N A N O P R O B E S     E - N E W S

Vol. 7, No. 8          August 21, 2006

Updated: August 21, 2006

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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A Nanogold®-Quenched RNA Molecular Beacon

We have previously described the application of Nanogold® as a quencher for molecular beacons. Conventional molecular beacons are hairpin loops of DNA, conjugated to a fluorescent label at one end and a quencher at the other. The two ends comprise complementary sequences; before binding to the target, the two ends are hybridized, holding the fluorophore and quencher in close proximity and resulting in fluorescence quenching. The loop is complementary to the target sequence: upon binding to the complementary target, the beacon opens and the two ends are forced to separate. The fluorophore and quencher move apart, and as a result, fluorescence is no longer quenched and the fluorescent signal appears.

The quenching ability of a molecule depends upon the degree of overlap between its electronic absorption spectrum and the emission spectrum of the fluorophore. Nanogold absorbs strongly across a wide range of the UV/visible spectrum, and therefore provides very efficient quenching for many fluorophores; the "signal-to-noise ratio," or ratio of fluorescence intensity with the beacon open to that when it is closed, has been found to be up to 2,000 or more with Nanogold, but limited to 100 or less with organic quenchers such as DABCYL (4-([4'-(dimethyl-amino)-phenyl]-azo) benzoic acid).

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 - the distance at which 50% of fluorescence is quenched - is between 6 and 7 nm. How molecular beacons work, and the relationship between gold-fluorophore separation and fluorescence according to the Förster model, are shown below.

[The Hammerhead Ribozyme beacon, and Frster Energy Transfer for Fluorescein and Nanogold (54k)]

Left: The hammerhead ribozyme, showing substrate binding, cleavage and product release; Right: Relationship between fluorophore-Nanogold separation and relative fluorescence intensity showing Förster distance and application to molecular beacons.

Following their recent report on lifetime quenching by 1.5 nm gold particles, Jennings and co-workers describe, in the July issue of Nano Letters, the use of a RNA beacon, using Nanogold as the quencher, in which the fluorescence is activated by cleavage rather than unfolding. Nanometal surface energy transfer (NSET), which describes an energy transfer event from optically excited organic fluorophores to small metal nanoparticles, was used to measure Mg2+-induced conformational changes for a hammerhead ribozyme, and these measurements confirmed using FRET. NSET differs from FRET theoretically in that the distance dependence of quenching is related to the inverse fourth power rather than the inverse of the sixth power of the separation distance (1/R4 rather than 1/R6). These studies are particularly useful because they use optical interactions to gain understanding of the kinetic pathways for this ribozyme at the molecular level.

The hammerhead ribozyme is a naturally occurring RNA motif. It comprises a conserved core loop flanked by three stems, and can catalyze internal strand cleavage. Because it undergoes a controllable conformational change and displays well-characterized cleavage kinetics, it provides a good demonstration of the utility of using NSET as a molecular beacon to follow such a process. At high Mg2+ concentrations, the native ribozyme folds about the scissile bond, bringing stems I and III into close proximity. This initiates cleavage of the substrate strand, releasing the cleaved strands. The authors demonstrated the validity of NSET molecular beacon techniques by measuring the structural changes and cleavage kinetics in a synthetic 40-nt hammerhead complex, consisting of a ribozyme strand and a substrate strand.

To prepare the hammerhead complex, purified RNA strands were deprotected and desalted. Binding of Monomaleimido Nanogold (small size: 6 nmol aliquot) to the free thiol of the ribozyme strand was performed by dissolving 600 pmol of the ribozyme into 20 µL RNase-free ultrapure water; 20 µL of 50mM TCEP was added to the ribozyme solution to regenerate free thiols from any disulfide bonds formed by thiol oxidation. After reacting for 30 minutes at room temperature, the ribozyme solution was brought up to 100 µL and separated from TCEP using successive spin columns. The Nanogold was added to the ribozyme by dissolving a single vial (6 nmol) into 100 µL of water, which was added to the purified ribozyme solution, vortexed, and placed in the refrigerator at 4°C for at least 24 hours to allow coupling. Longer reaction times did not influence the activity of the system.

