Updated: October 14, 2005

N A N O P R O B E S     E - N E W S

Vol. 6, No. 10          October 14, 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.

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NTA-Ni(II) Gold Finds the Photosystem II PsbH Subunit in Synechocystis 6803

You've tried gold-antibody probes and Fab'-Nanogold® probes, but you still want higher resolution? If you can use a smaller targeting agent, the gold can get closer to the target. One of the smallest targeting agents around is the Nickel (II) nitriloacetic acid (NTA) chelate; this nickel coordination compound has two loosely coordinated water molecules that are readily displaced by histidines, and the complex therefore targets polyhistidine sequences in proteins.

At Nanoprobes, we have developed a NTA-Ni(II)-Nanogold, a novel probe in which this chelate is linked to gold nanoparticles and used to target the gold labels to protein sites marked with engineered polyhistidine tags. This probe has significant advantages over conventional antibody and protein probes:

  • The nitrilotriacetic acid - Ni(II) chelate is much smaller than an antibody or protein, so labeling resolution is higher: once it has bound, the gold is much closer to its target. This makes NTA-Ni(II)-Nanogold ideal for localizing sites in protein complexes at molecular resolution.

  • Because it is so small, NTA-Ni(II)-Nanogold is better able to penetrate into specimens and access restricted sites within them.

  • NTA-Ni(II)-Nanogold is made with a modified gold particle, with very high solubility and stability. At 1.8 nm in size, is readily visualized by electron microscopy.

  • Binding constants for Ni(II)-NTA are very high due to the chelate effect of multiple histidine binding and multiple Ni(II)-NTA functionalization. Dissociation constants are estimated to be between 10-7 to 10-13 M-1. For many applications, this provides binding strengths comparable to antibodies.

Bumba and co-workers use these strengths to achieve macromolecular localization of the Photosystem II (PSII) PsbH subunit in Synechocystis 6803, described in their recent paper in the Journal of Structural Biology. Following the recent localization of His-tagged proteins on viral capsids, and an earlier study of PsbH in PSII by Büchel and co-workers, Bumba and group labeled the PsbH subunit with high precision even though it resides on the side of the complex that lies in contact with the grid in its preferred orientation.

[Structure, PsbH binding and STEM image of NTA-Ni(II)-Nanogold (43k)]

left: Structure of Ni-NTA-Nanogold® showing interaction with a His-tagged protein. Inset shows localization of PsbH subunit within Photosystem II using NTA-Ni(II)-Nanogold, showing resolution; right: Knob protein from adenovirus cloned with 6x-His tag, labeled with Ni-NTA-Nanogold, column purified from excess gold, and viewed in the scanning transmission electron microscope (STEM) unstained (Full width approximately 245 nm).

To identify the location of the His-tagged PsbH subunit within PSII, dimeric PSII complexes (obtained by solubilization of thylakoid membranes with 1% dodecyl-maltoside (DM) in thylakoid buffer with 100mM NaCl, centrifugation for 30min at 60,000 g, and affinity chromatography of the supernatent on EMD Chelate gel charged with Ni2+) were applied to grids that were glow discharged for ten seconds prior to use. The PSII sample was applied first, at a concentration of 5.6 mg/ml Chl in 50 mM MES (pH 6.5) and 0.03% DM. Excess buffer was removed using filter paper. The grid was then incubated upside-down on a droplet of NTA-Ni(II)-Nanogold label solution (170 nM in 50 mM MES, pH 6.5) for ten minutes at 4°C. Labeling was terminated by removing the grid from the droplet, rinsing thoroughly with water and staining with 1% uranyl acetate. This labeling procedure provides the advantage of a more accurate localization of the targeted site, since no additional protein densities are present, as compared with conventional immunogold labeling procedures, and gave specific labeling of multiple His sites on the protein complexes. Its effectiveness was confirmed by performing the labeling in a buffer containing 30mM imidazole: this competes with the His-tag for the NiNTA sites, and eliminated any labeling of proteins.

