTES-derived SANDs have 777
several advantages including 778
substrate-independent functionalization, high water 779
solubility, biocompatibility, negligible cytotoxicity, 780
strong photoluminescence, and high photostability. 781
Moreover, we demonstrate the feasibility of SANDs 782
as a biomolecular labeling probe through direct 783
functionalization to obtain surface coatings for 784
specific biomolecular conjugates without secondary 785
chemical multi-modification or difficult 786
functionality processing. Hence, this new type of 787
nanodot has strong potential for the development of 788
in vitro biochemical assays and could be used 789
advantageously for in vivo imaging, disease 790
detection, and cancer diagnosis. 791
792
Acknowledgements 793
M. L. Kung and P. Y. Lin contributed equally to this 794
work. The authors would like to thank the Ministry 795
of Science and Technology (NSC 796
101-2113-M-110-013-MY3) and (NSC 797
101-2811-M-110-034 and NSC 102-2811-M-110-008) of 798
Taiwan and the National Sun Yat-sen University 799
Center for Nanoscience and Nanotechnology for 800
financial support of this work. Prof. Hsieh also 801
thanks Dr. Ming-Hong Tai for providing the 802
fluorescent microscope and SK-Hep-1 cell line, and 803
Dr. David Beck for helpful discussions. 804
805
Electronic Supplementary Material: Supplementary 806
material (Additional SANDs characterization data 807
including FTIR, Raman, and XPS, as well as 808
biolabeling and bioimaging applications of SANDs.) 809
is available in the online version of this article at 810
http://dx.doi.org/10.1007/s12274-***-****-* 811
812
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S1
Electronic Supplementary Material
Correct Aqueous Self-Assembly and Surface-Functionalized Nanodots for Live Cell Imaging and Labeling
Mei-Lang Kung,1 Pei-Ying Lin,
1 Chiung-Wen Hsieh,
1 and Shuchen Hsieh
1,2 ()
1Department of Chemistry and Center for Nanoscience and Nanotechnology, “National Sun Yat-sen University”, Kaohsiung, 80424 2School of Pharmacy, College of Pharmacy, Kaohsiung Medical University, Kaohsiung, 80707
Supporting information to DOI 10.1007/s12274-****-****-* (automatically inserted by the publisher)
S2
Supplementary Figures
Figure S1 FTIR spectra of SAND particles deposited on a silicon substrate. The presence of the anchored propyl
group was confirmed by the presence of symmetric and anti-symmetric –CH2– stretching peaks at 2935 and 2885
cm-1
. The major contribution of the vibrational modes at 1570 and 1485 cm-1
arises from surface amine groups.
Peaks corresponding to C–N stretching vibrations were observed at 1125 and 1041 cm-1
.
S3
Figure S2 Raman spectra of SAND particles deposited on a silicon substrate. The peak at 669 cm-1
and 961 cm-1
correspond to a Si–O stretch. The peak at 2892 cm-1
, 2936 cm-1
and 2980 cm-1
correspond to the ν(CH2)sym,
ν(CH2)asym, and ν(CH3) stretching peaks, indicating the presence of alkyl groups.
S4
Figure S3 Near-surface elemental analysis of the SAND using XPS. XPS data of SANDs deposited on gold
substrate, showing (a) Si 2p, (b) O 1s, (c) C 1s, and (d) N 1s binding energy peaks. The Si 2p peak at 103.4 eV
corresponds to SiO2, and the lower binding energy peak at 99.4 eV indicates pure silicon. Figure (b) shows the O
1s spectra at 533.2 eV which indicates a Si–O–Si bonding environment. Figure (c) shows the C 1s spectrum with a
binding energy peak at 286.0 eV. This corresponds well to C–N carbon and confirms that the alkyl chains from the
precursor are preserved in the NP product. Figure (d) shows the N 1s spectra with a peak at 400.2 eV which is
attributed to NH2 groups.
S5
Figure S4 UV-Vis absorption spectra of SANDs.
S6
Table S1 Summary for the QY for SAND particle solution at different emission wavelengths.
S7
Figure S5 Red fluorescence monitored by flow cytometry. SK-Hep-1 cells were treated with different
concentrations of SANDs for 48 h. (a-e) In these histogram plots, grey filled regions indicate control normal cells;
red unfilled regions indicate SAND-treated cells. (f) Bar graph representation of the red fluorescence intensity at
various concentrations of SANDs. *p < 0.01.