The Nanogold-labeled Hammerhead ribosome was analyzed by PAGE to confirm labeling and separate the labeled complex. Equimolar amounts of Nanogold-ribozyme (NG-Rib) and FAM-bound substrate strand were combined in a microcentrifuge tube and heated to 95°C for two minutes, allowed to cool to room temperature for ten minutes, then cooled with ice for an additional 10 minutes to allow formation of the Nanogold-hammerhead complex (NG-HHComp). 60 pmol RNA per lane was loaded onto a pre-equilibrated non-denaturing 15% polyacrylamide gel (PA gel) and electrophoresis performed at 4°C. Reactivity of the labeled complex was confirmed by reacting an identical sample with 100 mM Mg2+ for 2 hours before loading onto the gel. The gel was imaged initially using only FAM luminescence, then stained with ethidium bromide and imaged again.

PAGE and optical analyses of Nanogold-labeled ribosome were then correlated in order to derive values for the rate constants for the reaction and its rate determining step, the cleavage reaction. Nanogold-labeled ribosome and FAM-labeled substrate strand were annealed in the 20 mM PBS 0.1 M NaCl pH 6.5 buffer and the solution placed in a clean, dry 50 µL cuvette, which was placed in a Peltier temperature controlled Fluorimeter and equilibrated at 37°C for 10 minutes. 1.2 µL of 5 M MgCl2 was mixed in to start the reaction, bringing the solution in the cuvette to 20 mM Mg2+. Photoluminescence (PL) was monitored continuously for 250 minutes: the data was baseline-subtracted based on the minimum intensity after Mg2+ addition and normalized to pre-Mg22+ intensity to obtain product fraction information. PAGE data were obtained simultaneously for the same sample by removing a 7 µL aliquot (70 pmol hammerhead complex) at specific time points. Each aliquot was quenched by rapid mixing with a denaturing loading buffer containing formamide and 10 mM EDTA, followed by heating to 95°C for 2 minutes, then electrophoresed on a denaturing 15% polyacrylamide gel. The gel was imaged using only the FAM emission only taking care not to saturate CCD pixels of the gel documenter. Line scans of intensity vs. pixel position for each gel lane were background corrected and fit to Gaussian line shapes, and the integrated intensity normalized to a control sample of known concentration (70 pmol of substrate strand) run on the same gel. Gold-labeled ribosome could be observed visually under ambient light as a dark brown band, which was colocalized with the ethidium bromide signal, confirming co-migration and hence labeling.

The kinetic traces derived from the PAGE data and the continuously monitored photoluminescence for the FAM at 518 nm were plotted versus time. Photoluminescence data show that upon rapid mixing of Mg2+ with the Nanogold-ribosome complex, the intensity of the FAM decreases immediately as the complex folds to bring the FAM into closer proximity of the NG. After 1-2 minutes, the intensity of the FAM photoluminescence begins to increase as the cleavage reaction progresses, approaching a maximum value near t = 250 min. After the initial incubation (mixing) period, an increase in the fluorescence of FAM is observed both in the PA gel and in the emission intensity plot, which correlates directly to the kinetics of the cleavage reaction (k2) and can be fit to a first-order reaction for substrate at the limit of low ribozyme concentration. First-order kinetics for substrate binding were confirmed by the linear plot of log(FAM intensity) vs. time: the rate constant of binding was measured to be kc1 = 14.9 ± 1.5 µM-1min-1, and the rate of cleavage for a preannealed Nanogold-labeled ribosome was 0.013 ± 0.001 min-1 in pH 6.5 buffer and at 37°C, which is similar to reports in the literature for these pH conditions.

The authors conclude that using a metallic gold nanoparticle as an acceptor for energy transfer distinguishes NSET from FRET in two significant aspects. Firstly, the change in the distance dependence from 1/R6 to 1/R4 extends the usable distances for measurement; and secondly, the same nanoparticle is able to quench dyes of different emission frequencies, spanning the visible range into the near-infrared. NSET may therefore be very useful for studies in which distances are expected to extend beyond 10 nm, or studies in which multiple dyes need to be quenched. It is likely that a single nanoparticle has the ability to quench several dyes of different energies simultaneously, thus allowing the analysis of several distances in a single experiment. Furthermore, the maintenance of catalytic activity in the presence of Nanogold, which is demonstrated here, is critical for the application of NSET-based molecular beacons or molecular rulers are to biophysical studies. NSET has shown to be advantageous for the measurement of very fast processes that are otherwise difficult to observe.


