View Cart     Email Updates

Updated: July 1, 2005

N A N O P R O B E S     E - N E W S

Vol. 6, No. 7          July 1, 2005

In this Issue:

This monthly newsletter is to inform you about techniques to improve your immunogold labeling, highlight interesting articles and novel applications of metal nanoparticles, and answer your questions. We hope you enjoy it and find it useful; as always, let us know if we can improve anything.

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Double Labeling: Nanogold®, 10 nm Gold, and Gold Enhancement

In our previous reports on double labeling, we highlighted procedures in which one target was labeled with Nanogold® and silver enhanced, then a second target was labeled with colloidal gold which would then be distinguished by its smaller size compared with the silver-enhanced gold.

Brainard and group now report an alternative approach, in which two different targets were labeled with Nanogold and with a 10 nm colloidal gold probe respectively, then gold enhanced, and used this approach in combination with coimmunoprecipitation, immunoblotting and immunocytochemistry to study the interactions of cell signaling components in human myometrial smooth muscle cells (hMSMCs). In particular, the authors wished to determine the association between large-conductance, voltage- and Ca2+-sensitive K+ channel (maxi-K channel) activity, alpha-actin, and the caveolins, and used immunoelectron microscopy to investigate the colocalization of actin with the maxi-K channel.

In preparation for immunoelectron microscopy, thin slices of NP or LP human uterine tissue sample were fixed overnight at 4&176;C in 2% paraformaldehyde and 0.25% glutaraldehyde in phosphate-buffered saline (PBS), and inactive aldehyde groups quenched with 0.05 M glycine in PBS for 15 minutes. The tissue was then permeabilized with 0.01% Triton X-100 in PBS (15 minutes at room temperature), blocked in PBS containing 5% bovine serum albumin (BSA), 0.1% coldwater fish skin gelatin, and 5% fetal bovine serum (FBS), washed in PBS, and incubated overnight at 4&176;C with gentle agitation with a mouse monoclonal anti-actin antibody (Sigma) diluted 1 : 500 in PBS. After washing with PBS, the tissue was incubated with a Nanogold goat anti-mouse IgG (1:2,500 dilution) for 2 hours at room temperature. Next, after washing with PBS, the tissue was blocked as described above and incubated with rabbit anti-maxi-K polyclonal antibody (1:250 dilution) in PBS overnight at 4&176;C. After being washed in PBS, the tissue was then incubated with a goat anti-rabbit 10 nm gold-conjugated secondary antibody (diluted 1 : 2,500 in PBS) for 2 hours at room temperature. Slices were then washed with PBS, postfixed with 2% paraformaldehyde and 0.25% glutaraldehyde in PBS for 5 minutes, and washed with PBS and distilled water. The tissue slices were then enhanced with gold enhancement.

The tissue slices were prepared for transmission electron microscopy by osmication for 90 min in osmium fixation solution (0.2 M sodium cacodylate, 1% osmium, and 6% potassium ferrocyanide at a ratio of 2:1:1), washed with 0.1 M sodium cacodylate for 20 minutes then distilled water for 20 minutes. The tissue was then dehydrated in 50% ethanol for 15 min at RT, 75% ethanol for 15 min at 4&176;C, 95% ethanol for 30 min at -20&176;C, and 100% ethanol for 1 h at -20&176;C. The tissue was subsequently incubated in two parts ethanol and one part Epon-812 (Electron Microscopy Sciences) for 30 minutes at room temperature, then one part ethanol and two parts Epon-812 for 2 hours at room temperature, and finally in Epon-812 overnight at room temperature. The tissue was placed in a Beem capsule in fresh Epon-812 and cured overnight at 60&176;C. After the blocks had cooled, 40 nm sections were cut and mounted on no. 300 nickel grids backed with Formvar and carbon. The grids were subsequently stained with 5% uranyl acetate and lead citrate using a standard electron microscopy grid-staining technique, and viewed using a Hitachi H-7000 electron microscope.

