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Updated: May 4, 2005

N A N O P R O B E S     E - N E W S

Vol. 6, No. 5          May 4, 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)-Nanogold® Identifies His Tag Locations on Viral Capsids

Nitrilotriacetic acid (NTA)-Nickel (II) Nanogold is a new type of gold probe. Instead of an antibody, it is targeted by a nickel chelate that has a high affinity for polyhistidine tags. This new way to target Nanogold® gives it several important advantages:

[NTA-Ni(II)-Nanogold Structure, binding and STEM (46k)]

left: Structure of Ni-NTA-Nanogold® showing interaction with a His-tagged protein. Inset shows comparison between labeling resolution using Fab'-Nanogold and NTA-Ni(II)-Nanogold; 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).

Advantages of this probe include:

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

  • Its small size means that NTA-Ni(II)-Nanogold is also better able to penetrate into specimens and access restricted sites within them.

  • NTA-Ni(II)-Nanogold is made with a modified gold particle, stabilized slightly differently to that used for other Nanogold reagents. This particle has very high solubility and stability, and at 1.8 nm in size, is also readily visualized by electron microscopy.

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

Polyhistidine tags are may be engineered easily into a wide variety of proteins, and are often used to separate an expressed protein by means of chromatography over Ni(II) chelate resins. This new probe therefore has a wide range of potential applications:

  • High-resolution localization of specific sites in protein complexes and other supramolecular structures.

  • Cytochemical labeling, using polyhistidine-tagged recombinant primary antibodies or probes.

  • Targeting of polyhistidine-tagged green fluorescent protein (GFP) fusion proteins, thus providing a novel method for combined fluorescent and gold labeling.

Chatterji and co-workers illustrated the resolution of this probe by using it to selectively label Cowpea Mosaic Virus (CPMV) capsids modified with 6x His sequences inserted at different locations on the capsid surface; these experiments were part of a study to show control of the electrostatic properties of the capsid through protonation of the histidine residues.

6x His peptide was expressed at five different sites on the capsid surface known to tolerate genetic modifications, using well-established oligonucleotide directed mutagenesis techniques: two (vaBC-H6 and vaCC-H6) in which the His tag sequence is displayed in the loops of the small subunit; the vcEFH6 mutant with the 6x His peptide presented in the betaE-betaF loop of the large subunit of the capsid; and another two mutants, vaCTr-H6 and vaCT-H6, with the His tag presented as linear epitopes on the C-terminus of the small subunit. The mutants were purified by Ni(II) chelate chromatograpy, but bound to the column to differing extents, indicating differences in accessibility of the 6x His tag. To demonstrate the influence of the 6x His tag on the surface properties of viral capsids as a function of pH, the purified mutants were chromatographed using a mono Q anion exchange column. Changing the pH from 5.5 (the isoelectric point of the wild-type virus) to 9.0 did not change the retention time of the native virus significantly, but produced a dramatic change in the binding affinity of the HIS mutants due to deprotonation of the histidine residues (pKa 6.7-7.1).

The specific addressability of the His-tagged mutants was verified by labeling with NTA-Ni(II)-Nanogold, which targets His sequences. Cryo-electron microscopy combined with three-dimensional reconstruction techniques were used to determine the structure of gold-labeled vcEF-H6 mutant. 10 nmol of NTA-Ni(II)-Nanogold, dissolved in 90 microliters of 0.1M sodium phosphate, pH 7.5, was mixed with 10 microliters (100 micrograms) of vcEF-H6 mutant and the reaction was incubated for one hour at room temperature, then overnight at 40°C. Excess gold was then removed by gel filtration (Superose 6 column). Fractions containing the gold-labeled virus were collected, concentrated to 1 mg/ml and used for cryo electron microscopy. Samples (4 microliters) were applied to glow-discharged Quantifoil R 2/1 grid, quickly blotted with filter paper, and plunged into liquid nitrogen-cooled ethane to embed the virus particles in a thin layer of vitreous ice. The virus-gold complexes were then examined using a Philips CM120 transmission electron microscope operating at 100kV maintained at near liquid nitrogen temperature. Micrographs were recorded at an instrument nominal magnification setting of 60,000X, and objective lens defocus of 0.94 m.