Reference for gold-labeled DNA beacons:

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

More information:

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Storing Nanogold® Conjugates

Once you have prepared and isolated a Nanogold® conjugate, how do you store it?

Nanogold® is a stable coordination compound. Its surface is protected by coordinated organic ligands, and it is stable to wide ranges of pH - from about pH 4 to pH 10 - and to most commonly used buffer salts. The reactions used for Nanogold conjugation all produce stable covalent linkages which are also resistant to all of these conditions. Monomaleimido Nanogold produces a thioether linkage; Mono-Sulfo-NHS-Nanogold results in an amide link; and the most commonly used cross-linking reaction of Monoamino Nanogold - reaction with an activated ester or carboxylic acid ("peptide" type coupling) also gives an amide link. All three are stable to all the conditions listed above for Nanogold.

The high stability of Nanogold means that any sensitivity of the conjugate to storage conditions is likely to arise either from the properties of the conjugate biomolecule, or from changes in the conjugate biomolecule during the Nanogold labeling reaction, such as the breaking of intrachain disulfide bonds or disruption of hydrophobic interactions reducing the integrity of the biomolecule and making it more vulnerable. In general, the stability of the conjugate biomolecule should therefore predict the stability of the conjugate.

Because of the high affinity of thiols for gold, you should avoid any thiol-containing compounds in solutions used to store Nanogold conjugates. If it is necessary to use thiol-containing compounds during the purification or use of Nanogold conjugates, we strongly receommend keeping the concentration to 1 mM or less and the exposure time to 10 minutes or less.

We recommend the following in order to ensure the stability and integrity of your Nanogold conjugate:

  • For proteins or peptides, maintain a salt concentration of at least 0.1 M. We usually suggest using 0.02 M sodium phosphate buffer containing 0.15 M Sodium chloride for storing conjugates. This may be substituted with any buffer in which the conjugate biomolecule is stable, provided that it does not contain thiols. However, we recommend maintaining a concentration of at least 0.1 M because it helps prevent adsorption to the vial walls, and also helps prevent protein aggregation and precipitation.

  • Store at 4°C (refrigerator) where possible. This will slow any chemical changes.

  • Make sure handling of conjugates prior to storage is as close to sterile as is reasonably possible to minimize the risk of bacterial contamination and subsequent degradation. We recommend filtering conjugate solutions through a 0.22 µm filter immediately before storage to remove any bacteria.

  • If you need to freeze Nanogold conjugates, add a cryoprotectant such as 30 or 50% glycerol. This is because the formation of ice crystals upon freezing can denature proteins and may degrade Nanogold conjugates. Addition of a cryoprotectant will prevent damage due to ice microcrystals during freezing.

If product stability is critical to your application and you know that the structure will be significantly impacted by the labeling procedure, you may wish to consider using an alternative labeling strategy. For example, if you intend to label with Monomaleimido Nanogold, but know that your molecule contains a number of intrachain disulfides which will be reduced during the preparation, you should consider labeling at an amino- site using Mono-Sulfo-NHS-Nanogold (this reaction can often be made selective for more reactive N-terminal amines by conducting labeling at close to pH 7.5), or labeling at a carboxyl group by activation with EDC / Sulfo-NHS followed by reaction with Monoamino Nanogold

What exactly is the thermal stability of Nanogold®? What temperature ranges can I use during immunolabeling or other staining experiments in which specimens are stained with Nanogold conjugates?

Generally, we caution against elevated temperatures during immunolabeling procedures using Nanogold® conjugates and reagents. However, we have found that degradation at elevated temperatures usually requires a complicating factor that increases sensitivity. Specifically, we have found that thermal degradation is more likely to occur under the following conditions:

  • Acidic pH - 5.0 or lower.
  • High ionic strength - 0.3 M or higher.
  • Extensively hydrogen-bonded buffer salt, such as ammonium acetate.

