N A N O P R O B E S     E - N E W S

Vol. 9, No. 8          August 31, 2008

An important aspect of the study of endocytosis is tracing the participant entities as they move through the process, and this requires labels that can bind to the species involved without changing their size or mobility properties significantly: this requires small molecules. While a number of probes exist for this at the light level, probes for electron microscopic tracing are more problematic. Because they are difficult to functionalize and require additional macromolecules for stabilization, colloidal gold probes are not well suited to this task; however, these requirements are met by the smaller Charged Nanogold® probes. Nanogold can be prepared with positively or negatively ionizing surface functionalization, enabling targeting to negatively charged and positively charges species; for example, Positively Charged Nanogold can be used to label oligonucleotides for scanning transmission electron microscope (STEM) observation and as a potential method for making conductive nanowires.

Onelli and co-workers have continued their earlier work in their recent paper in the Journal of Experimental Botany, in which they used Positively Charged Nanogold and Negatively Charged Nanogold as probes to trace the internalization of plasma membrane (PM) domains carrying negatively charged residues at the ultrastructural level; these experiments were combined with immunogold labeling experiments with 20 nm gold to confirm the nature of compartments.

Tobacco cells were grown in a culture chamber at 25±2°C with continuous lighting for 7 days, using MS medium containing 0.1 mg/mL myo-inositol, 0.5 mg/mL thiamine chlorohydrate, 0.5 mg/mL glycine, 0.5 mg mg/mL nicotinic acid, 0.5 mg/mL pyridoxine, 8.84 mg/mL 2,4-dichlorophenoxyacetic acid, and 20 g/L sucrose was used. Protoplasts (8 g) were obtained by digesting the cell walls with 20 mL of digestion solution (0.45 M pH, 5.5 mannitol, 1.5% cellulase, 0.15% pectinase and protease inhibitors) for 3 hours at room temperature. Protoplasts were then filtered (100 µm mesh), centrifuged at 380 x g for 3 minutes, then resuspended in 10 mL culture medium with 0.45 M mannitol and protease inhibitors for 15 minutes before Nanogold addition.

For time-course experiments, protoplasts were incubated with 30 nmol of Positively Charged Nanogold, resuspended into 200 µL distilled water (MilliQ grade). Samples were taken at 5, 15, 30, 45, and 120 minutes. Protoplasts were fixed overnight at 4°C by direct addition of formaldehyde and glutaraldehyde to give final concentrations of 2% and 0.2% respectively, or with 2% glutaraldehyde. Cell viability was checked in all experiments by staining the protoplasts with fluorescein diacetate (FDA) before fixation; samples were observed by fluorescence microscopy. After fixation, specimens were centrifuged at 380/400 x g for 3 minutes, then resuspended with an equal volume of 2% low melting agarose in HEPES buffer (50 mM HEPES, 1 mM MgCl₂, 5 mM EGTA) to form solidified drops, which were then rinsed for 1 hour in HEPES buffer. Protoplasts were treated with ammonium chloride 50 mM in HEPES for 30 minutes at room temperature, then dehydrated with increasing concentrations of methanol. Infiltration and polymerization were carried out at low temperature (20°C) with a CS-Auto cryo-substitution apparatus, according to the protocols furnished with LR GOLD resin. 80 nm ultra-thin sections, obtained using an Ultracut E microtome, were collected on nickel grids.

Positively and Negatively Charged Nanogold were enhanced with HQ silver for 2 minutes in time-course and control experiments and for 1 minute for immunogold labeling. Sections were then stained with 3% uranyl-acetate for 20 minutes and observed with a Jeol SX100 electron microscope at 80 kV. Quantitation of Positively and Negatively Charged Nanogold in different compartments was performed by counting the number of gold particles observed at a fixed magnification of 20,000 of the electron microscope. Total numbers obtained for different labeled protoplast profiles were used to calculate the percentage of Nanogold internalized in different membranous compartments. Standard errors (SE) were calculated for all the experiments using Positively or Negatively Charged Nanogold for graphs. ANOVA test was used to evaluate the difference between single compartments under different experimental conditions and between different incubation times in the time-course and pulse experiments. Tukeys Post Hoc test of Honestly Significant Difference (HSD) was used to assess the significance of each comparison.

