N A N O P R O B E S     E - N E W S

Vol. 9, No. 11          November 30, 2008

They are also potentially important building blocks for nanostructured materials and nanodevices. As shown below, decoration of DNA with positively charged Nanogold is a potential method for preparing conductive nanowires, and these molecules may be used for the attachment of multiple conjugate biomolecules to make polyfunctional probes. They can also provide Enhanced efficiency in DNA transfection.

In their recent paper in the Journal of Bacteriology, Matias and Beveridge describe the use of Positively Charged Undecagold, in addition to Mono-Sulfo-NHS-Nanogold-antibody labeling, to identify components of the periplasm in Bacillus subtilis bacteria by cryoelectron microscopy. Like other gram-positive bacteria, this organism possesses a thick cell wall that plays a major role in survival. Gram-positive cell walls are highly dynamic structures that are constantly remodeled during the cell cycle, and precise assembly of the polymers that make up the cell wall into a complex functional 3-dimensional network is critical for their biology; however, conventional high-resolution imaging methods such as electron microscopy of thin sections reveal few structural details, typically showing only a thick and undifferentiated cell wall tightly apposed to the plasma membrane - possibly due to the effect of harsh chemicals used during specimen preparation. In contrast, cryoelectron microscopy affords observation of this complex structure in a close-to-native state. A distinctive feature revealed by cryo-EM in comparison with conventional EM methods is a gram-positive periplasmic space between the plasma membrane and an often differentiated cell wall. Since almost nothing was known about the composition of this feature, the authors used cryo-EM of frozenhydrated sections in combination with gold nanoparticles and key hydrolytic enzymes to probe for major periplasmic components in B. subtilis.

B. subtilis 168 was grown in tryptic soy broth at 37°C up to mid-exponential phase (optical density at 600 nm, 0.5 to 0.8). Cell wall fragments were isolated from cells broken by a French press by boiling cell wall fragments in 4% sodium dodecyl sulfate (SDS) for 2 hours to remove cytoplasmic and membrane contaminants, then washing in deionized water to remove SDS. Protoplasts were generated by first washing cells three times in a phosphate-buffered saline (PBS) buffer (20 mM phosphate, 50 mM NaCl, 10 mM MgCl₂, pH 7.4) supplemented with 10% (wt/vol) sucrose and then incubating cells in PBS-sucrose buffer with 1 mg/mL of lysozyme at 37°C until cells rounded up (usually for 45 to 60 minutes, monitoring with a light phase-contrast microscope.

The cell envelope of B. subtilis was initially probed for the distribution of negatively charged sites, a characteristic feature of its wall polymers (wall teichoic acids and peptidoglycan), which in B. subtilis account for almost all of the wall mass, using positively charged undecagold (PCU). For labeling, prewashed pellets of cells (75 µL), cell wall fragments (70 µL), and protoplasts (80 µL) were incubated with 150, 135, and 160 nM PCU, respectively. Cells were incubated for 5 minutes and then were cryoprotected and immediately frozen, whereas cell wall fragments were further washed three times in buffer before being cryoprotected and frozen. For cryo-EM, cells, cell walls, and protoplasts were cryoprotected in 17% (wt/vol) dextran, 20% dextran, and 10% dextran plus 10% sucrose, respectively; dextran was generally used for cryoprotection to ensure that frozen-hydrated sections had no to few crevasses. In all cases, the amount of cryoprotectant added ensured good freezing (vitrification) of samples. Specimens were frozen at a high pressure and cryosectioned, and cryo-EM was performed under relatively low electron dose conditions (estimated electron dose, 1,000 to 2,500 e^- nm^-2). Images were analyzed using NIH ImageJ software. The PCU particles were small enough to diffuse through the cell wall network and label negatively charged surface components from the outer face of the membrane through the periplasmic space to the outer surface of the wall. The particles are too small to be individually visualized, but bind with sufficient density to produce improved contrast of the structures to which they bound. Undecagold particles accumulated almost uniformly across the envelope, making it difficult to distinguish the interface between the periplasmic space and the cell wall, but did not significantly alter the envelope thickness from that of unlabeled cells. In isolated cell wall fragments, however, denser labeling was observed at the inner and outer surfaces, consistent with new wall material being added at its inner surface and the old wall being shed by autolytic enzymes at its outer surface.