An investigation into the effects of heating using UV/visible spectroscopy to monitor changes in UV/visible absorption revealed that even at close to boiling, changes in the UV/visible spectra of Nanogold were slow, and most of the Nanogold (80%) remained even after several hours.


Hainfeld, J. F., and Furuya, F. R.: Silver-enhancement of Nanogold and undecagold: in Immunogold-Silver Staining: Principles, Methods and Applications; M. A. Hayat (Ed.), CRC Press, Boca Raton, FL, 1995, pp. 71-96.

If you are concerned about thermal sensitivity, we recommend the following precautions:

  • If you plan to use silver enhancement, complete this before embedding. This will reduce thermal sensitivity and you can then use a higher temperature resin.
  • Use a low-temperature embedding medium if you are embedding without silver enhancement.
  • Use the lowest ionic strength and nearest to neutral pH that you can afford.

More information:

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FluoroNanogold: Correlative Fluorescence Microscopy and FESEM

In addition to its use for combined and correlative fluorescence and transmission electron microscopic labeling, the combined fluorescent and gold probe FluoroNanogold is useful for a number of other correlative microscopy methods. In recent articles, we have highlighted its use for correlative fluorescence and scanning electron microscopy, and for correlative optical and X-ray fluorescence microscopy. Our Alexa Fluor®* FluoroNanogold probes now offer superior performance:

  • Increased fluorescence brightness and higher quantum yield.
  • Improved solubility: lower background signal and higher signal-to-noise ratios.
  • Fluorescence remains high and consistent across a wider pH range.

In addition, we now offer both Alexa Fluor®* 488 and Alexa Fluor®* 594 FluoroNanogold, enabling the differentiation of multiple targets using different colored fluorescence. The structure of Alexa Fluor®* 488 FluoroNanogold-Fab', and some examples of the results obtained with it, are shown below.

[Alexa Fluor 488 FluoroNanogold-Fab' and results with it (87k)]

Left: Structure of Alexa Fluor®* 488 FluoroNanogold - Fab' and Streptavidin, showing covalent attachment of components. Center: Fluorescent staining obtained using Alexa Fluor 488 FluoroNanogold as a tertiary probe to label red blood cells. Specimen is a slide from the NOVA Lite ANA HEp-2 test, an indirect immunofluorescent test system for screening anti-nuclear antibodies in human serum, stained using positive pattern control human sera, a Mouse anti-Human secondary antibody, and Alexa Fluor 488 FluoroNanogold tertiary probe. Specimens were washed (PBS, 30 minutes) between each step, then blocked by addition of 7% nonfat dried milk to the tertiary antibody solution (original magnification x 400). Right: Scanning electron micrograph of a peg-like terminal constriction of an Oziroë biflora (plant, Hyacinthaceae) chromosome. The image shows both chromosome topography (secondary electron signal) and hybridized enhanced gold signals (superimposed back-scattered electron signals, yellow) labeling 45S rDNA in the nucleolus organizing region with Alexa Fluor®* 488 FluoroNanogold-Streptavidin (micrograph courtesy of Elizabeth Schröder-Reiter and Gerhard Wanner)

At Microscopy and Microanalysis 2006, Barton and Overall presented their studies, in which correlative immunofluorescence and field emission scanning electron microscopy (FESEM) were used to map microtubule distribution within elongating leaf epidermal cells of Tradescantia virginiana. In interphase plant cells, microtubules form a complex array extending over the inner surface of the plasma membrane, involved in the directed growth of cellulose microfibrils within the cell wall as well as acting as tracks for the intracellular movement of cellular components. A number of methods have been developed to image and investigate the organization and dynamics of microtubule arrays within both live and fixed cells, but the precise organization of individual microtubules within interphase arrays, and their interaction with microtubule associated proteins (MAPs) remains unclear. In order to preserve interphase microtubule arrays within the elongating leaf epidermal cells, a novel stabilization method was developed using the microtubule stabilizing drug, taxol, and the microtubule depolymerizing drug, oryzalin. Colocalization of specific antibodies against MAPs to microtubules was investigated with confocal microscopy and FESEM, using FluoroNanogold secondary antibodies combined with gold enhancement for FESEM visualization.