For immunogold experiments, after silver enhancement of Positively Charged Nanogold (1 minute), sections were saturated with 2% bovine serum albumin (BSA) in Tris-buffered saline (TBS), then incubated with SYP21 primary antibody (1 : 200). After rinses in 0.1% BSA/TBS, sections were incubated with 20 nm gold anti-rabbit secondary antibody. Finally, specimens were stained with 3% uranyl-acetate for 20 minutes, and observed using a Jeol SX100 electron microscope at 80 kV.

The probes revealed distinct endocytic pathways within tobacco protoplasts, and allowed detailed characterization of the morphology of the organelles involved in endocytosis. Putative early endosomes with a tubulo-vesicular structure, similar to that observed in animal cells, were described. A new compartment, characterized by interconnected vesicles, was identified as a late endosome using the Arabidopsis anti-syntaxin family Syp-21 antibody and immunogold labeling. Endocytosis dissection using Brefeldin A (BFA), pulse chase, temperature- and energy-dependent experiments combined with quantitative analysis of Nanogold particles in different compartments, suggested that recycling to the PM predominated with respect to degradation. Further experiments using ikarugamycin (IKA), an inhibitor of clathrin-dependent endocytosis, and Negatively Charged Nanogold confirmed that distinct endocytic pathways coexist in tobacco protoplasts.

Reference:

• Onelli, E.; Prescianotto-Baschong, C.; Caccianiga, M., and Moscatelli, A.: Clathrin-dependent and independent endocytic pathways in tobacco protoplasts revealed by labelling with charged nanogold. J. Exp. Bot., 59, 3051-3068 (2008).
  • Abstract (courtesy of Oxford Journals).
  • reprint (PDF) (courtesy of Oxford Journals).

More information:

• Catalog information: Nanogold reagents
• Positively Charged Nanogold: Product information and instructions
• Negatively Charged Nanogold: Product information and instructions
• Technical help: Nanogold reagents
• References: Charged Nanogold

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Customize your Silver Enhancement

How can I change the reaction rate? Can I dilute the silver enhancer?

Our silver enhancement reagents can certainly be diluted, but this may not be the best way to modify their reaction rate, as we explain below. Dilution is actually more appropriate if you wish to limit the degree of enhancement. In our experience, the best method for slowing down the silver enhancement reaction is to reduce the pH of the silver enhancement reaction. HQ Silver has a pH buffered close to 6.8, so the addition of a drop of buffer with a lower pH, such as 5.5, will slow down the reaction. Make sure to use a buffer which does not precipitate silver: citrate buffer is a good choice, and usually helps keep background low as well. One drop of sodium citrate buffer, pH 5.5, and one extra drop of solution B should slow down the reaction rate. Lower pH buffer may be necessary if the reaction is not slowed down sufficiently.

LI Silver already has a relatively low pH (and does in fact develop more slowly than HQ Silver). However, if you wish to slow it down further, adding or increasing the amount of thickening agent is a good method, since this slows down the diffusion of ions through the solution. If you wish to slow down the performance of LI Silver, you can add a thickening agent such as 50% gum arabic, or a high MW polyethylene glycol solution (or, if you have HQ Silver as well, you can use some of the Solution B from this). If you want a thicker, slower HQ Silver, use more Solution B ("Moderator").

If you wish to dilute our HQ Silver silver enhancer, you should dilute the two components that contain the silver source and the reducing agent - Solutions A ("Initiator") and C ("Enhancer") - equally. Since the mixed reagent is buffered, the pH will not change much; therefore, the reaction and results will be similar.

Component B - the moderator - actually has twice the concentration of thickening agent as the mixed final solution, and does not contain either silver or reducing agent. Therefore, a 1 : 1 mixture of Solution B with deionized water is actually just about the best diluent, since mixtures prepared with this will have the same viscosity properties as regular HQ Silver. Alternatively (or if you have used all the Solution B), a 30% solution of gum arabic in deionized water will provide approximately the same effect. If you wish to dilute LI Silver, you should dilute both components equally in deionized water: the reagent is buffered, so this will not change the pH much.

Our silver enhancement reagents are already at the optimum concentration for many applications, and we not quantitatively investigated the relationship between concentration and development rate for significant variations in concentration. In the absence of thickening agent, our experience is that dilution will slow down the development rate slightly, but not by as much as might be expected; the reduction might be about a factor of 2 for a 10-fold dilution. It is also not clear how the effect of dilution would change in the presence of a thickening agent.