Labeled, negatively stained PSII complexes were visualized in the electron microscope. A typical EM image showed dispersed particles with uniform size and shape, almost free of contaminants: these are dimeric PSII particles, mostly in top-view projections. Relatively few PSII dimers exhibited gold label; the proportion was not significantly changed by using different labeling conditions, such as incubation time, label concentration, pH and temperature. All the projections had the same type of handedness and no mirror images were detected, indicating preferred orientation of the PSII dimers with their stromal side to the carbon support film. Since the PsbH His-tag is located on the stromal side of the complex, the relatively low labeling probably reflects the preferential binding of the PSII particles, which renders the labeling site somewhat inaccessible when the particles are orientated with their stromal sides to the carbon support film; another reason for the lower gold-label affinity may be that in cyanobacteria, the N terminus of the PsbH protein is 19 amino acids shorter compared to the C. reinhardtii and the labeling site is further inside the complex.

The shorter N terminus of the Synechocystis PsbH protein enabled more precise identification of the location of the His site within the PSII complex. A careful comparison of the location of the gold clusters in C. reinhardtii and Synechocystis PSII revealed that gold label in Synechocystis preparation is slightly shifted with respect to the longer edge of the complex. The location of the PsbH subunit is in good agreement with the assignment of the PsbH subunit in the recent model of Ferreira, and suggests that a single transmembrane helix close to the CP47 subunit corresponds to the PsbH protein. These results suggest that the resolution of labeling and freedom from additional protein density makes labeling of His sites using NiNTA gold clusters a powerful approach to locating specific proteins within multisubunit protein complexes.


    Bumba, L.; Tichy, M.; Dobakova, M.; Komenda, J., and Vacha, F.: Localization of the PsbH subunit in photosystem II from the Synechocystis 6803 using the His-tagged NiNTA Nanogold labeling. J. Struct. Biol., 152, 28-35 (2005).

Previous references:

  • Hainfeld, J. F.; Liu, W.; Halsey, C. M. R.; Freimuth, P., and Powell, R. D.: Ni-NTA-Gold Clusters Target His-Tagged Proteins. J. Struct. Biol., 127, 185-198 (1999).

  • Buchel, C.; Morris, E.; Orlova, E., and Barber, J.: Localisation of the PsbH subunit in photosystem II: a new approach using labelling of His-tags with a Ni(2+)-NTA gold cluster and single particle analysis. J. Mol. Biol., 312, 371-379 (2001).

More information:

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Bigger NTA-Ni(II) Gold

The hexahistidine (6x-histidine, or His) tag is one of a number of genetic "tags" which are inserted into expressed proteins by incorporating a "tail" that codes for the protein in the DNA used to construct the protein of interest. Other genetic tags include Flag, that binds to anti-flag antibodies, strep-tag that binds to streptactin, c-myc that binds to antibody, calmodulin binding domain that binds to calmodulin, cellulose binding domain that binds cellulose, glutathione-S-transferase that binds glutathione, and streptavidin binding peptide that binds streptavidin. These tags are used to incorporate an affinity - often, this is an affinity towards a metal chelate column used to purify the target protein, or an antibody used to bind it.

These tags can also function as targets for tracking, detecting or labeling the protein: a detection label is conjugated to the tag-binding entity and the conjugate used to localize the protein. For example, green fluorescent protein expressed as a fusion protein with the protein of interest is widely used for studying proteins in living cell by light microscopy. Electron microscopy, however, requires a dense label, such as a gold particle. One approach has been to use gold-labeled anti-GFP antibodies; but another approach, with significant advantages, is to derivatize gold nanoparticles with the binding couple for the tags used in protein expression. These tags may be much smaller that GFP, and the corresponding binding couples can be much smaller than antibodies. One example is a 1.8 nm Nanogold particle with a synthetically introduced peripheral NTA-Ni(II) group, which has now been used to successfully localize a number of His-tagged proteins with high resolution by electron microscopy.