A total number of 1072 full particles were manually selected using the program BOXER. Gold particles were clearly identifiable under the microscope, allowing a selection of only those particles with strong density spots corresponding to gold: 900 were used for final calculations. The model-based procedures were used for all subsequent orientation and phase origin refinement. Image processing was carried out using the program SPIDER imposing completed icosahedral (532) symmetry for density map calculations. A threshold value was determined that included 100 % of the expected volume for the CPMV-gold complex capsids and CPMV WT capsids. Spherically averaged radial density profiles were calculated for both maps, normalized, and scaled to match the fit between both profiles; a difference map was then obtained by subtraction. The atomic models of CPMV and gold particles were fit into the electron density maps manually by using O; images were rendered using Bobscript and Chimera.

Structural comparison between WT virus and the gold-labeled vcEF-H6 capsids revealed additional electron density located on the virus surface, attributable to the attached gold. In the final image, the structure of the wild-type virus may be identified, with additional density, mainly around the 5-fold axes; by subtraction of the density for the wild-type particles from the density associated with the gold-labeled particles, the location of the gold was clearly defined. The density associated with the gold is located on the particle surface, just on top of the betaE-betaF loop where the His tag was inserted; this gold density was elongated, perhaps as a result of either flexibility of the histidine peptide, or averaging of the position of the gold particles binding to multiple histidine residues.

Reference:

Chatterji, A.; Ochoa, W. F.; Ueno, T.; Lin T., and Johnson, J. E.: A virus-based nanoblock with tunable electrostatic properties. Nano Lett., 5, 597-602 (2005).

Abstract (courtesy of PubMed (Medline)).

The nature of the binding interaction is different from that of antibodies or proteins such as streptavidin, and different concentrations, reagents and conditions may be appropriate for blocking and for generating the highest labeling with lowest background interaction. Because binding occurs by coordination of electronegative atoms, usually aromatic nitrogen, to the nickel (II) ion, other aromatic nitrogens, such as other histidine residues, may also bind. This may be prevented by treatments that reduce this interaction. The following steps may be useful:

  • Wash with a buffer containing imidazole. Imidazole is the active coordinating group in histidine; treatment with imidazole will displace isolated single histidine binding, but will not overcome the chelate effect and stronger binding of NTA with polyhistidines, and therefore will disturb polyhistidine binding much less. Try increasing the imidazole concentration from 10 mM progressively to 200 mM until background is controlled to your satisfaction.

  • Increase the ionic strength of the solution. The nitrilotriacetic acid moiety is negatively charged, and increasing the ionic strength will prevent it binding by ion-pairing with positively charged regions of a target. Try 300 mM NaCl, and, if your biomolecule can tolerate it, increase to 1.0 M if necessary.

  • Other possible factors include:

    • hydrophobic interactions. Try adding detergents to reduce these interactions: start with 0.05 % Tween-20, and increase to 0.1 % if necessary.

    • pH: it may be helpful to vary the pH to find an optimum pH at which charge interactions such as those mentioned above are reduced.

    • Transition metals may also promote interactions between the NTA group and electron donating groups in your specimen. Remove these by washing with 0.05 M disodium ethylene diamine tetraacetic acid (EDTA).

    • Interaction of thiols (sulfhydryls) with gold. Thiols have a strong affinity for gold, and thiols in your specimen may bind to any gold particle species, including Ni-NTA-Nanogold. Avoid the use of reducing buffers or preservatives containing thiols, such as dithiothreitol (DTT), mercaptoethanol, or mercaptoethylamine hydrochloride(MEA). If your specimen contains exposed thiols, they may be blocked with N-ethylmaleimide.

If you are seeing background signal appearing after silver or gold enhancement, a number of methods are available for stopping these reactions and preventing further reaction after the desired end-point by reagents that have diffused into specimens.

NTA-Ni(II)-Nanogold contains multiple NTA-Ni(II) groups, necessary for the best overall combination of labeling selectivity, density and sensitivity. However, it can interact with polyhistidine tags on several protein molecules simultaneously, and hence may aggregate proteins, or perturb the formation of protein complexes in solution. To avoid this, use a ratio of reagent to protein such that the stoichiometry reduces or eliminates this possibility. For example, if your protein has only one polyhistidine tag, use an excess of the Ni-NTA-Nanogold reagent to guard against the possibility of multiple interactions. You can also avoid crosslinking by carefully selecting when to add the reagent, for example after complex assembly.

Original reference:

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).