On cells whose cell walls were enzymatically hydrolyzed (protoplasts), a surface diffuse layer extending ~30 nm from the membrane was revealed. In order to explore the fine structure of this layer in more detail, lipoteichoic acid (LTA) was localized using a Nanogold-labeled antibody. LTA is a characteristic envelope component of a large number of gram-positive bacteria, with an essential role in the physiology of these organisms. LTAs are macroamphiphiles, composed of long, negatively charged hydrophilic chains (poly(glycerophosphate)), inserted into the outer leaflet of the membrane through a glycolipid anchor. For labeling of lipoteichoic acid (LTA) on the surfaces of protoplasts, mouse monoclonal anti-LTA IgG was conjugated to Mono-Sulfo-N-hydroxysuccinimido-Nanogold. Sufficient Mono-Sulfo-NHS-Nanogold was added to 1.25 mg of anti-LTA IgG to give a ratio of 3.5 nanogold particles bound to each IgG molecule. The anti-LTA-Nanogold conjugate was separated from unbound Nanogold using a microconcentrator with a cutoff of 50 kDa. To label LTA on protoplast surfaces, protoplasts were washed twice in the sucrose-PBS buffer (to remove hydrolyzed wall components) and once in a blocking buffer (sucrose-PBS buffer with 0.4% skim milk), incubated with the anti-LTANanogold for 40 minutes in the blocking buffer, then washed four times in the blocking buffer (to remove unbound IgG-Nanogold complexes). As a labeling control, mouse serum IgG (Sigma) was similarly labeled with Nanogold and incubated with protoplasts. Immunolabeling of protoplasts with anti-LTANanogold caused clumping of protoplasts; this was expected because of the surface localization of LTA and the bivalent nature of IgG binding. At a medium level of magnification, some accumulation of Nanogold along the membrane surface could be observed. At a high magnification, a concentration of Nanogold complexes either between two adjacent protoplast membranes or on the membrane surface of a noncontacting protoplast clearly marked the presence of LTA in the surface diffuse zone of protoplasts, and suggested a high abundance of LTA in this surface zone. In closely associated protoplasts, a higher-density band was observed between the adjacent membranes, indicating that the binding of LTA epitopes took place on both membranes.

Protoplasts were also treated with a nonspecific protease (proteinase K) to determine the impact of removing membrane-associated proteins (such as lipoproteins) on the visualization of the diffuse surface layer: lipoproteins and membrane-anchored proteins were thought to represent the only other characteristic components of the B. subtilis envelope that could have contributed to the mass of the surface diffuse layer observed here by cryo-EM. Proteinase K-treated protoplasts showed a diffuse layer only slightly thinner than that of untreated protoplasts, with a similar level of contrast, implying that (lipo)proteins are not a major component of the surface diffuse zone. The combined results show that the LTA layer spans most of the thickness of the periplasmic space, which strongly suggests that LTA is a major component of the B. subtilis periplasm.

Reference:

• Matias, V. R., and Beveridge, T. J.: Lipoteichoic acid is a major component of the Bacillus subtilis periplasm. J. Bacteriol., 190, 7414-7418 (2008).
  • Abstract (courtesy of the Journal of Bacteriology).

More information:

• Nanogold labeling reagents: Catalog information
• Mono-Sulfo-NHS-Nanogold: Product information, instructions and protocols
• Nanogold labeling reagents: Technical help
• Undecagold: Catalog information
• Positively Charged Undecagold: Product information, instructions and protocols
• Undecagold: Technical help
• References: Nanogold labeling for cryoEM

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Nanogold® Labeling for Other Functional Groups

Hydroxyls

• Use a hydroxyl-reactive cross-linker.