Using FESEM, individual microtubules were easily resolved. Two spatially distinct subpopulations of microtubules were identified within the arrays: (a) bundled microtubules adjacent to the plasma membrane, located within 25 nm of their neighbors, interlinked by crossbridges and highly aligned within the main axis of each array; and (b) discordant microtubules, lying deeper in the cytoplasm, that crossed the paths of the microtubules lying adjacent to the plasma membrane. The discordant microtubules were randomly aligned throughout the arrays. Comparison of the confocal microscopy and FESEM revealed that individual microtubules, particularly discordant microtubules, were resolved by electron microscopy but were often not visible in the confocal image. Taxol treatment promoted microtubule co-alignment by increasing bundling and decreasing the number of discordant microtubules. After treatment with oryzalin, discordant microtubules were rarely found, and highly aligned, long, single microtubules lay on the plasma membrane. In addition, it is suggested that structural features corresponding to specific microtubule-associated proteins could be identified from the images. This work demonstrates that the higher resolution of electron microscopy provides important insights into antibody localization on the macromolecular scale, and the relationship between microtubules and their associated proteins.


  • Barton, D. A., and Overall, R. L.: Structure and Organisation of Interphase Microtubule Arrays: FESEM Provides a MAP. Microsc. Microanal., 12, (Suppl. 2: Proceedings); Kotula, P.; Marko, M.; Scott, J.-H.; Gauvin, R.; Beniac, D.; Lucas, G.; McKernan, S., and Shields, J. (Eds.); Cambridge University Press, New York, NY; 426CD (2006).

More information:

* Alexa Fluor is a registered trademark of Invitrogen (Molecular Probes), Inc.

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Nanogold® Characterization of MYPT2, a Target Subunit of Myosin Phosphatase

Nanogold-labeled Fab' fragments are the smallest commercially available immunogold probes, and this gives them the ability to penetrate readily into cells and tissues, access restricted antigens, and localize targets with very high resolution. In their recent paper in Cellular Signaling, Okamoto and co-workers use these advantages to determine the distribution and function of MYPT2, a target subunit of myosin phosphatase in heart muscle. In smooth muscle, phosphorylation of light chain of myosin II (MLC) by the Ca2+-calmodulin-dependent myosin light chain kinase (MLCK) initiates contraction by promoting cross-bridge cycling. In striated muscle, phosphorylation of MLC does not induce contraction but may increase Ca2+-sensitivity, i.e. by inducing a shift to lower [Ca2+]i in the forceCa2+ relationship. Since the phosphorylation of MLC is determined by the balance of activities of MLCK and myosin phosphatase (MP), the latter plays an important role in regulating cell functions involving myosin II phosphorylation: for example, inhibition and activation of MP in smooth muscle induces Ca2+-sensitization and Ca2+-desensitization of contractile behavior.

Smooth muscle MP is composed of three subunits: a 38 kDa catalytic subunit of type 1 protein phosphatase delta isoform (PP1cdelta), and two regulatory subunits of 110 kDa (MYPT1) and 20 kDa (M20). MYPT1 has several important functions, including activation and regulation of phosphatase activity. The interaction of MYPT1 with PP1cdelta increases phosphatase activity toward MLC, and phosphorylation of MYPT1 at Thr696 by Rho-kinase inhibits MP activity. The role of MP in cardiac and skeletal muscle is less well characterized. An isoform of MYPT, MYPT2, is predominantly expressed in striated muscle and brain [12]. Because of its structural similarity to MYPT, MYPT2 also functions as a target subunit in MP of striated muscle and brain. In order to facilitate an understanding of the role of MYPT2 in cardiac muscle, the authors have investigated and describe the biochemical properties, regulation, localization and function of MYPT2.

Like MYPT1, MYPT2 has a specific interaction with the catalytic subunit of type 1 phosphatase, delta isoform (PP1cdelta), an interaction of MYPT2 with the small heart-specific MP subunit; interaction of the C-terminal region of MYPT2 with the active form of RhoA phosphorylation by Rho-kinase at an inhibitory site, Thr646 and thiophosphorylation at Thr646 inhibited activity of the MYPT2PP1cdelta complex. MYPT2 activates PP1cdelta activity, using light chains from smooth and cardiac muscle, by reducing Km and increasing kcat. The extent of activation (kcat) was greater than for MYPT1 and could reflect distinct N-terminal sequences in the two MYPT isoforms. Adenovirus-mediated gene transfer of MYPT2 and PP1cdelta reduced the phosphorylation level of cardiac light chains following stimulation with A23187. Overexpression of MYPT2 and PP1cdelta blocked the angiotensin II-induced sarcomere organization in cultured cardiomyocytes.