More information:

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Fluorescence Quenching, FluoroNanogold and Molecular Beacons

What are the interactions between fluorescent labels and gold particles, and how do they affect labeling applications? In fact, at short distances, gold particles effectively quench fluorescence. This is why, if you wish to prepare combined fluorescent and gold probes for correlative labeling, you need to position the two labels sufficiently far apart that this quenching is reduced enough to see the fluorescence. For example, in our FluoroNanogold conjugates, the fluorescent label and Nanogold® particle are attached separately. Over distances in the nanometer range, we have found that resonance energy transfer (Förster mechanism) predicts fluorescence behavior reasonably well, and we have described this in our 1998 paper in Microscopy Research and Technique:

Powell, R. D.; Halsey, C. M. R., and Hainfeld, J. F.: Combined fluorescent and gold immunoprobes: Reagents and methods for correlative light and electron microscopy. Microsc. Res. Tech., 42, 2-12 (1998).

For Nanogold and fluorescein, the Förster distance is between 6 and 7 nm. The structure of fluorescein Fab'-FluoroNanogold and the Förster relationship between gold-fluorophore separation and fluorescence are shown below.

[Structure of Fab'-FluoroNanogold, and Frster Energy Transfer for Fluorescein and Nanogold (42k)]

left: Structure of fluorescein Fab'-FluoroNanogold, showing the separate attachment of the Nanogold and fluorescein labels; right: Schematic showing effect of fluorophore-Nanogold separation on fluorescence, Förster distance, and application to molecular beacons.

Although this presents an obstacle to preparing smaller FluoroNanogold probes such as peptides or substrate analogs, it enables other important applications. Molecular beacons are loops of DNA labeled with both a fluorophore and a quencher. Unbound, the two are close together, and fluorescence is quenched: upon target binding, the loop opens, the quencher and fluorophore move apart, quenching is removed, and the fluorescent signal appears. This property makes molecular beacons very useful for homogeneous systems, and for fluorescent labeling in situations where it is difficult to remove unbound probes.

Because of its strong UV-visible absorption across a wide wavelength range, Nanogold is highly effective as a quencher. Dubertret and co-workers, using Monomaleimido Nanogold to label hairpin loop beacons, found that the Nanogold particle is significantly more effective than the conventional DABCYL quencher, yielding signal-to-noise ratios (i.e. the fluorescence intensity for open : closed configuration) of up to 1,000 or higher:

Dubertret, B., Calame, M., and Libchaber, A.; Single-mismatch detection using gold-quenched fluorescent oligonucleotides. Nat. Biotechnol., 19, 365-370 (2001).

Nanoprobes is currently developing new technology for this application.

However, Förster energy transfer is only one of several mechanisms for fluorescence quenching. Especially at shorter distances, others may contribute and produce even higher quenching, resulting in even higher signal-to-noise ratios. Dulkeith and co-workers previously described the effect on radiative and non-radiative fluorescence lifetimes for systems in which fluorophores were linked to 1 to 30 nm metal nanoparticles; they found both an increase in the radiative lifetime and a decrease in the non-radiative lifetime, implying fluorescence quenching greater than predicted by Frster theory alone, and with a gold-fluorophore separation of 1 nm observed about 99.8 % quenching even with 1 nm particles. At those small distances, the large fluorescence quenching efficiency is due to two effects of equal importance: (a) increased nonradiative rate of the molecules by the gold particles due to energy transfer, and (b) the radiative rate of the molecules is decreased because the molecular dipole and the dipole induced on the gold particle radiate out of phase if the molecules are oriented tangentially to the gold surface.

Now, this group reports a study investigating the effect of separation upon quenching in more detail. They used time-resolved photoluminescence studies to determine the radiative and non-radiative rates for Cy5 fluorophores spaced between 2 and 16 nm from attached 6 nm gold particles using single-stranded DNA spacers. They find that the distance dependent quantum efficiency is almost exclusively governed by the radiative rate, and the energy transfer is of minor importance. This may be explained by the fact that the photoluminescence emission wavelength of Cy5 is red shifted with respect to the gold nanoparticle plasmon resonance at about 520 nm: suppression of the radiative rate is due to a phase difference between the molecular dipole and the gold nanoparticle plasmon. Energy uptake of a driven resonator like the gold nanoparticle plasmon is known to relax more quickly with detuning than does its phase lag.

These results suggest that where the gold plasmon resonance plays a role in quenching, it would be greater than predicted by Förster theory alone, but the increase may not be as great as suggested by some other models such as the Gersten-Nitzan. Given the enhanced quenching at shorter distances, an optimum configuration of gold nanoparticle and fluorophore could yield beacons with a higher signal-to-noise value than predicted by either approach.