The Nanogold particle is directly visible in ice or on a thin film, but may be more difficult to see in stained or thicker samples, such as tissue sections. Although it may be enlarged using silver enhancement or gold enhancement, a larger particle, such as 5 nm, may be seen directly without enhancement, and its use would reduce the steps required for preparation and avoid the sometimes large size variations that can occur with autometallography. Now we've got rid of the big targeting protein, there's room to make the gold bigger - and there are plenty of applications for a larger NTA-Ni(II) gold probe. Insertion of His tags is now feasible for a wide range of different proteins and constructs, so larger gold targeted to His tags would be a simple, versatile and effective labeling and detection reagent. At Microscopy and Microanalysis 2005, we presented preliminary results describing the preparation of a nitrilotriacetic acid - nickel (II) - 5 nm gold probe. The extended abstract of this presentation is now available as a research application on our web site.

The NTA-Ni(II)-derivatized particles were prepared similarly to the NTA-Ni(II)-Nanogold particles described previously. A transmission electron micrograph is shown below, together with a chromatogram showing the formation of a new peak corresponding to the gold conjugate when the functionalized gold particles were incubated with the protein ISWI from the ACF chromatin remodeling complex, synthesized with a His tag. In a control experiment in which the NTA-Ni(II)-derivatized particles were incubated with ISWI without the 6x-His tag, virtually no binding to the protein was seen, as evidenced by the absence of the conjugate peak.

[Chromatographic separation of NTA-Ni(II)-[Au5nm] labeled protein, and TEM image (60k)]

left: Chromatogram showing new peak (arrow) when 6x-His protein (ISWI) is incubated with nitrilotriacetic acid (NTA) - Nickel (II) - 5 nm gold. Right: TEM image of functionalized 5 nm gold particles.


Reddy, V.; Lymar, E.; Hu, M., and Hainfeld, J. F.: 5 nm Gold-Ni-NTA binds His Tags. Microsc. Microanal., 11 (Suppl. 2),; Price, R.; Kotula, P.; Marko, M.; Scott, J. H.; Vander Voort, G. F.; Nanilova, E.; Mah Lee Ng, M.; Smith, K.; Griffin, P.; Smith, P., and McKernan, S. (Eds.), Cambridge University Press, 1118CD (2005).

More information:

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Nanogold® Labeling: pH and Solvent

We are frequently asked about using Nanogold® in non-standard or non-aqueous buffers and solvents, for example with insoluble peptides and small molecules. Nanogold is not just for use in aqueous systems - it is also soluble, and may be used, in a wide range of other solvents. However, while the particles themselves are mostly stable, the conjugation chemistry has been developed primarily for use in aqueous systems, and you may need to explore alternatives if you need to label under very different conditions.

What is the solubility of Nanogold in alternative solvents?

  • Aqueous buffers: Nanogold is soluble in water and in most aqueous buffers, in most cases to at least 1,000 nmol/mL, providing a solution that is concentrated enough for most labeling experiments. However, solution may require gentle agitation or even vortexing for a short time before the reagent dissolves completely.

  • Alcohols: Nanogold is soluble in alcohols, especially ethanol, but it is more soluble in alcohol-water mixtures than in alcohols alone. If you are using ethanol precipitation for a Nanogold-labeled oligonucleotide, the degree to which the labeled oligonucleotide will be precipitated will vary depending on the length (and hence contribution to the solubility properties of the conjugate) of the oligonucleotide. Ethanol precipitation alone may give inadequate separation of Nanogold-labeled oligonucleotide, so we recommend that chromatographic separation (reverse-phase, gel filtration and hydrophobic interaction) and UV/visible spectroscopy are used in addition to ethanol precipitation to separate and characterize the reaction products in each phase of the reaction mixture. For a more detailed discussion of oligonucleotide labeling, see our May 2004 newsletter.