  A hydroxyl-reactive cross-linker is commercially available: p-Maleimidophenylisocyanate. Unfortunately, the maleimide group, which this reagent introduces, is not directly reactive towards Nanogold labeling reagents: but simple treatment with mercaptoethylamine hydrochloride will add an amino- group, which you can then label with Mono-Sulfo-NHS-Nanogold.

• Make it more reactive towards amines, so you can label with Monoamino Nanogold®.

  This can be done relatively easily: react it with tosyl chloride (p-toluene-sulfonato chloride). This converts it to a tosyl group, which is now strongly reactive towards amines: simply react with Monoamino Nanogold.

  Tosylation of hydroxylated molecule and reaction with Monoamino Nanogold.

• Oxidize it to a ketone or aldehyde, then react with a bifunctional hydrazido cross-linker with a Nanogold-reactive functionality at the other end.

  Many methods are available for the oxidation of hydroxyls to carbonyls. Once this is achieved, you can use a heterobifunctional hydrazide cross-linker to introduce a reactive functional group. Succinimidyl 6-(3-[2-pyridyldithio]-propionamido) hydrazide (SPDP hydrazide) introduces a disulfide, which is readily reduced to a sulfhydryl and labeled with Monomaleimido Nanogold:

  Oxidation of hydroxyl to carbonyl, followed by reaction with hydrazido cross-linker, activation of disulfide, and labeling with Monomaleimido Nanogold.

  Many carbonyl-reactive hydrazido- maleimide cross-linkers are available: you can use these to introduce maleimides, which may then be reacted with mercaptoethylamine hydrochloride to convert them to amines and labeled with Mono-Sulfo-NHS-Nanogold. For examples, the list of heterobifunctional cross-linkers from Molecular Biosciences, cross-linker selection guide from Pierce are useful references.

  If you have a primary hydroxyl and can convert it to an aldehyde, you have a simpler option: you can react it directly with Monoamino Nanogold. This is the same reaction used, after periodate oxidation, to label RNA and glycoproteins, and details are in our application note on RNA labeling.

If you have a cis-1,2-dihydroxy group, you can oxidize with sodium periodate to generate the dialdehyde, then react with Monoamino Nanogold. The full procedure is given in our application note on RNA labeling: you can also use this procedure for any molecule containing a cis-1,2-dihydroxy group, such as a glycoprotein or other carbohydrate:

Periodate oxidation of carbohydrate (cis-1,2-diol) followed by reaction with Monoamino Nanogold.

Aldehydes and Ketones

The procedure for this is described on our web site, in the application note on labeling RNA and glycoproteins. In this procedure, the compound to be labeled is a glycoconjugate that has been oxidized using periodate to yield a dialdehyde: this is then reacted directly with Monoamino Nanogold. An important consideration is that the conjugation reaction yields a Schiff base, which must then be reduced to a secondary amine. An example is shown below.

Example showing labeling of an aldehyde with Monoamino Nanogold

Example procedure:

1. Dissolve 30 nmol of Monoamino Nanogold in 100 microliters anhydrous dimethyl sulfoxide (DMSO). Mix Nanogold-DMSO solution (16.7 microliters, 10-fold excess) in 100 mM PIPES (30 microliters) with oxidized biomolecule in buffer solution at pH 7.2 or higher (100 microliters). Incubate at 4°C for 60 minutes.

2. Add 1.5 microliters of fresh 20 mg/mL borane tert-butylamine complex (stable for about 15 minutes). incubate at 4°C for 30 minutes. Stop reaction with 2 microliters of acetone. After 3 minutes at 4°C, separate labeled biomolecule using an appropriate gel filtration column.

Alternatively, it may be possible to convert the aldehyde, or even the ketone, to a carboxylic acid. If so, you can then convert the carboxylic acid to a reactive ester using either 1-Ethyl-3-[3-dimethylaminopropyl] carbodiimide Hydrochloride (EDC) with Sulfo-N-hydroxysuccinimide or N,N-carbonyldiimidazole in DMF, then react the activated ester with Monoamino Nanogold.