Examination of the resulting electron micrographs shows that the two enlarged particles are still clearly distinct in size, and both are clearly visible. Both the maxi-K channel and actin are localized to the same caveolar invaginations in human myometrium. In addition, disruption of the actin cytoskeleton in cultured hMSMCs by cytochalasin D and latrunculin A greatly increased the open-state probability of the channel, while stabilization of actin cytoskeleton with jasplakinolide negated this effect. Immunocytochemistry indicated that both alpha and gamma-actin, and the maxi-K channel, colocalize with caveolin. Taken together, these results indicate that the actin cytoskeleton is involved as part of a caveolar complex in the regulation of myometrial maxi-K channel function.


Brainard, A. M.; Miller, A. J.; Martens, J. R., and England, S. K.: Maxi-K channels localize to caveolae in human myometrium: a role for an actin-channel-caveolin complex in the regulation of myometrial smooth muscle K+ current. Am. J. Physiol. Cell Physiol., 289, C49-57 (2005).

More information:

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Labeling Other Functional Groups: Carboxylic Acids

More from our technical help files...

I want to label a molecule with Nanogold and the only functional group I have is a carboxylic acid. What should I do?

You can do this using Monoamino Nanogold®. To do so, you need to do here is to convert the carboxylic acid group to a reactive ester, which will then react with the amine - the same reaction that occurs when Mono-Sulfo-NHS-Nanogold is used to label an amine. Some suitable reaction schemes are shown below:

[Carboxylic Acid Nanogold Labeling (13k)]

Reactions for labeling carboxylic acids, using Monoamino Nanogold with (a) EDC (1-Ethyl-3- [3-dimethylaminopropyl] carbodiimide Hydrochloride) / Sulfo-NHS, and (b) 1,1-carbonyl-diimidazole (CDI).

The reaction used in peptide synthesis usually works well - react the carboxylic acid with EDC (1-Ethyl-3- [3-dimethylaminopropyl] carbodiimide Hydrochloride) and Sulfo-NHS to convert it to a reactive Sulfo-N-hydroxysuccinimide ester. You can purchase EDC from a number of sources; its use is described by Pierce.

EDC reacts with a carboxyl group on the molecule to be labeled, forming an amine-reactive O-acylisourea intermediate. This intermediate could then react with Monoamino Nanogold; however, it is also susceptible to hydrolysis, making it unstable and short-lived in aqueous solution. The addition of Sulfo-NHS (5 mM) stabilizes the amine-reactive intermediate by converting it to an amine-reactive Sulfo-NHS ester, thus increasing the efficiency of EDC-mediated coupling reactions. The amine-reactive Sulfo-NHS ester intermediate has sufficient stability to permit two-step cross-linking procedures, which allow the carboxyl groups on one protein to remain unaltered.


Staros, J. V.; Wright, R. W., and Swingle, D. M.: Enhancement by N-hydroxysulfosuccinimide of water-soluble carbodiimide-mediated coupling reactions. Anal. Biochem., 156, 220-222 (1986).

Another reagent that works well in non-aqueous systems is 1,1-carbonyl-diimidazole (CDI). The molecule to be labeled should be dissolved in a small amount of the organic solvent and a small (5-fold to 10-fold) excess of CDA added; the pH is then raised to 7.5 or higher by the addition of aqueous reaction buffer, and the Monoamino Nanogold added.


Staab, H. A., and Rohr; W.; Newer Methods Prep. Org. Chem., 5, 61 (1968).

Can I use Nanogold to label functional groups on a surface rather than in solution?

There is no reason why Nanogold labeling should not work on other types of systems in addition to biological macromolecules, including labeling functional groups on surfaces. Nanogold is well suited to this application because it is soluble in other solvents as well as water and aqueous buffers. In particular, it is highly soluble in dimethyl sulfoxide (DMSO), N,N-dimethylacetamide (DMA), or isopropanol, and these give the option to conduct labeling in non-aqueous, or even in aprotic or hydrophobic environments.