References:

  • Dulkeith, E.; Ringler, M.; Klar, T. A.; Feldmann, J.; Munoz Javier, A., and Parak, W. J.: Gold nanoparticles quench fluorescence by phase induced radiative rate suppression. Nano Lett., 5, 585-589 (2005).

  • Dulkeith, E.; Morteani, A. C.; Niedereichholz, T.; Klar, T. A.; Feldmann, J.; Levi, S. A.; van Veggel, F. C. J. M.; Reinhoudt, D. N.; Enschede, A. E.; Möller, M., and Gittins, D. I.: Fluorescence Quenching of Dye Molecules near Gold Nanoparticles: Radiative and Nonradiative Effects. Phys. Rev. Lett., 89, 203002 (2002).

More information:

*Alexa Fluor is a trademark of Molecular Probes, Inc.

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Nanogold® Optical Sensors for Chemical Warfare Agents and Pesticides

Molecular Beacons are not the only types of probes that are enabled by the ability to modify fluorescence. As Simonian and co-workers report in this month's Analytica Chimica Acta, the principle can also be used for the detection of an enzyme substrate, by measuring the change in fluorescence when a fluorescently labeled substrate decoy is displaced from a Nanogold®-labeled enzyme. The twist here is that fluorescence is actually enhanced upon binding, suggesting that Nanogold can actually amplify the fluorescence of a fluorophore at distances of 10 to 40 nm.

Neurotoxic organophosphates (OP) have found widespread use for insect control, and there is also an increasing threat that OP-based chemical warfare agents will be used in both ground based warfare and terrorist attacks. These trends necessitate the development of simple and specific methods for the discriminative detection of very low quantities of organophosphate neurotoxins. The enzymatic hydrolysis of organophosphate neurotoxins by organophosphate hydrolase (OPH) generates two protons in each hydrolytic turnover through reactions in which PX bonds are cleaved, and this reaction previously provided the basis for a potentiometric biosensor. However, practical use of this method is limited.

Measuring the change in a fluorescence signal is simpler. Therefore, to test the feasibility of this approach, OPH-gold nanoparticle conjugates were prepared using Mono-Sulfo-NHS-Nanogold and Monomaleimido Nanogold. Native OPH was isolated from a recombinant Escherichia coli strain: the crystal structure shows that this molecule contains six primary amines on each OPH monomer, in the form of lysine residues, and only two sulfhydryl groups, in the form of cysteine residues, on the OPH molecule. The OPH/mono-Sulfo-NHSNanogold conjugate was prepared by dissolving Mono-Sulfo-NHS-Nanogold in 1 ml deionized water, mixing with the protein (1 mg/ml final concentration) and reacting overnight at pH 7.58, at 4°C. Sufficient reagent was supplied to label 6 nmol of amine sites. Unbound Nanogold particles were removed by ultrafiltration, using Millipore Microcon YM-10 tubes (MW cutoff 10,000). The extent of labeling was calculated from the UVvisible absorption spectrum of the conjugate. Labeling with Monomaleimido Nanogold was also conducted according to the product instructions. Nanogold conjugates were stored in 0.02M sodium phosphate buffer with 150 mM sodium chloride.

Detection reactions were conducted in solution, using a spectrophotometer to monitor enzyme activity and a spectrofluorimeter for fluorescence measurements. A stock solution of the fluorescent substrate decoy, 7-Hydroxy-9H-(1,3-dichloro-9,9-dimethylacridin-2-one (DDAO phosphate) fluorophore was prepared at concentrations of 10-6 to 3 10-6 M in deionized water. The fluorescence intensity (IF1) of DDAO was measured and used as a background signal level. OPH-Nanogold conjugate was added and intensity of fluorescence of the conjugatedecoy complex (IF2) measured again. Paraoxon was then added in different concentrations, fluorescence intensities (IF3) were measured, and relative fluorescence intensity change was calculated. Upon incubation of the OPH-Nanogold with the fluorescent enzyme inhibitor or decoy, the fluorescence intensity of the decoy was found to increase significantly. Introduction of different paraoxon concentrations resulted in a decrease in fluorescence proportional to the paraoxon concentration added; the greatest sensitivity to paraoxon was obtained when decoys and OPHgold nanoparticle conjugates were present at near equimolar levels.

Reference:

Simonian, A. L.; Good, T. A.; Wang, S.-S., and Wild, J. R.: Nanoparticle-based optical biosensors for the direct detection of organophosphate chemical warfare agents and pesticides. Anal. Chim. Acta, 534, 69-77 (2005).