You can also cross-link directly using one of the many carbonyl-reactive hydrazido- maleimide cross-linkers that are available. Succinimidyl 6-(3-[2-pyridyldithio]-propionamido) hydrazide (SPDP hydrazide) introduces a disulfide, which is readily reduced to a sulfhydryl and labeled with Monomaleimido Nanogold. Others introduce maleimides, which may then be reacted with mercaptoethylamine hydrochloride to convert them to amines, and labeled with Mono-Sulfo-NHS-Nanogold. You can find suitable cross-linkers from the list of heterobifunctional cross-linkers from Molecular Biosciences, or the cross-linker selection guide from Pierce.

Carboxylic Acids

If I want to label a molecule with Nanogold and the only functional group I have is a carboxylic acid, what should I do? You can do this using Monoamino Nanogold®. To carry out the labeling reaction, you need to convert the carboxylic acid group to a reactive ester, which will then react with the amine - the same reaction that occurs when Mono-Sulfo-NHS-Nanogold is used to label an amine. Some suitable reaction schemes are shown below:

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

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

Reference:

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

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

Reference:

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

Phosphates in Lipids

Options for conjugating Nanogold to this molecule are very limited, because the PI(3)P molecule does not have any reactive functional groups to which Nanogold may be readily conjugated. Thiols or primary aliphatic amines would be best; however, as we have explained in previous articles, cross-linking chemistry is also possible for carboxylic acids or hydroxyls, although these procedures are slightly more difficult. However, PI(3)P has none of these, and contains only three groups which can be readily modified in any way. Is it possible to label any of them with Nanogold?

Considering each group in turn:

• The phosphate group: This is the best choice for labeling, because cross-linking chemistry has already been described in the literature. The simplest and best-documented approach is to activate the phosphate group in the same manner as a carboxylic acid: either by EDC/Sulfo-NHS coupling, activating with EDC (1-ethyl-3,3'-dimethylaminopropyl carbodiimide) followed by sulfo-NHS, or by treatment with CDI (N,N'-carbonyldiimidazole) in DMF (dimethylformamide). In either case, you could then react the activated ester with Monoamino-Nanogold.

  Reference:

  Chu, B. C. F.; Kramer, F. R., and Orgel, L. E.: Synthesis of an amplifiable reporter RNA for bioassays. Nucleic Acids Res., 14, 5591-5603 (1986).

  • Abstract (courtesy of Nucleic Acids Research).
  • Reprint (PDF) (courtesy of PubMed Central).

  Note that there is another option besides direct reaction of the activated ester with Monoamino Nanogold. The authors react their activated ester with cystamine, which is cleavable to give a thiol group: an alternative approach for Nanogold labeling is use cystamine exactly as described, cleave, and label using Monomaleimido Nanogold. Both reactions are described in more detail in the technical help section of our web site.

• The double bonds: While modification of the double bonds is possible chemically, it is likely to be undesirable because modification of these groups may change the biological or liposomal behavior of the molecule.

• The carbonyl (C=O) group: However, because this is actually part of an ester linkage, reactivity will be difficult and most methods will involve cleavage of the ester group, so part of the PI(3)P will be lost. Two potentially useful reactions are the Wittig or Horner-Wadsworth-Emmons reactions; however, these generally do not work well with esters, and you will probably need to find a variation that is adapted to this functionality. If your experiment would work with a cleaved derivative, this may be an option.

The reactions used for phosphate group labeling are shown below:

One consideration in selecting functional group for labeling is whether the site participates in, or might hinder, the activity that you will be using the labeled probe to study; if so, then this may not be the best strategy. It is often preferable to conjugate Nanogold to the hydrophilic region of the molecule rather than the hydrophobic domain, both because Nanogold itself is hydrophilic, and also because the assembly of lipids into liposomes and other structures is mediated by the interactions of the hydrophobic domain: this activity would be impacted by the attachment of a large label at this site.