  • Organic solvents: Nanogold is highly soluble in dimethylsulfoxide (DMSO). Dissolving in a small quantity of DMSO before adding water will usually speed up solution and ensure an effective reaction. Up to 20% DMSO is also well tolerated by many biological molecules. Therefore we recommend predissolving Nanogold reagents in a small quantity of DMSO (10% of the final reaction volume) then adding water. Nanogold is also soluble in acetonitrile, and it is highly soluble in mixtures of isopropanol, another organic solvent that is well tolerated by many biologicals, with water: predissolving in 10% isopropanol is a good alternative if DMSO is not suitable for your application. Nanogold is not compatible with N,N-dimethylformamide (DMF): however, in many applications you can use N,N-dimethylacetamide (DMA), which has very similar properties, and Nanogold is soluble and stable in this solvent.

  • Halogenated solvents: If you are labeling lipids or other hydrophobic entities, Nanogold is soluble in mixtures of up to 50% dichloromethane (methylene chloride) or trichloromethane (Chloroform) with alcohols, and these should sufficiently solubilize both lipids and Nanogold to allow reaction.

What is the stability of Nanogold towards acids and bases? Are there any conditions I need to be careful to avoid?

  • Acids: Nanogold is stable to about pH 2-3 for short periods (up to a few hours at room temperature). However, prolonged exposure to pH values below about 4 can lead to degradation of the Nanogold, and is not recommended. The degradation risk is particularly high if the ionic strength or salt concentration of the solution is high. Under such conditions, Nanogold tends to form larger 'colloidal' particles that may be red or blue-purple in color. While brief exposure to dilute acetic acid is usually not problematic, exposure to trifluoroacetic acid at concentrations higher than 0.1 M for longer than an hour is much more likely to cause problems, and we recommend that you avoid trifluoroacetic acid exposure whenever possible.

  • Bases: Nanogold is generally stable towards bases up to about pH 10, although high pH values can cause solubility problems. Addition of 10 - 20% DMSO may help. For stronger bases, test a small amount of Nanogold in the solution you plan to use: monitor by UV/visible absorption spectroscopy. If the spectrum remains unchanged, the Nanogold is stable.

  • Other compounds: Thiols have a very high affinity for gold. Thiols will displace the surface functionalities of Nanogold, and exposure to greater than 1 mM concentrations of thiols for longer than about an hour will result in breakdown of the gold core into smaller gold-containing species. Thiol-containing compounds should be blocked (use N-ethylmaleimide) or removed by gel filtration before Nanogold is introduced; if you need to use a thiol-containing reducing agent, such as dithiothreitol (DTT) or mercaptoethylamine hydrochloride (MEA), to reduce a disulfide to a thiol before labeling with Monomaleimido Nanogold, the excess reducing agent must be removed completely by gel filtration before the Nanogold is added so that it will not react with the maleimide. Dialysis does not give acceptable purification in this situation.

How do alternative buffers and solvents affect reactivity, and what can I do to ensure a successful reaction?

The reaction of maleimides with thiols proceeds best at near-neutral pH values of 6.0 to 7.0, while the reaction of primary amines with Sulfo-NHS- groups is most selective at pH 7.5-8.2. At higher pH values, hydrolysis of the reactive group begins to compete with the specific reaction, and at lower pH values, the desired reaction may not occur, or other undesirable reactions may interfere. If you plan to conduct labeling in different buffers or solvents, you should either select conditions which parallel these, or look for an alternative reaction scheme.

  • Controlling pH in organic solvents: Addition of a small amount of an organic-soluble base, such as triethylamine, may help the reaction of Mono-Sulfo-NHS-Nanogold under these conditions but make sure that your base does not contain primary or secondary amines that may react. If you are using Monomaleimido Nanogold, avoiding the addition of either acids or bases should help produce a successful reaction. Maintaining a proportion of water in the reaction mixture is helpful in this case because it will dissolve the buffer salts with which this reagent is packaged, and help keep the pH in the appropriate range.

  • Alternative reactions: If you require acidic or basic conditions to solubilize the compound you are labeling, you may need to look for an alternate labeling reaction. The best approach is to use Monoamino Nanogold, and look for either a cross-linker which is compatible with your reaction conditions, or a functional group such as a carboxylic acid, an aldehyde or ketone, or an alcohol in your molecule that can be activated to give a reaction site for Monoamino Nanogold.