For immunogold labeling and silver enhancement, 10-µm-thick sections from adult mouse (C57BL6/J) hearts were washed for 10 minutes with Tris-buffered saline (TBS) containing 5% bovine serum albumin (BSA) and 0.02% saponin, then incubated for 48 hours with the primary antibody rabbit anti-LZ antibody at 1 µg/mL in TBS containing 1% BSA and 0.005% saponin. After washing with TBS containing 1% BSA and 0.005% saponin (3 x 1 hour), the sections were incubated with a Nanogold Fab' anti-Rabbit secondary antibody conjugate, diluted 1 : 40 in TBS containing 1% BSA and 0.005% saponin. The sections were then washed again and the sample-bound gold particles enhanced using HQ Silver at 18°C for 12 minutes. The samples were again washed, and postfixed with 0.5% osmium oxide in a buffer containing 100mM cacodylate, pH 7.3. After dehydration by passage through a graded series of ethanol (50%, 70%, 90%, and 100%) and propylene oxide, they were embedded in epoxy resin. Ultrathin sections were cut, stained with uranyl acetate and lead citrate, and then observed in the electron microscope.

Electron microscopy indicated locations of MYPTs, at, or close to, the Z-line, the A band and mitochondria. Similarity of the two MYPT isoforms suggests common enzymatic mechanisms and regulation. Cardiac myosin is a substrate for the MYPT2 holoenzyme, but the Z-line location raises the possibility of other substrates, and pathways or mechanisms that involve MYPT2 could form part of the dynamic role of the Z-lines in intracellular signaling and cardiac function.


  • Okamoto, R.; Kato, T.; Mizoguchi, A.; Takahashi, N.; Nakakuki, T.; Mizutani, H.; Isaka, N.; Imanaka-Yoshida, K.; Kaibuchi, K.; Lu, Z.; Mabuchi, K.; Tao, T.; Hartshorne, D. J.; Nakano, T., and Ito, M.: Characterization and function of MYPT2, a target subunit of myosin phosphatase in heart. Cellular Signalling, 18, 1408-1416 (2006).

Immunogold method:

  • Mizoguchi, A.; Yano, Y.; Hamaguchi, H.; Yanagida, H.; Ide, C.; Zahraoui, A.; Shirataki, H.; Sasaki, T., and Takai, Y.: Localization of Rabphilin-3A on the synaptic vesicle. Biochem. Biophys. Res. Commun., 202, 1235-1243.

More information:

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Custom Syntheses and Distributor Pricing

We are occasionally asked about differences between our prices and those charged by our distributors. Nanoprobes does not stipulate the prices charged by distributors; the only restriction we impose is that distributor prices must not be less than those charged by Nanoprobes for the same items. Apart from this, our distributors are free to set their own prices according to their costs of business and the costs to get the product to their customers.

Costs for products ordered from distributors typically can include both the cost for shipment from Nanoprobes to the distributor, and the cost of shipping from the distributor to the customer. Given current security concerns in many regions of the world, high shipping costs are to be expected to cover rigorous inspection of any packages coming into the country. These costs likely also include any duties, taxes, and other fees on incoming goods. When you order products directly from us, shipping charges are added to the invoice, and in addition, duties and taxes are usually billed to you directly by the shipping company. Customers are always free to seek quotations or order from us directly if they wish, but should be aware of other costs that may be added to the invoice or billed separately.

Custom syntheses are a special situation. Each custom preparation is unique, so we cannot negotiate prices in advance with our distributors. Furthermore, prices for custom syntheses are calculated based only on cost and likelihood of success - because of the unpredictability of the work, there is no calculated profit margin as there is with our catalog products. The price we offer to the person placing the order is our best price, and we cannot negotiate a different price should that customer then decide to place their order through a distributor instead. Therefore, we ask that when you inquire about a custom synthesis, that you complete the transaction through the same channel: i.e. if you first approached us directly, place the order directly, and if you first approached us through a distributor, place the order through them.