I want to enlarge my particles in solution. How should I do this?

The key to enhancement in solution is to stop the enhancement reaction as quickly as possible once the particles have reached the desired size, either by removing the enhancement reagent, or by terminating the reaction. Dilution can be used to limit the reagent, but it is very difficult to quantitate the amount of silver required since not all the gold particles will be enlarged. Slowing down the reaction will allow easier monitoring and control, and give more time to effect a final separation.

The reaction may be further slowed or stopped once the particles have reached the required size by lowering the pH: be careful to use a buffer that does not precipitate silver. Low pH citrate buffer is a good choice. Then separate the enhanced particles as rapidly as possible. Membrane centrifugation or spin columns may be helpful for this; gel filtration over a gel such as Superose-6 may also be helpful. However, the media should be tested first to make sure that the filtered enlarged particles may be resuspended afterwards.

Can I use silver enhancement on synthetic surfaces?

This should be no problem - provided the surface in question does not precipitate silver, or coordinate silver ions which can then produce background. Test the surface (in the absence of gold particles, but after any other treatments or steps in the procedure you plan to use) with a small amount of the enhancement reagent for an appropriate enhancement time, then wash thoroughly. Leave to dry if this is how the final sample will be treated: then examine carefully for background. If you find background, then a wash with 0.1% Tween-20 or a similar detergent in sodium citrate buffer before application of the silver enhancer may help to reduce or eliminate background; for other methods to control background, see our previous article on this subject.

Can I use silver enhancement to demonstrate other targets besides gold nanoparticles?

Actually, silver enhancement was not originally used for gold particles, but for the histological demonstration of heavy metals and compounds in tissues; these included gold and silver, but also elements such as zinc, selenium and bismuth. Gorm Danscher, who first described the process, has used it to demonstrate a variety of elements, and has recently applied it to the tracing of quantum dots, which include species prepared from some of these elements. Therefore, silver enhancement can be used for the demonstration of a variety of species.

Transition metal ions are thought to be one cause of background deposition in the silver enhancement reaction when silver enhancing gold particles for blots and light microscopy, and therefore we are careful to exclude sources of transition metal ions from procedures where silver enhancement is required, and to wash with treatments which can remove any extraneous transition metals. Washing with a chelating agent - such as an EDTA (ethylene diamine tetraacetic acid) salt can chelate and remove transition metal ions before silver enhancement, and we have fund reduced background in blots after such treatments. Therefore, it is also reasonable to use silver enhancement to demonstrate deposits of transition metals.

For activity towards silver enhancement, what we look for is an active redox chemistry where it is likely that one of the redox couples of the target can couple with that of the silver/hydroquinone reduction. Iron and Manganese are potential candidate for this because they have extensive redox chemistry; compounds are known for every oxidation state from 0 to +7, so there's a good chance that this chemistry includes redox couples that fit with the silver enhancement reaction, and that they have some reactivity towards silver enhancement.

Reference:

• Danscher, G., and Stoltenberg, M.: Silver enhancement of quantum dots resulting from (1) metabolism of toxic metals in animals and humans, (2) in vivo, in vitro and immersion created zinc-sulphur/zinc-selenium nanocrystals, (3) metal ions liberated from metal implants and particles. Prog. Histochem. Cytochem., 41, 57-139 (2006).
  • Abstract (courtesy of Science Direct).

More information:

• Catalog and general information - Silver enhancement
• Product information and instructions - HQ Silver
• Product information and instructions - LI Silver
• Technical help - Silver enhancement
• References sorted by Application - Silver enhancement

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Towards Correlative Double Labeling with FluoroNanogold

Currently, little is known about the arborization of the cardiac sympathetic neural network, and acquisition of such information from living heart tissue is difficult because of the movement of beating. Analysis in healthy and diseased heart of the cardiac sympathetic nervous system would be valuable for understanding heart disease processes, and could lead to better treatments for catastrophic cardiac circumstances. Determination of the structural relationship of sympathetic fibers to the surrounding cardiomyocytes may provide the basis for methods to assess areas deprived of sympathetic input, hyperinnervated regions, and understanding of alterations in injured myocardium. Implementation requires mice with fluorescent reporter genes in conjunction with high resolution TPLSM to explore the innervation of the intact, live heart. The experimental objective was comparison of the localization of dopamine hydroxylase (hDBAH) using EGFP expression, placing EGFP under control of DBH promoter, with immunolocalization using antibodies against DBH and tyrosine hydroxylase.