More information:

• Guide to Gold Cluster Labeling
• Catalog information: Nanogold labeling reagents
• Product instructions: Monomaleimido Nanogold
• Product instructions: Mono-Sulfo-NHS-Nanogold
• Product instructions: Monoamino Nanogold
• Technical help: Nanogold labeling reagents
• References sorted by application - Nanogold labeling
• Nanogold labeling: plan your reaction for best results

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Tracking Down Food Poisoning with NanoVan

McIntyre, Beniac and group have successfully used NanoVan in their studies identifying bacteria responsible for food poisoning outbreaks in British Columbia. The Bacillus cereus group, which is responsible for many food poisoning cases, contains five species: B. cereus sensu stricto, B. thuringiensis, B. anthracis, B. mycoides, and B. weihenstephanensis. These are difficult to distinguish using standard biochemical schemes, chemotaxonomic methods, or phylogenetically relevant target genes; furthermore, many distinguishing pathogenicity markers in this group can be attributed to mobile plasmids. B. cereus sensu stricto carries the plasmid-borne emetic toxin cereulide (ces), while B. thuringiensis carries insecticidal crystal protein (ICP) (cry) genes on one or more plasmids. Because the identification of B. cereus group members using molecular techniques is not a standard part of food-poisoning diagnoses, species such as B. thuringiensis may be underreported. For example, initial investigation of a 2005 food-poisoning event implicating imported strawberries identified the isolate phenotypically as B. cereus, but a later PCR analysis showed it to be B. thuringiensis. In order to better assess the actual proportions of different B. cereus group species causing food poisoning outbreaks, the authors characterized the isolates collected from food or clinical specimens in food-borne outbreaks between 1991 and 2005 using a combination of molecular and phenotypic typing methods.

Analyses were conducted using multiplex PCR and electron microscopy. Frozen isolates were retrieved onto blood agar plates and incubated at 35°C for 24 hours. Pathogenicity genes for emetic cereulide toxin (nonribosomal peptide synthetase, NRPS) and ICP (cry1 or cry2) were detected in multiplex PCR assays. Strains positive for NRPS were designated as B. cereus, those positive for ICP (by microscopy or PCR) as B. thuringiensis, those with rhizoidal growth on nutrient agar as B. mycoides, and all other strains with the typical B. cereus phenotype as B. cereus NRPS^- ICP^-.

Samples of B. cereus group bacteria were prepared for electron microscopy by fixation with 2% glutaraldehyde and 1% paraformaldehyde, then adsorbed to a glow-discharged carbon-coated Formvar film on a 400-mesh copper grid for 1 minute, then negatively contrasted with NanoVan (2% methylamine vanadate). The negatively stained specimens were then imaged in an FEI Tecnai 20 TEM operated at 200 kV, using nominal instrument magnifications of 5,000 to 9,600X. Images were acquired with a charge-coupled-device camera. The majority of the 155 food-poisoning isolates were identified as B. cereus NRPS^- ICP^- (58%, n = 90): no plasmid markers for emetic toxin or ICPs were detected or visualized. Thirty-eight (24.5%) isolates were identified as B. cereus NRPS⁺, and 23 (15%) isolates identified as B. thuringiensis: these were positive for either cry1, cry2, or both (in one case, ICPs were detected only by TEM). A small number (2.6%) were identified as B. mycoides based on rhizoid growth demonstrated on nutrient agar. Overall, B. cereus isolates were identified in 23 of the 39 outbreaks (B. cereus NRPS⁺ in 5 [12.8%] and B. cereus NRPS^- ICP^- in 18 [46.1%]), B. thuringiensis in 4 (10.3%), B. mycoides in 1 (2.6%), and mixed species of Bacillus in 11 (28.2%). No B. cereus group-like bacterium reviewed here was consistent with B. weihenstephanensis or B. anthracis.

The rapid identification of Bacillus species implicated in food poisonings can be facilitated by PCR, and the combination of PCR with negative stain electron microscopy provides an unambiguous method for differentiating between similar species. PCR in tandem with phenotypic tests is recommended to assist in the identification of all B. cereus group species implicated in food-borne illnesses.