Cross-linker resources:

More information:

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SEM-ISH: Bacterial Detection by rRNA Sequence using Gold-Enhanced Nanogold®

Our gold enhancement technology, which is similar to silver enhancement except that gold, not silver, is deposited onto gold nanoparticles, has significant advantages for scanning electron microscopy:

  • Gold gives a much stronger backscatter signal than silver.
  • May be used in physiological buffers (chlorides precipitate silver, but not gold).
  • The metallographic reaction is less pH sensitive than that of silver.
  • May safely be used before osmium tetroxide - it is not etched.
  • GoldEnhance is near neutral pH for best ultrastructural preservation.
  • Low viscosity, so the components may be mixed easily and accurately.

Kenzaka and co-workers combine two SEM detection with in situ hybridization in a novel method for bacterial detection based on their rRNA sequence, "SEM-ISH," described in their recent paper in Applied and Environmental Microbiology. The authors used their method to characterize the bacterial community composition on the surface of river sediment particles before and after cell dispersion treatment by sonication. Target cells were hybridized with oligonucleotide probes, which were then detected with Nanogold®-streptavidin. Gold enhancement was used for amplification of probe signals from hybridized cells: the hybridized cells generated a strong backscatter electron signal due to internal accumulation of gold atoms.

Surface natural river water samples from sites considered to be polluted by organic carbon (total organic carbon 5 to 7 mg/liter) and from a pond were collected in sterile 500-ml glass bottles, and densities of heterotrophic bacterioplankton determined with DAPI (4',6'-diamidino-2-phenylindole) at a final concentration of 1 microgram/mL were 1.4 X 107, 2.0 X 107, 5.3 X 105, and 1.0 X 106 cells per mL, respectively. 40 ml portions were fixed with 4% paraformaldehyde at 4°C for 16 hours. After fixation, 0.5 to 10 mL portions were filtered through gelatin [0.1% gelatin, 0.01% CrK(SO4)2]-coated polycarbonate white filter (0.2 micron pore size, 25-mm diameter) and rinsed twice with filtered deionized water. Sediment was taken from sand oxidized surface (upper 2 cm) at Kuwazu; sand particles were fixed with 4% paraformaldehyde at 4°C for 16 h.

The following oligonucleotide probes were used for in situ hybridization: (i) EUB338 complementary to a region of the 16S rRNA conserved in the domain bacteria; (ii) NON338, used as a negative control; (iii) ALF1b, complementary to a region of the 16S rRNA specific the alpha subclass of Proteobacteria; (iv) BET42a, complementary to a region of the 23S rRNA specific for the beta subclass of Proteobacteria; (v) GAM42a, complementary to a region of the 23S rRNA specific for the gamma subclass of Proteobacteria; (vi) CF319, complementary to a region of the 16S rRNA specific Cytophaga-Flavobacterium phylum; and (vii) ES445, complementary to a region of the 16S rRNA specific for Escherichia-Shigella. All probes were labeled with biotin or CY3 at the 5' end.

for SEM-ISH on membrane filter for water samples, polycarbonate filters with bacterial cells from water samples were cut into 12 sections. Each filter section was transferred to a 1.8 mL microtube and dehydrated in vacuo. For the EUB338, NON338, ALF1b, BET42a, and GAM42a probes, hybridization and washing were performed in hybridization buffers consisting of 0.9 M NaCl, 20 mM Tris-HCl (pH 7.4), 0.01% sodium dodecyl sulfate (SDS), and a variable concentration of formamide; For ES445 probe, hybridization was performed in a moisture chamber at 41°C for 2 h with hybridization buffer (0.45 M NaCl, 20 mM Tris-HCl, pH 8.0, 0.1% sodium dodecyl sulfate) containing 2 ng of probe per microliter. The washing step was done at 41°C for 30 minutes with washing buffer (0.08 M NaCl, 20 mM Tris-HCl, pH 8.0, with 0.1% sodium dodecyl sulfate). Samples were incubated with 100 microliters of alkali solution (0.5 M NaOH, 1.5 M NaCl) for 10 minutes at room temperature (ca. 25°C) to allow the gold labeling reagents to enter the cell, and neutralized with 100 microliters of 0.5 M HCl in 1.5 M NaCl. The filter sections were rinsed with deionized water and blocked with 100 microliters of PBS containing 0.1% gelatin and 0.1% Tween 20. Nanogold-streptavidin diluted 1 : 1,000 in 1% bovine serum albumin in PBS) was then added and the specimens incubated for 30 minutes at room temperature. The filter sections were then soaked in PBS containing 0.1% gelatin and 0.1% Tween-20 at room temperature for 10 minutes, then soaked in filtered, deionized water. For gold autometallography, they were soaked in 100 microliters of GoldEnhance EM, incubated for 10 minutes at room temperature in the dark.