FluoroNanogold was used in varied levels of microscopy, including two-Photon Laser Scanning Microscopy (TPLSM), Confocal Laser Scanning Microscopy (CLSM), and Transmission Electron Microscopy (TEM). TPLSM images were taken of an intact, live transgenic heart; this was then fixed and vibratome sections were cut. Intact sympathetic ganglion (positive control) and heart sections were exposed to a primary antibody against eGFP for 48 hours after proper permeabilization and blocking. The primary antibody was localized with secondary FluoroNanogoldTM overnight, washed and imaged with a Zeiss LSM-510 Meta CLSM. Sections were then processed for TEM via silver enhancement, osmication, and embedding in Embed 812. Thin sections were stained with uranyl acetate, and viewed in the transmission electron microscope.

The presentation also included double labeling to demonstrate colocalization of EGFP expression with tyrosine hydroxylase. Tyrsosine hydroxylase was labeled with FluoroNanogold as above, and EGFP was stained using biotinylated anti-GFP followed by Nanogold-streptavidin: the two labels were enhanced using the differential silver enhancement method of Hong Yi: first labeling with anti-TH and Alexa Fluor 594 FluoroNanogold followed by a first round of silver enhancement, then EGFP labeled with biotinylated anti-GFP, followed by Nanogold-streptavidin and a second round of silver enhancement. This procedure produced two populations of enlarged particles which could be distinguished by particle size:

Determination of innervations in the substructure in the myocardium is accomplished, in part, by using TEM in conjunction with CLSM fluorescent techniques through the use of FluoroNanogold. However, the experiments also illustrated some of the challenges that come with FluoroNanogold: that the optimum use concentration for the two labels may be slightly different. Heart tissue is optically very dense, and concentrations that provided an adequate fluorescence signal resulted in significant background in the TEM, while concentrations producing specific TEM signal gave low fluorescence. In these situations, control of non-specific binding may need to be quite rigorous, since both labels can interact non-specifically with biological components. We have found that the following methods may help ensure the brightest, cleanest signal:

• The most effective blocking agent we have tested is 5% nonfat dried milk. This was particularly effective when it was used in the FluoroNanogold conjugate incubation buffer in addition to the blocking step. Cold-water fish gelatin has also been found to be helpful for gold probes generally.

• Adjusting camera exposure: manual control of exposure can help obtain the best signal-to-noise ratio. FluoroNanogold is frequently compared with commercially available fluorescently labeled IgG conjugates. Since these are larger and more highly labeled, they give brighter fluorescence. If automatic exposure adjustment is allowed with FluoroNanogold-stained specimens, the greater exposure can lead to higher apparent backgrounds. Setting the camera exposure manually can be used to overcome this effect.

• For reducing the background in electron microscopy, sodium citrate buffer was found to be more effective than other buffers when used as a wash before silver enhancement. 0.02 M sodium citrate at pH 7.0 works well with HQ Silver, while pH 3.5 works best with the Danscher silver formulation.

• Background binding is often attributed to hydrophobic interactions (both the gold and fluorescent labels have some hydrophobicity), and therefore adding reagents that reduce hydrophobic interactions to the wash buffer may help remove non-specific binding. Examples include:
  • 0.6 M triethylammonium bicarbonate buffer (prepared by bubbling carbon dioxide into an aqueous suspension of degassed triethylamine with stirring; see Safer et al., reference below.
  • 0.1% to 1% detergent, such as Tween-20, or Triton X-100. 0.1% saponin may also be useful since its effects are reversible, so ultrastructural preservation may be improved if it is removed in later steps.
  • 0.1% to 0.5% of an amphiphile, such as benzamidine or 1,2,3-trihydroxyheptane.