Some issues which may arise when labeling surfaces:

  • How much reagent is needed?
    Because of the greater hindrance in labeling surface targets (since a gold particle with one functionality needs to approach in the right orientation for reaction), you may need a slightly larger excess of reagent than you would for antibody or protein labeling. Make an estimate of the number of reactive sites on the surface you are treating; since Nanogold is about 2.5 nm in diameter (including its coordinated ligands), the highest labeling density you will be able to achieve will probably be close to one Nanogold per 6 nm2.

  • Which solvent is best?
    The wetting properties of the surface may impact whether your reaction is successful or not, since these may also hinder the particles from approaching the surface. Organic solvents such as DMSO or DMA are likely to wet surfaces such as hydrophobic polymers much more effectively than aqueous buffers, and therefore will be much more effective in promoting labeling.

  • What other modifications may be needed?
    If you change the solvent or reaction media, you may need to consider other modifications to ensure that your reaction proceeds. If you are labeling amine sites or using Monoamino Nanogold in an aprotic organic solvent, you should add a small amount of a non-coordinating base to ensure that the amines are deprotonated and react readily. Triethylamine is useful for this application: dissolve a few drops of triethylamine in the reaction solvent, then add a few drops of this solution to the reaction mixture.

More information:

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Biocatalytic Growth of Shaped Gold Nanoparticles

We have described previously how enzymes can deposit metal from solution. Well, in their recent paper in Langmuir, Xiao, Willner and co-workers report that they can also be used to grow odd-shaped gold nanoparticles. Shaped gold nanoparticles were prepared by NAD(P)H-mediated growth of gold nanoparticles in the presence of ascorbic acid, [AuCl4]-, and cetyl-trimethylammonium bromide, which results in the formation of dipods, tripods, and tetrapods. These are distinct in color and spectroscopic properties from spherical gold nanoparticles, and therefore may have potential applications as both enzyme sensors and microscopy labels.

The system used to generate gold nanoparticles is an aqueous solution containing [AuCl4]- (2.4 x 10-4 M, L-ascorbic acid (AA) (6.9 x 10-4 M, cetyltrimethylammonium bromide (CTAB) (4.8 x 10-2 M), and variable concentrations of NADH.

When the pH of this system was adjusted to about pH 11, the rapid formation of gold nanoparticles was observed. In the absence of NADH, a single plasmon absorbance is observed close to 530 nm, the gold nanoparticles produced are spherical (5-8 nm), and their absorbance reaches a constant value after two minutes. In the presence of NADH and ascorbate, the resulting color changes from red to dark blue, and at high concentrations of NADH the UV/visible spectra include two bands at 530 nm and 680 nm. The plasmon band corresponding to the spherical gold nanoparticles (530 nm) decreases, while the new band at 680 nm is intensified as the concentration of NADH is increased. In the absence of NADH, only particles absorbing at 530 nm were formed, and are generated by reduction of the Au(III) salt by ascorbate. In the absence of ascorbate, NADH does not reduce [AuCl4]-, at pH 11, to give any species exhibiting plasmon absorbance bands.

Addition of NADH to a mixture that includes gold nanoparticles pre-synthesized by ascorbate reduction did not result in formation of the blue-colored particles, and the plasmon band at 530 nm remained unchanged, suggesting that the pre-synthesized nanoparticles are protected by CTAB ligands that render them inaccessible to the catalytic action of NADH. Furthermore, mixing the [AuCl4]-/AA/CTAB/NADH system at neutral pH does not yield any nanoparticles. The formation of the blue-colored NPs is also time dependent: the absorbance at 680 nm increases with time and reaches a saturation value. As the concentration of NADH in the system increases, the level of saturated absorbance is higher, and these results allow the derivation of a mechanism for the controlled growth of gold nanoparticles in the presence of NADH.