For sediment particles, the filter sections were soaked in PBS containing 0.1% gelatin and 0.1% Tween 20 for 10 minutes and in filtered deionized water for 10 minutes. Sediment particles of 0.5 to 1.5 mm were transferred to a 1.8 mL microtube, resuspended in filtered deionized water, sonicated for 5 minutes at 400 kHz and 125 W using a bath sonicator, and washed three times with filtered deionized water. Both sonicated sediment particles and untreated particles were subjected to SEM-ISH; the sediment particles were transferred to a new microtube, and SEM-ISH in the microtube was performed as described for the membrane filters for the water samples.

For SEM, slides, polycarbonate filters, or sediment particles were placed on an SEM pore connected to an LV cooling holder, and incubated with 50 microliters of t-butyl alcohol at room temperature for 10 minutes. The holder was then covered with a cooling cap and chilled with liquid nitrogen for 30 seconds, the cap removed, and the holder placed immediately in the microscope specimen chamber. Freeze-drying was carried out in the chamber. After 20 minutes, samples were observed in reduced vacuum (30 Pa), i.e., low vacuum mode. Samples were further sputter coated with evaporated gold for 2 minutes for observation under high vacuum. A 5 to 20 kV accelerating voltage was used for both low and high vacuum mode imaging. Specimens were placed 14 mm from the base detector, and electron micrographs obtained at magnifications of X 60 to X 20,000 using both secondary electron (SE) and backscatter electron (BSE) detectors for high vacuum imaging and BSE detectors for low vacuum imaging. For quantitative evaluation, at least 200 cells in different fields were counted in triplicate. Counting results were always corrected by subtracting signals observed with the NON338 probe.

When SEM-ISH was applied to analyze bacterial community composition in freshwater samples, bacterial cell counts determined by SEM-ISH with rRNA-targeted probes for major phyla within the domain Bacteria were highly correlated to those obtained by fluorescent in situ hybridization (FISH). The bacterial composition on surface of river sediment particles before and after cell dispersion treatment by sonication was successfully revealed by SEM-ISH, and direct enumeration of bacterial cells on the surface of sonicated sediment particles by SEM-ISH demonstrated that members of Cytophaga-Flavobacterium existed tightly on the surface of particles. SEM-ISH allows the user to define the number and distribution of phylogenetically defined cells adhering to material surfaces, which is difficult with FISH, and provides a new method for the electron microscopic study of microorganisms in their natural environment.


Kenzaka, T, Ishidoshiro, A, Yamaguchi, N, Tani, K, Nasu, M.: rRNA sequence-based scanning electron microscopic detection of bacteria. Appl. Environ. Microbiol., 71, 5523-5531 (2005).

More information:

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Who You Gonna Call? Contact Information