Reference:

• McIntyre, L.; Bernard, K.; Beniac, D.; Isaac-Renton, J. L., and Naseby, D. C.: Identification of Bacillus cereus group species associated with food poisoning outbreaks in British Columbia, Canada. Appl. Environ. Microbiol., 74, 7451-7453 (2008).
  • Abstract (courtesy of Applied and Environmental Microbiology).

Nanoprobes also offers a second negative stain, Nano-W (methylamine tungstate), which is based on tungsten rather than vanadium. While Nano-W shares the non-reactive, amorphous properties of NanoVan, because it contains the higher-Z element tungsten, it provides significantly higher contrast. The two stains provide users with a choice of negative stain contrast densities that can be used in several applications:

• NanoVan and Nano-W are completely miscible: they may be mixed in different proportions to give any desired intermediate stain density.
• Near-neutral pH results in better ultrastructural preservation.
• NanoVan is less susceptible to electron beam damage than uranyl acetate.
• Fine grain allows high imaging resolution.

More information:

• Catalog information: Negative stain reagents
• NanoVan: Product information and instructions
• Nano-W: Product information and instructions
• Technical help: Negative staining
• Negative staining: References
• NanoVan Negative stain: Our paper from Microscopy and Microanalysis 1994

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Nanogold® Particles and In Vitro Budding of Intralumenal Vesicles

In eukaryotic cells, early endosomes act as receptacles for the collection of molecules from the plasma membrane, ligands and solutes for recycling back to the plasma membrane, transportation to the trans-Golgi network, or - particularly in the case of ubiquitinated signaling receptors that need to be down-regulated - delivery to late endosomes and lysosomes. These receptors are incorporated into invaginations of the early endosomal membrane that form within their lumen, producing nascent multivesicular bodies (MVBs) with a characteristic multivesicular appearance, or endosomal carrier vesicles (ECVs), which function as intermediates between early and late endosomes. Eventually, intralumenal vesicles are delivered to lysosomes, where they and their contents are degraded. The invaginated lumenal membranes usually incorporate down-regulated EGF receptor destined for degradation, but the mechanisms that control their formation are poorly characterized. Falguières and group, in a recent issue of Molecular Biology of the Cell, report a novel, quantitative biochemical assay using 8-hydroxypyrene-1,3,6-trisulfonic acid (HPTS) incorporation for the formation of lumenal vesicles within late endosomes in vitro. They then used light microscopy, and electron microscopy with non-functional Nanogold, to determine accessibility of endocytosed components to the compartment supporting the budding process.

Experiments were conducted in baby hamster kidney cells (BHK-21) expressing human green fluorescent protein (GFP)-tagged epidermal growth factor (EGF) receptor. Cells were mock-treated or treated with siRNAs against ALG-2 interacting protein X (Alix), endosomal sorting complex required for transport type I (ESCRT-I) subunit Tsg101, or both for 3 days to generate knockdown cells. Postnuclear supernatants (PNSs) were prepared and used in for the in vitro assay: knockdown of Alix and Tsg101 was always controlled by Western blotting.

For the assay, membranes were incubated with HPTS before (protocol 1) or after (protocol 2) endosome fractionation, using either PNS (protocol 1) or isolated late endosome fractions (protocol 2). The PNS (215 µL) was adjusted to a final volume of 300 µL with 1 mM HPTS, 100 mM KCl, 12.5 mM HEPES-NaOH, pH 7.4, 1.5 mM Mg(OAc)₂ and 1 mM DTT (final concentrations) in the presence of an ATP-regenerating or -depleting system, supplemented if necessary with 1 µM bafilomycin A1 or 5 µM nigericin. This mix was incubated at 37°C, the reaction was stopped on ice, and excess 8-hydroxypyrene-1,3,6-trisulfonic acid (HPTS) quenched with 100 µL of 200 mM DPX in 20 mM HEPES-NaOH, pH 7.4 with 150 mM NaCl. Early and late endosome fractions were prepared by flotation in a step sucrose gradient. In some experiments, PNS was prepared from cells incubated with 2 mg/mL horseradish peroxidase for 10 minutes at 37°C then for 40 minutes without HRP to enzymatically label late endosomes. At the end of the assay, the reaction mixture was centrifuged at 120,000 X g, and HRP was quantified in pellets and supernatants to determine endosome latency after in vitro incubation. To measure EGF receptor incorporation in intralumenal vesicles in vitro, late endosomes from cells expressing GFP-EGF receptor were prepared on gradients 30 minutes after stimulation with EGF. The fractions were incubated for 15 minutes at 37°C with or without 250 µg TPCK (N-tosyl-l-phenylalanine chloromethyl ketone)-treated trypsin (Sigma) per mg of endosome protein. Reaction was stopped on ice by addition of soybean trypsin inhibitor. After 15 minutes, samples were processed for SDS-PAGE and Western blotting analysis. EGF receptor was quantified from the blots.