Reference:

• Jackson, K.; Miller, C.; Gattone, V., and Rubart, M.: Imaging Cardiac Sympathetic Neurons: a Correlative Study Using Fluorescence and TEM. Microsc. Microanal., 14 (Suppl. 2: Proceedings),; Marko, M.; McKernan, S.; Shields, J.; Scott, J.-H.; Kotula, P.; Anderson, I., and Woodward, J. (Eds.), Cambridge University Press, 1514CD (2008).

Reference for preparation of triethylammonium bicarbonate buffer:

• Safer, D.; Bolinger, L., and Leigh, J. S.: Undecagold clusters for site-specific labeling of biological macromolecules: simplified preparation and model applications. J. Inorg. Biochem., 26, 77-91 (1986).
  • Abstract (courtesy of Science Direct).

Doug Keene and group recently described a simplified method for correlative microscopy that may be appropriate for FluoroNanogold experiments. Correlative confocal and electron microscopy was used to study rat chondrosarcoma cells expressing GFP/YFP fusion proteins labeled using immunogold labeling. Cells were grown on plastic coverslips, lightly fixed in paraformaldehyde / glutaraldehyde, quickly dehydrated to 90% ethanol (all ice cold), then infiltrated twice at for 45 minutes in cold LR white, and embedded at 60°C overnight. Confocal images from 0.5 µm thick sections are overlaid atop TEM images of the same cells, which were collected from the next serial ultrathin 80 nm section after a separate immunogold labeling procedure, using a rabbit anti-GFP primary and 6 nm colloidal gold-anti-rabbit secondary. This procedure demonstrates important advantages for the proposed work. Fluorescence is observed in LR White, confirming that confocal and electron microscopic images can be acquired from the same specimen. However, this procedure would be further simplified with FluoroNanogold: the need for a separate immunogold labeling step would now be eliminated, thus yielding a quicker procedure with even less potential for differences between the two sections.

Reference:

• Keene, D. R.; Tufa, S. F.; Lunstrum, G. P.; Holden, P, and Horton, W. A.: Confocal/TEM overlay microscopy: a simple method for correlating confocal and electron microscopy of cells expressing GFP/YFP fusion proteins. Microsc. Microanal., 14, 342-348 (2008).
  • Abstract (courtesy of Cambridge Journals).

More information:

• Catalog information: FluoroNanogold
• Alexa Fluor® 488 FluoroNanogold-Fab': Product information and instructions
• Alexa Fluor® 594 FluoroNanogold-Fab': Product information and instructions
• Alexa Fluor® 594 FluoroNanogold-Streptavidin: Product information and instructions
• FluoroNanogold: Technical help
• FluoroNanogold: References, sorted by application

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Coffee, Calcium and the Ryanodine Receptor - Localized by Nanogold®

• High penetration into cells and tissues.
• High labeling density.
• Close to quantitative labeling of antigenic sites.
• High labeling resolution.

Together with HQ Silver, this provides the ideal combination of features to produce high-quality staining:

• The only commercial silver enhancer to feature a protective colloid for highest size uniformity and morphologically consistent enhancement.
• Near neutral pH and low ionic strength ensure maximum specimen integrity.
• High proportion of gold particles are enlarged.
• Highly selective reaction with low background.

Resolution advantage: size comparison of Nanogold-Fab' with conventional 5 nm colloidal gold-IgG probe, showing overall probe size and distance of gold from target. Due to its position at the hinge region, Nanogold is positioned closer to the target upon binding, yet does not hinder or interfere with binding.

Shimizu and group demonstrated these advantages recently in their study of use-dependent amplification of presynaptic Ca²⁺ signaling by axonal ryanodine receptors at the hippocampal mossy fiber synapse, reported in a recent issue of the Proceedings of the National Academy of Sciences of the USA. The authors were investigating the hypothesis that presynaptic Ca²⁺ stores regulate Ca²⁺ dynamics within the nerve terminals at certain types of synapse. Little is known about their mode of activation, molecular identity, and detailed subcellular localization, and hence the authors used a variety of methods to study the distribution and actions of the species involved in presynaptic Ca²⁺ signaling. The authors showed that ryanodine-sensitive stores exist in axons and amplify presynaptic Ca²⁺ accumulation at the hippocampal mossy fiber synapses, which display robust presynaptic forms of plasticity. Caffeine, a potent drug inducing Ca²⁺ release from ryanodinesensitive stores, causes elevation of presynaptic Ca²⁺ levels and enhancement of transmitter release from the mossy fiber terminals. Blockers of ryanodine receptors, TMB-8 or ryanodine, reduce presynaptic Ca²⁺ transients elicited by repetitive stimuli of mossy fibers; however, they do not affect those evoked by single shocks, and this suggests that ryanodine receptors amplify presynaptic Ca²⁺ dynamics in an activity dependent manner.