More information:

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How to Label Aromatic Amines

If you plan to use Mono-Sulfo-Nanogold® to label aromatic amines, it is important to note that the reactivity of these is different to that of aliphatic amines, and modify the conditions accordingly. Aromatic amines are much less reactive towards NHS esters than aliphatic amines. Chemically, this is because reaction proceeds via nucleophilic attack of the lone pair of electrons on the amino- nitrogen atom on the Sulfo-NHS-ester, displacing the NHS group to form an amide bond. However, in aromatic amines, this lone pair is partially conjugated with the aromatic pi-system, and much less available for reaction. Therefore, reaction with NHS esters is more difficult.

As a result, while aliphatic amines will react if the pH is simply kept high enough to prevent protonation of the amine to the ammonium ion R-NH3+, aromatic amines must be partially deprotonated so that there is a contribution from the Ar-NH- form in order to facilitate reaction. This requires more forcing conditions, and usually a much stronger base.

To conduct such labeling reactions, we suggest the following strategy:

  • Conduct the reaction in an aprotic organic solvent rather than in water, aqueous buffers or alcohols. Dimethylsulfoxide (DMSO) or N,N-dimethylacetamide (DMA) make good choices because they are good solvents for Nanogold and are often compatible with water-soluble molecules.

  • You will need a strong, hindered base. Triethylamine is a good choice to try first: add a couple of drops of triethylamine to 1 mL of DMSO or DMA solution of the compound you wish to label, then add the Mono-Sulfo-Nanogold, and stir at room temperature for one hour. Use an excess of a smaller molecule, but use an excess of Nanogold if you are labeling a molecule that is larger than Nanogold.

  • If the reaction is complete, you can transfer the reaction mixture to aqueous or alcoholic solution, then separate the conjugate from unconjugated Nanogold or any excess of the molecule you are labeling using gel filtration or other usual methods.

  • Test the conjugate for reactivity. We recommend either:

    • A blot test, in which the target is spotted in serial dilutions onto a nitrocellulose membrane, then detected by application of the Nanogold conjugate followed by silver enhancement or gold enhancement blot, or;

    • If the probe target is larger than the probe, you can mix the target with the labeled probe in solution, then separate the mixture using a size fractionation column such as Pharmacia Superose-6 or Superose-12. If binding occurs, a large labeled species will elute, and you will observe little or no peak for free Nanogold; if it does not, then the target and labeled probe will elute separately.

  • If this does not produce labeling, consider a stronger base, such as LHMDS (lithium hexamethyldisilylamine) or lithium diisopropylamide. However, these should be used cautiously, since they are much stronger than triethylamine and may produce undesirable reactions elsewhere in your molecule.

If this approach is unsuccessful or other groups in your molecule prevent it, you may need to use the aromatic amine as a starting point to attach a more reactive group. An example of such a reaction is shown below in Scheme 1: you can use either a very activated ester such as a tosyl ester, or even an acid halide, to react with the amine and attach a new functional group that is easier to work with, in this case an ester which you can subsequently hydrolyze, activate to an NHS ester or other reactive species, and label using Monoamino Nanogold

[Aromatic amine labeling (8k)]

Schematic showing reactions for functionalizing and labeling aromatic amines.

If you need help in finding a suitable synthesis, we would be glad to advise. Contact us with your questions.

More information:

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See you in Hawaii!

Microscopy and Microanalysis 2005 promises to be the largest Microscopy and Microanalysis meeting yet. Together with our collaborators, we are contributing four papers to the effort:

  • Vishwas Joshi's paper on "High Z Metal Carbonyls for Imaging and Microspectroscopy" will be presented at symposium on Confocal Microscopy, at 3:00 pm in room 317B, on Tuesday, August 2.

  • Rick Powell and colleagues will be reporting their progress "Towards Bigger Nanogold: Preparation of Covalent 3nm Gold-Fab' Probes" in Symposium on Biological Specimen Preparation. This is included in the poster session beginning at 3:30 pm in the Exhibit Hall on Tuesday, August 2.

  • Jim Hainfeld and group, in a second poster, will be reporting on "In Vivo Vascular Casting" as part of Symposium on Vascular Corrosion Casting. See it at the poster session beginning at 3:30 pm in the Exhibit Hall on Wednesday, August 3.

  • Jim Hainfeld will also be describing larger NTA-Ni(II) gold probes, in "5 nm Gold-Ni-NTA Binds His Tags." This will be part of the symposium titled "From the Static to the Dynamic: Correlative and High Resolution Biological Imaging and Labeling." This presentation will also be in the poster session, beginning at 3:30 pm in the Exhibit Hall, Wednesday, August 3.