The structural features of gold nanoparticles generated with and without NADH were analyzed by high-resolution transmission electron microscopy (HRTEM). In the absence of NADH, spherical particles 5 to 8 nm in diameter were observed. In the presence of high concentrations of NADH (4.0 x 10-6 M), nanoparticles of variable shapes were observed. Statistical analysis revealed that in addition to spherical particles (30%), most particles were composed of distinct structures. comprising L-shaped dipods (12%), tripods (45%), and tetrapods (13%). HRTEM analysis of the tripod structure showed a lattice of planes exhibiting an interplanar distance of 0.235 nm, corresponding to {111} type planes of crystalline gold). The orientation of the crystallite was found to be [011], and possible growth directions for the three pods are of (211) type. A similar HRTEM analysis of the tetrapod indicates that the crystallite is oriented along the [010] direction (only {220} type interplanar distances of 0.144nm are observed). At lower NADH concentrations (5.4 x 10-7 M) stepwise formation of nanoparticle shapes was observed. While at high NADH concentrations, nanoparticle shapes with "arms" about 20 nm long and 2 - 5 nm in width are formed, substantially shorter pods about 6-8 nm long are observed in these "embryonic-type" particles generated at lower NADH concentrations. Statistical analysis indicated that the resulting gold nanoparticle mixture includes about 60% spherical particles, and about 10%, 25%, and 5% respectively of "embryonic-type" dipod, tripod, and tetrapod shapes.

The structural features of the NADH-grown shaped nanoparticles correlate well with the absorbance spectra of the solutions. The blue color and 680 nm absorbance of the system containing well-developed dipods, tripods, and tetrapods was attributed to a longitudinal plasmon exciton that exists in rodlike gold nanoparticles; similar absorbance features have been observed at 680 - 700 nm for tripod and tetrapod structures prepared chemically. The less intensive band at 530 nm was attributed to residual spherical particles. The spectrum of the system that includes less developed shaped nanoparticles generated at a lower NADH concentration consists of a significant band at 530 nm, corresponding to the spherical NPs, and a broad red shifted shoulder that indicates the formation of longitudinal-shaped structures. The distinctive morphological and spectroscopic properties make these particles potentially useful as biosensors and markers at both the optical and electron microscopic levels.

From the spectral and structural features of the nanoparticles and from control experiments, a mechanism for the growth of the shaped nanoparticles was inferred. The reduced cofactor NADH is unable to reduce [AuCl4]- to Au(0) (or to Au(I)) due to its low concentration; however, ascorbate reduces, however, [AuCl4]- to Au(0) nanocrystals, and their formation then activates the NADH-mediated reduction to produce shaped nanoparticles. The electrocatalytic oxidation product of NADH bound to single-crystalline gold surfaces (NAD+) binds with different affinities to the different faces, with a predominantly favored affinity to the (011) face. This increases the local concentration of the reduced cofactor at the nanoparticle surface, thus facilitating the catalyzed reduction of [AuCl4]- to Au(0) and enlargement of the particle. However, the resulting NAD+ binds effectively to the (011) face, blocking further NADH-mediated growth on that face. Adsorption of CTAB to the gold nanoparticles then controls their direction of growth in the (211) directions. Further NADH-mediated growth along these directions yields the tripod structures.


Xiao, Y.; Shlyahovsky, B.; Popov, I.; Pavlov, V., and Willner, I.: Shape and color of au nanoparticles follow biocatalytic processes. Langmuir, 21, 5659-5662 (2005).

More information:

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Nanogold® Labeling Demonstrates Adenoviral Targeting Modification

We have reported previously on the utility of Nanogold® labeling of engineered viruses as a means of making nanopatterned arrays of gold particles. Kreppel and colleagues have taken the same approach and applied it to genetherapy, using both protein engineering and chemical modification of adenovirus in order to overcome its limitations as a vector for gene therapy. The adenovirus vectors used were Ad5-based E1-deleted first-generation vectors, generated by transfection of the corresponding infectious plasmids into the E1-transcomplementing transcomplementing N52E6 cell line followed by subsequent vector amplification. Three Ad5-based, E1-deleted adenovirus vectors were prepared, genetically modified to present in the solvent-exposed fiber HI-loop the short cysteine-containing motifs LIGGGCGGGID (Ad1Cys), LIGCGCGCGID (Ad3Cys), and LICCCCCID (Ad5Cys). During vector particle purification in the presence of atmospheric oxygen, Ad5Cys formed macroscopically visible aggregates, which were not observed in the case of Ad1Cys and Ad3Cys or unmodified control vector Ad1stGFP; these aggregates could be resolved by addition of the reducing reagents tris (caroxylethyl) phosphine (TCEP) or dithiothreitol (DTT) (final concentration 10 mM), suggesting that the genetically introduced thiol groups of the cysteine residues had crosslinked the vector particles by forming interparticle disulfide bonds.