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Other Recent Publications

York-Dieter Stierhof and group continue to use Nanogold® labeling with silver enhancement to good effect in their recent paper in the Journal of Cell Biology, in which they report the use of pre-embedding Nanogold labeling with HQ Silver enhancement to study the distribution and role of rootletin, a protein distantly related to C-Nap1 and previously identified as a structural component of the ciliary rootlet in murine photoreceptor cells and human T lymphoblastoid cells, as a centrosome linker. For pre-embedding immunoelectron microscopy of whole cells, U2OS cells, grown on coverslips and fixed with 4% PFA for 10 min, were permeabilized with PBS-0.5% Triton X-100 for 2 minutes, blocked with 1% bovine serum albumin (BSA) in PBS for 10 minutes, incubated with primary antibodies (rabbit antirootletin serum (R145; 1:1,000), mouse antigamma-tubulin mAb (1:1,000; GTU-88), and rabbit antiC-Nap1 affinity-purified IgG, then incubated with goat antirabbit IgG Nanogold (1:501:100), and enhanced with HQ Silver. The results show that endogenous rootletin forms striking fibers emanating from the proximal ends of centrioles; rootletin was also shown to interact with C-Nap1, a protein previously implicated in centrosome cohesion. Similarly to C-Nap1, rootletin is phosphorylated by Nek2 kinase and is displaced from centrosomes at the onset of mitosis. While rootletin overexpression results in the formation of extensive fibers, small interfering RNAmediated depletion of rootletin or C-Nap1 causes centrosome splitting, suggesting that both proteins contribute to maintaining centrosome cohesion. The ability of rootletin to form centriole-associated fibers suggests a dynamic model for centrosome cohesion based on entangling filaments, rather than continuous polymeric linkers. component


Bahe, S.; Stierhof, Y. D.; Wilkinson, C. J.; Leiss, F.; and Nigg, E. A.: Rootletin forms centriole-associated filaments and functions in centrosome cohesion. J. Cell. Biol., 171, 27-33 (2005).

Kim, Lee and co-workers take a look at the factors behind gold nanoparticle aggregation in their recent paper in Langmuir. Colloidal gold nanoparticles 20 to 50 nm in diameter were prepared by trisodium citrate reduction, then aggregated by the addition of benzyl mercaptan to manipulate the interparticle interaction in the absence of cross-linking effects. UV-visible absorption spectroscopy, surface-enhanced Raman scattering (SERS), quasi-elastic light scattering, and zeta-potential measurement were used to characterize the resulting nanoparticle aggregates. The results suggest that the replacement of the trivalent citrate ions adsorbed on the nanoparticle surface with monovalent benzyl mercaptan ions should destabilize the particles, causing aggregation and hence an increase in the measured size of nanoparticle aggregates. These experimental results may be successfully rationalized by the classical DLVO (Derjaguin-Landau-Vervey-Overbeek) theory that describes the interparticle interaction and colloidal stability in solution: according to this, replacement of the trivalent citrate with monovalent benzyl mercaptan reduces the energy barrier to interaction, leading to increased interaction and aggregation. These results suggest that the control of surface potential is crucial in the design of stable gold nanoparticle aggregates.


Kim, T.; Lee, K.; Gong, M.-S., and Joo, S.-W.: Control of gold nanoparticle aggregates by manipulation of interparticle interaction. Langmuir, 21, 9524-9528 (2005).

Meanwhile, Arnold Kell goes further by showing that when you get rid of the thiols, stability is reduced still further, in another Langmuir paper. Acyl and alkyl radicals generated photochemically by irradiation of pinacolone or pivalophenone with UV radiation of more than 300 nm wavelength, when conducted in a solution containing monolayer-protected gold nanoparticles (MPNs) prepared with hexanethiolate, dodecanethiolate, and octadecanethiolate ligands, are shown to efficiently liberate the alkylthiolate ligands into the solution as the thioacetyl or alkyl sulfide, respectively. This radical-induced reaction initiates a coalescence of the gold cores to facilitate an increase in core size, and the authors show that this may be used to increase the core size in a controlled manner from 1.7 ± 0.5 nm prior to irradiation to 2.7 ± 0.8 and 5.1 ± 0.9 nm after 90 and 210 min, respectively. No evidence was found for precipitation of the gold nanoparticles. The photoinitiated radical reaction was also found to liberate monolayers from two-dimensional gold surfaces.


Kell, A. J.; Alizadeh, A.; Yang, L., and Workentin, M. S.: Monolayer-protected gold nanoparticle coalescence induced by photogenerated radicals. Langmuir, 21, 9741-9746 (2005).

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