Are you interested in working at Nanoprobes, either as a Microscopy Technician or biotechnology Research Associate? We are looking for someone to help us with microscopy, and for two others to join our research and development team. If you are interested, please check our employment page for full details.

More information:

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Other Recent Publications

Itamar Willner and group describe another novel biosensor application in their recent Nano Letters article. The hydrolysis of acetylthiocholine by acetylcholine esterase, AChE, yields a reducing agent, thiocholine, that stimulates the catalytic enlargement of 2-3 gold nanoparticle seeds in the presence of tetrachloroaurate, [AuCl4]-. This reductive enlargement of the gold nanoparticles is controlled by the concentration of the substrate and by the activity of the enzyme, but it is inhibited by 1,5-bis(4-allyldimethylammoniumphenyl)-pentane-3-one dibromide, or by diethyl p-nitrophenyl phosphate (paraoxon). This inhibition enables a colorimetric test for the presence of these AChE inhibitors, based on the change in the UV/visible absorbance spectra of the nanoparticles; this assay was also developed on glass supports, using glass slides functionalized with nanoparticles.

Reference:

Pavlov, V.; Xiao, Y., and Willner, I.: Inhibition of the acetycholine esterase-stimulated growth of au nanoparticles: nanotechnology-based sensing of nerve gases. Nano Lett., 5, 649-653 (2005).

The streak of references citing our negative stain reagents, NanoVan and Nano-W, is continued this time by Norcum and colleagues in their studies of the bacteriophage T4 primosome assembly. Helicase and primase are enzymes that participate in primosome assembly during replication of DNA; in bacteriophage T4, they are separate polypeptides, for which little structural information is available and whose mechanism of association within the primosome is not yet understood. Equimolar ratios of protein were allowed to assemble into complexes in the presence of a 45-mer ssDNA substrate and MgATPgammma S, then adsorbed to thin carbon films and embedded in NanoVan (methylamine vanadate). Transmission electron micrographs were taken with minimum dose focusing at a magnification of 63,000. Three-dimensional structural information was obtained by reconstruction from the electron microscopic images, and structures were calculated for complexes of each of these proteins. Both the helicase (gp41) and primase (gp61) complexes are asymmetric hexagonal rings. The gp41 structure suggests two distinct forms that have been termed "open" and "closed." The gp61 structure is clearly a six-membered ring, which may be a trimer of dimers or a traditional hexamer of monomers. This structure provides conclusive evidence for an oligomeric primase-to-ssDNA stoichiometry of 6:1.

Reference:

Norcum, M. T.; Warrington, J. A.; Spiering, M. M.; Ishmael, F. T.; Trakselis, M. A, and Benkovic, S. J.: Architecture of the bacteriophage T4 primosome: electron microscopy studies of helicase (gp41) and primase (gp61). Proc. Natl. Acad. Sci. USA,, 102, 3623-3626 (2005).

Want to preserve your immunofluorescence longer in non-aqueous samples? Espada and group describe a solution in the most recent issue of Histochemistry and Cell Biology. An aqueous mounting medium is generally assumed to be necessary for the preservation of immunofluorescent-labeled specimens, and polyvinyl alcohol-based solutions are the most frequently used mounting media; however, the quality and intensity of the fluorescence signal in most immunolabeled preparations slowly diminish with time after aqueous mounting, and become unsuitable for examination. In this study, dehydration through 70% and 100% ethanol, clearing in xylene (each step lasting 1020 s) followed by mounting in 330 DePeX (a non-fluorescing solution of polystyrene in xylene; Serva) was used, and compared with both direct mounting of wet preparations with 25% glycerol in PBS, and mounting with the hydrophilic medium Mowiol, prepared as 8% Mowiol 4088 in glycerol-PBS (1:3, v/v). While both the latter preparations eventually showed degradation of the fluorescence signal, specimens mounted in 330 DePeX showed comparable fluorescence even after storage for one year. DePeX mounting is permanent but not irreversible. The glass coverslip of the preparations is easy to remove after immersion in xylene for several days and, after rehydration, samples can be subjected to further processing.

Reference:

Espada J.; Juarranz A.; Galaz S.; Canete M.; Villanueva A.; Pacheco M., and Stockert, J. C.: Non-aqueous permanent mounting for immunofluorescence microscopy. Histochem. Cell Biol., 123, 329-334 (2005).

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