More information:

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Other Recent Publications

Willner and group report another novel development in detection and labeling in a recent Angewandte Chemie article: the amplification of biorecognition events based on the catalytic evolution of a biocatalytic label that leads to a readable signal of the biosensing process. Put more simply, and specifically, the authors used targeted ecarin (EC) conjugates. Ecarin catalyzes the transformation of prothrombin (PTh) to thrombin (Th); the thrombin that is produced then acts as a biocatalyst for the hydrolysis of the nonfluorescent substrate bis(p-tosyl-Gly-Pro-Arg)rhodamine-110 to a fluorescent product. The surface concentration of the catalytic EC conjugate is low due to the low coverage by the recognition sites of the analyte units. The EC-mediated conversion of PTh into Th 'evolves' the catalyst for the hydrolysis of 1 and, because of the multiplication inherent in the double catalytic process, leads to the formation of a large number of fluorescent product molecules. This amplification method was applied to the detection of antigenantibody and DNA recognition complexes, and was tested as an analytical procedure for the detection of telomerase in cancer cells. This approach offers the potential for the development of very highly sensitive biodetection methods.


  • Shlyahovsky, B.; Pavlov, V.; Kaganovsky, L., and Willner, I.: Biocatalytic Evolution of a Biocatalyst Marker: Towards the Ultrasensitive Detection of Immunocomplexes and DNA Analysis. Angew. Chem. Int. Ed. Engl., 45, 4815-4819 (2006).

We have previously described a number of novel probes and labeling methods developed by Meyer, Albrecht and colleagues, including multiple labeling using metal particles of different shape or composition, as well as correlative fluorescence and electron microscopy labeling in which different fluorophores were combined with nanoparticles of different metals to label proteins in muscle tissue. This work was reviewed, illustrated and consolidated at Microscopy and Microanalysis 2006, in which the quadruple labeling in platelet whole mount was accomplished using 18 nm colloidal gold particles conjugated to mouse anti-human p-selectin, umbonate ("popcorn-shaped") 18 nm colloidal palladium particles conjugated to human Factor X, hexagonal 15 nm colloidal platinum particles sheep anti-human Factor V IgG, and 5 nm colloidal gold particles conjugated to exogenous human Factor V.


  • Meyer, D. A.; Bleher, R.; Kandela, I. K.; Oliver, J. A., and Albrecht, R. M.: The Development of Alternative Markers for Transmission Electron Microscopy and Correlative Transmission Electon and Light Microscopies. Microsc. Microanal., 12, (Suppl. 2: Proceedings); Kotula, P.; Marko, M.; Scott, J.-H.; Gauvin, R.; Beniac, D.; Lucas, G.; McKernan, S., and Shields, J. (Eds.); Cambridge University Press, New York, NY; 32 (2006).

Also at Microscopy and Microanalysis 2006, Krieger, Bode and group reported on some recent developments in reliable and reproducible digital immunogold analysis for electron microscopy. The authors used the 11 Megapixel CCD camera Morada in combination with the software Solution EMarker for iTEM. The resolving power of this CCD camera combined with the reliability and flexibility of the software combination provides for fast automated analyzing and archiving of digital images of immunogold labeled samples. Overview images of around 10,000 x magnification can easily be analyzed, as well as detailed views of tissues. This combination can be applied to most transmission electron microscopy platforms. It may be used not only on standardized labeled samples with single labels, but for the assay of both double-labeling and multi-labeling probes with the same simplicity and reliability.


  • Krieger, J.; Bartels, M.; Chen, W.-J., and Bode, M.: Recent Developments in Reliable and Reproducible Digital Immunogold Analysis in Electron Microscopy. Microsc. Microanal., 12, (Suppl. 2: Proceedings); Kotula, P.; Marko, M.; Scott, J.-H.; Gauvin, R.; Beniac, D.; Lucas, G.; McKernan, S., and Shields, J. (Eds.); Cambridge University Press, New York, NY; 1674CD (2006).

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