In order to localize ryanodine receptors, the group generated a specific antibody against the type 2 ryanodine receptor (RyR2; originally referred to as the cardiac type) and examined its cellular and subcellular localization using immunohistochemistry. Under deep pentobarbital anesthesia, mice were perfused transcardially with either 4% paraformaldehyde in the 0.1 M sodium phosphate buffer (PB, pH 7.2) or 4% paraformaldehyde/0.1% glutaraldehyde in the PB. Microslicer (50 µm) sections of the brain and cryostat sections (50 µm) of the heart and skeletal muscles were used. All immunohistochemical incubations were done at room temperature. For immunofluorescence, microslicer sections were incubated with 10% normal donkey serum for 20 minutes, a mixture of primary antibodies overnight (1 µg/ml), and a mixture of Alexa 488-, Cy3-, and Cy5-labeled species-specific secondary antibodies for 2 hours at a dilution of 1: 200. Phosphate-buffered saline (PBS, pH 7.4) containing 0.1% Tween-20 was used as diluent of antibodies and a washing buffer. Images were taken with a light microscope equipped with a digital camera or with a confocal laser scanning microscope. For immunoelectron microscopy, microslicer sections immunoreacted for the RyR2 were subjected to silver-enhanced immunogold, using 1.4-nm Nanogold-labeled anti-rabbit IgG, followed by enhancement with HQ silver. Sections were further treated with 1% osmium tetroxide and uranyl acetate, then and embedded in Epon 812. Ultrathin sections (70 nm in thickness) were prepared with an ultramicrotome, and photographs were taken with an H7100 electron microscope.

RyR2 is highly expressed in the stratum lucidum of the CA3 region and mostly colocalizes with axonal marker NF160 but not with terminal marker VGLUT1. Immunoelectron microscopy revealed that RyR2 is distributed around smooth ER within the mossy fibers but is almost excluded from their terminal portions. The above results strongly suggested that axonal RyR2 mediated activity dependent presynaptic CICR at the mossy fiber synapse. However, they do not exclude the possibility that other types of ryanodine receptors were involved. To address this possibility, the cellular distribution of RyR1 was examined using the specific antibody for this receptor. In contrast with the intense labeling of RyR2 within mossy fiber axons, RyR1 immunoreactivity in the hippocampus was generally weaker than in the cerebellum and predominantly expressed in postsynaptic neurons, indicating that RyR1 was less likely to contribute to presynaptic CICR at the hippocampal mossy fiber synapse. These results suggest that axonal localization of RyR2 at sites distant from the active zones enables use dependent Ca²⁺ release from intracellular stores within the mossy fibers and thereby facilitates robust presynaptic forms of plasticity at the mossy fiber-CA3 synapse.

Reference:

• Shimizu, H.; Fukaya, M.; Yamasaki, M.; Watanabe, M.; Manabe, T., and Kamiya, H.: Use-dependent amplification of presynaptic Ca²⁺ signaling by axonal ryanodine receptors at the hippocampal mossy fiber synapse. Proc. Natl. Acad. Sci. USA, 105, 11998-12003 (2008).
  • Abstract (courtesy of the Proceedings of the National Academy of Sciences of the USA).

More information:

• Application note: Pre-embedding labeling with Nanogold and HQ Silver enhancement
• Nanogold conjugates: catalog information
• Nanogold-antibody probes: instructions and product information
• Nanogold conjugates: technical help
• Pre-embedding Nanogold labeling: references
• Silver enhancers: catalog information
• HQ Silver: instructions and product information

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Reach the Right Person: Contact Information

If you have a question about an order, such as whether it has been received or shipped, you should contact our main office (nano¤nanoprobes.com); telephone 1-877-447-6266 in North America, ++ (631) 205-9490 from elsewhere; fax (631) 203-9493. You should not contact technical support: details of individual orders, shipping, and billing are not accessible to technical support personnel.