S8
Figure S6 Green fluorescence monitored by flow cytometry. SK-Hep-1 cells were treated with different
concentrations of SANDs for 48 h. (a-e) In these histogram plots, grey filled regions indicate control normal cells;
green unfilled regions indicate SAND-treated cells. (f) Bar graph representation of the green fluorescence intensity
at various concentrations of SANDs. *p < 0.01.
S9
Figure S7 Uptake of SANDs nanodots by SK-Hep-1 cells. SK-Hep-1 cells were treated with SANDs (20 mM) for
48 h. The cells were fixed and then imaged at bright field (a). The blue, green, and red fluorescence (b-d) from
SAND were recorded using DAPI, FITC and R-DIL filters, respectively. Multicolor fluorescent emission as a
function of excitation wavelength was observed.
S10
Figure S8 The concentrations of avidin conjugation with SAND were determined by bicinchoninic acid-based
protein assay. Each experiment was repeated for three times. (Cavidin: avidin conjugated with SAND at different
concentrations)
S11
Figure S9 The contents of galactose binding to SAND were analyzed by the phenol-sulfuric acid method. Each
experiment was repeated for three times. (Cgalactose: galactose binding to SAND at different concentrations)
S12
Figure S10 Isolation of Avidin-SAND conjugates. Mixture of avidin (Sigma) and SAND solution was pH adjusted
to 7.0-7.3 using 20% acetic acid. After the avidin-SAND conjugate solution was filtered, centrifuged, and rinsed,
unconjugated SANDs and avidin-SAND conjugates were separated. The bright avidin-SAND conjugates which
remain on the upper column were collected. (a) The UV image and (b) protein absorbance (2 µl conjugates or
avidin protein) demonstrated that the unconjugated SAND particles were removed from avidin-SAND the
conjugate solution.
S13
Figure S11 FTIR spectra of SAND, avidin, and conjugated SAND-avidin solutions.
S14
Figure S12 A biotin pull-down experiment using biotin agarose to precipitate Avidin-SAND conjugates.
Avidin-SAND conjugates (100 µl) were co-incubated with 2-iminobiotin agarose (100 µl) for 10 min at 4˚C. After
a series of centrifugation and basic buffer rinsing, excess avidin-SAND conjugates were removed and the
2-iminobiotin agarose pull-down avidin-SAND conjugates were collected. (a) The UV image and (b) protein
absorbance (2 µl pull-down beads conjugates) were recorded and analyzed.
S15
Figure S13 SAND photostability compared to FITC. Consecutive images of SAND (a-c), avidin-FITC (d-f) and
avidin-SAND (g-i) expressed in SK-Hep-1 cells. These images represent the SAND, avidin-FITC and
avidin-SAND fluorescence intensity respectively through a 5 min period during which three fluorophores were
exposed to continuous excitation.
S16
Figure S14 Photographs of SAND solutions under different pH conditions illuminated with ambient light (a) and
365 nm UV radiation (b).
S17
Figure S15 Photographs of SAND solutions in two kinds of salt solutions under ambient light (a) and under 365
nm UV illumination (b).
S18
Table S2 Summary of SAND emission wavelengths under different pH conditions and under different salts
environments measured in the 280-520 nm range.
S19
Figure S16 Biotin competition assay. After the 2-iminobiotin agarose pull-down avidin-SAND
conjugates were collected, we incubated SK-Hep-1 cells with SAND (a-d), avidin-SAND (e-h) and
the 2-iminobiotin-avidin-SAND complex (50 μg/ml) (i-l) for 48 h. The nanoparticle uptake and cell
morphology were analyzed by fluorescence microscopy. SANDs were taken up and surrounded the
cellular nucleus. The avidin-SAND conjugates were much more internalized into the cytoplasm than
SANDs. Moreover, following the 2-iminobiotin-avidin-SAND complex treatment, cells exhibited
no fluorescence. This result shows that exogenous biotin interfered with avidin-SAND conjugate
cellular uptake, and supports our assertion that the avidin and SAND were conjugated. The cellular
multi-photoluminescence images were monitored using DAPI, FITC and R-DIL channels,
respectively. Original magnification was 400X.