To couple Monomaleimido Nanogold® particles to the different vector preparations, 1011 vector particles (purified under conditions with reduced oxygen) were incubated in degassed phosphate-buffered saline (PBS) with and without 10 mM TCEP as a reducing reagent for 4 hours at 48°C in a total volume of 500 microliters. Monomaleimido Nanogold, freshly dissolved in 20% DMSO in PBS, was added in a 100-fold molar excess over cysteine residues as calculated for Ad5Cys. Coupling was performed at 48°C overnight. To stop the reaction, a 2-fold molar excess of free cysteine over gold particles was added and incubation continued 2 hours at room temperature. An aliquot containing 1010 vector particles was supplemented with SDS-PAGE loading buffer devoid of reducing reagent and incubated at 37°C for 30 minutes to disrupt Ad particles before being loaded on an 8% polyacrylamide gel. The samples were not boiled in order to avoid compromising the organic shell surrounding the gold particles. After electrophoresis, silver staining was performed with the LI Silver Enhancement Kit.

The coupling products were analyzed by seminative SDS-PAGE and gold-specific silver staining. The gold-specific silver staining revealed that only vectors that had been genetically modified to bear cysteine residues on their surfaces reacted with the Monomaleimido Nanogold. To further analyze the specificity of coupling, gold-conjugated Ad5Cys vector particles were purified by ultrafiltration: the unpurified particles, the purified particles, and the flow-through were all then subjected to seminative SDS-PAGE and gold-specific silver staining. The results demonstrated that the maleimide-based chemistry could be used to specifically modify the surface of thiol-bearing vector particles. Other molecules could be linked in a similar manner, as was found using a 20,000 MW maleimido-polyethylene glycol derivative.

Staining of Nanogold-labeled entities in gels is an effective and highly sensitive method for visualization, although the shift due to the incorporation of the gold is unpredictable for the separation of gold conjugates. Often the shift in their migration is considerably less than that predicted from their molecular weight, possibly because the gold particles are spherical, smaller than biomolecules of equivalent mass (due to the high atomic mass of gold) and embed themselves into biomolecules. If you plan to use gel electrophoresis for the separation of Nanogold conjugates, we recommend that you try it on a small scale first. Do two runs side-by-side: stain one with a conventional general stain for the type of molecule (protein or oligonucleotide) that you are using, and develop the second using LI Silver or a gold-silver enhancement reagent (do not use a silver protein stain!). The staining pattern in the second run will tell you where the gold is located, while in the first, the stain will show all the bands: by comparing the two results, you will be able to check whether the gold is bound to the molecule of interest, and if so, which band.


Kreppel, F.; Gackowski, J.; Schmidt, E., and Kochanek, S.: Combined genetic and chemical capsid modifications enable flexible and efficient de- and retargeting of adenovirus vectors. Mol. Ther., 12, 107-117 (2005).

More information:

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Other Recent Publications

A new report by Farrer and co-workers in June's indicates that gold nanoparticles possess useful luminescence properties. This group used multiphoton-absorption-induced luminescence (MAIL), in which the absorption of multiple photons from a near-infrared ultrafast laser is used to produce luminescence, to generate efficient luminescence from gold nanoparticles from 125 nm to as small as 2.5 nm in diameter. Highly efficient photoluminescence was generated upon irradiation with sub-100-fs pulses of 790-nm light, and strong emission was observed at excitation intensities comparable to or less than those typically used for multiphoton imaging of fluorescently labeled biological samples. The gold particles showed polarized emission, can radiate more efficiently than single molecules, do not exhibit significant blinking, and were found to be photostable under hours of continuous excitation. Therefore, it is possible that metal nanoparticles would be a viable alternative to fluorophores or semiconductor nanoparticles (quantum dots) for biological labeling and imaging.