If you are looking for Nanogold® conjugate or reagent that is similar to our catalog items and the chemistry has already been established - such as a multiply functionalized Nanogold particle, or a Nanogold-labeled primary antibody - we can usually prepare such products as custom syntheses. Please contact technical support (tech¤nanoprobes.com) if you have a specific custom synthesis request, or fill out our custom synthesis request form. If your request includes steps that we do not do regularly, such as labeling a different type of protein or biomolecule, we can often consider it, but may need to treat it as a short-term contract research project, in which payment is required whether or not the synthesis is successful.

For your information, contact information is summarized below:

If you are looking for a specific person, our web site includes a listing of key personnel with individual extensions and e-mail addresses.

More information:

• How to Order
• Custom synthesis: general information
• Custom synthesis terms and conditions
• International Distributor list
• How to order from our international distributors
• News (holiday hours, closings)

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Other Recent Publications

Reference:

• Aili, D.; Enander, K.; Baltzer, L., and Liedberg, B.: Assembly of polypeptide-functionalized gold nanoparticles through a heteroassociation- and folding-dependent bridging. Nano Lett., 8, 2473-2478 (2008).
  • Abstract (courtesy of ACS Publications).

Electron microscopic analysis was used to characterize these structures. Huh7, HepG2, COS-7, and HeLa cells cultured on coverslips were fixed with 2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer, and post-fixed in a mixture of 1% osmium tetroxide and 0.1% potassium ferrocyanide in the same buffer. Isolated LDs adhering to coverslips were fixed with a mixture of 2.5% glutaraldehyde and 1% osmium tetroxide. For glucose-6-phosphatase histochemistry, cells fixed with 0.5% glutaraldehyde in 0.1 M sodium cacodylate buffer were processed using the lead-nitrate method. For pre-embedding immunoEM, cells were fixed and permeabilized, then labeled sequentially by a primary antibody and a FluoroNanogold secondary antibody and treated with GoldEnhance EM to visualize Nanogold particles. Labeling by filipin was performed as previously described. After dehydration through an ethanol series, samples were embedded in Quetol-812 resin; ultrathin sections were observed in the electron microscope at 100 kV. The ApoB-crescent was identified as a thin, cholesterol-rich ER cistern fused to a lipid droplet, and topologically this structure was equivalent to a lipid-ester globule between the two leaflets of the ER membrane. ApoB localized in the thin cisternal lumen: its binding to LDs was resistant to alkaline treatment. Overexpression of ADRP or TIP47 suppressed the increase in the number of ApoB-crescents, whereas knockdown of these proteins had the opposite effect. From these results, it was inferred that the ApoB-crescent was formed by an LD that is arrested in the ER membrane by tight binding of lipidated ApoB to its luminal surface, suggesting that ApoB processing and LD formation are closely linked.

Reference:

• Ohsaki, Y, Cheng, J, Suzuki, M, Fujita, A, and Fujimoto, T.: Lipid droplets are arrested in the ER membrane by tight binding of lipidated apolipoprotein B-100. J. Cell Sci., 121, 2415-2422 (2008).
  • Abstract (courtesy of Journal of Cell Science).

The use of Nanogold® as a mass label for mass spectrometric analysis of biological molecules has been described previously. The use of matrix-assisted laser desorption/ionization (MALDI) imaging mass spectrometry (IMS) as a method for investigating the distribution of targets in biological systems is reviewed in a recent paper in Histochemistry and Cell Biology. MALDI-IMS is a powerful tool for investigating the distribution of proteins and small molecules through the in situ analysis of tissue sections. MALDI-IMS can determine the distribution of hundreds of unknown compounds in a single measurement, and enables the acquisition of cellular expression profiles while preserving cellular and molecular integrity. Recent advances in the practice of imaging mass spectrometry which make the technique more sensitive, robust, and ultimately more useful are reviewed and described, and recent applications in basic research and in clinical settings are discussed.

Reference:

• Walch, A.; Rauser, S.; Deininger, S. O., and Höfler, H.: MALDI imaging mass spectrometry for direct tissue analysis: a new frontier for molecular histology. Histochem. Cell Biol., 130, 421-434 (2008).
  • Abstract (courtesy of SpringerLink).
  • Reprint (PDF) (courtesy of SpringerLink).

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