Farrer, R. A.; Butterfield, F. L.; Chen, V. W., and Fourkas, J. T.: Highly efficient multiphoton-absorption-induced luminescence from gold nanoparticles. Nano Lett., 5, 1139-1142 (2005).

In the same issue, Tseng and co-workers report another novel application for gold nanoparticles: using 30 nm polyaniline nanofibers decorated with 1 - 20 nm gold nanoparticles for non-volatile memory. A relatively uniform distribution of nanometer-sized gold nanoparticles is achieved by controlling the time and temperature of a redox reaction between 30-nm-diameter polyaniline nanofibers and chloroauric acid. A nonvolatile plastic digital memory device, consisting of the plastic composite film sandwiched between two electrodes, was fabricated by the deposition of a bottom (column) aluminum electrode with a thickness of 80 nm by thermal evaporation in a chamber under a pressure of 1 x 10-5 Torr. A 70-nm-thick active layer was then formed by spin coating an aqueous solution of ~0.1 wt % polyaniline nanofiber/gold nanoparticle composite in 1.5 wt % poly(vinyl alcohol); the poly(vinyl alcohol) serves as an electrically insulating matrix for the composite. An external bias was used to program the ON and OFF states of the device, that were separated by a 3-orders-of-magnitude difference in conductivity. ON-OFF switching times of less than 25 ns are observed by electrical pulse measurements. The devices possess prolonged retention times of several days after they have been programmed. Write-read-erase cycles are also demonstrated. The switching mechanism is attributed to an electric field-induced charge transfer from the polyaniline nanofibers to the gold nanoparticles.


Tseng, R. J.; Huang, J.; Ouyang, J.; Kaner, R. B., and Yang, Y.: Polyaniline nanofiber/gold nanoparticle nonvolatile memory. Nano Lett., 5, 1077-1080 (2005).

To keep the theme of novel nanoparticle constructs going...Wanunu and co-workers, in a recent Journal of the American Chemical Society paper, report the assembly of gold nanoparticles by metal coordination, using partial substitution of a coordinated ligand with a zinc-binding moiety to introduce a zirconium coordination site. Gold nanoparticle mono- and multilayers were constructed on gold surfaces. Hydrophilic Au NPs (6.4 nm average core diameter), capped with a monolayer of 6-mercaptohexanol, were modified by partial substitution of the 6-mercaptohexanol with bishydroxamic acid disulfide ligand molecules. A monolayer of the ligand-modified nanoparticles was assembled, via coordination with Zr4+ ions, onto a semitransparent gold substrate (15 nm Au, evaporated on silanized glass and annealed) precoated with a self-assembled monolayer of bishydroxamate disulfide. Layer-by-layer construction of nanoparticle multilayers was achieved by alternate binding of Zr4+ ions and ligand-modified nanoparticles onto the initial nanoparticle layer. Atomic force microscopy (AFM), ellipsometry, wettability, transmission UV-vis spectroscopy, and cross-sectional transmission electron microscopy all showed regular growth of nanoparticle layers, with a similar particle density in successive layers and gradually increased roughness. Step-by-step assembly of more elaborate nanostructures can be achieved by using different ligand-possessing components. This was demonstrated by introducing a nanometer-scale coordination-based organic multilayer, consisting of small branched hydroxamates assembled using Zr4+ ions, between the gold surface and the initial nanoparticle layer. Electrical characterization of the nanoparticle films using conductive AFM confirmed the barrier properties of the organic spacer multilayer.


Wanunu, M.; Popovitz-Biro, R.; Cohen, H.; Vaskevich, A., and Rubinstein, I.: Coordination-based gold nanoparticle layers. J. Amer. Chem. Soc., 127, 9207-9215 (2005).

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .


View Cart
© 1990-2015 Nanoprobes, Inc. All rights reserved. Sitemap