Alternatively, early and late endosomes were first prepared using the same gradient. Then, 165 µL of the endosomal fraction was incubated with 50 µL rat liver cytosol added to the assay before the reaction was stopped on ice, and excess HPTS was quenched with DPX, as above. HPTS trapped in endosomes was measured by fluorescence (lambdaexc: 413 nm/lambdaem: 510 nm), normalized to the protein content of the fractions. HPTS fluorescence (lambdaem = 510 nm) was also measured after excitation at the pH-sensitive wavelengths lambdaexc = 397 and 445 nm to measure the pH of intralumenal vesicles. The same protocol was used to measure the lumenal pH of endosomes, after 60-minute continuous HPTS internalization followed by preparation of early and late endosome fractions, as above. Fractionation of endosomes was routinely monitored using Rab5, Rab7 and LBPA. Formation of intralumenal vesicles was then visualized using electron microscopy. PNS was prepared and used in the in vitro assay (protocol 1) with 1 µM HPTS and non-reactive Nanogold particles (OD520 nm = 1.0; Nanoprobes, Yaphank, NY). Endosomes were then recovered and HPTS was quantified by fluorometry. Aliquots were concentrated by ultracentrifugation at 55,000 rpm for 30 minutes. Pellets were carefully recovered in 50 µL of 2% glutaraldehyde in PBS, incubated for 10 minutes at room temperature, and mixed with an equal volume of 4% low-melting agarose type VII (preheated at 37°C). The agarose was solidified on ice, and blocks were cut in small pieces and successively washed with PBS, 50 mM NH4Cl in PBS, PBS containing 1% BSA, and water. Gold enhancement was performed for 5 minutes at room temperature, and after washes, samples were processed for electron microscopy. After this enhancement time, gold particles showed a heterogeneous size distribution, which was attributed to variability in gold particle accessibility to the enhancer. In some experiments, the endosome lumen was prelabeled in vivo after continuous endocytosis for 90 minutes at 37°C of 15-nm BSA-gold (OD520 nm = 10) in medium containing 1% BSA. Cells were then used as above in the in vitro assay using Nanogold. Gold enhancement was then conducted for only 1 minute at room temperature, so that the diameter of enhanced gold particles remained close to 10 nm. Alternatively, endosomes were prelabeled in vivo with anti-EGF receptor gold particles. Cells expressing the EGF receptor were serum-starved and incubated on ice with monoclonal antibodies against the receptor, followed by 200 µg/ml EGF and 15-nm protein A-gold. The complex was endocytosed for 50 minutes at 37°C, PNS was prepared and used in the in vitro assay with Nanogold (gold enhancement for 1 minute, as above). For quantification, the number of gold particles per profile (inside or outside) was counted in 90 different profiles from three independent experiments for each siRNA condition, and 60 different profiles from two independent experiments for EGF receptor labeling, respectively. Results were expressed as averages of numbers of gold particles per isolated structure. The perimeter of isolated late endosomal structures after siRNA was determined for 20 isolated structures from three independent experiments using the NIH ImageJ software. The process of vesicle budding into the endosome lumen was found to be time-, temperature-, pH-, and energy-dependent, and required cytosolic factors and endosome membrane components. Light and electron microscopy data showed that the compartment supporting the budding process was accessible to endocytosed bulk tracers and EGF receptor, since both were localized inside the vesicles. However, 15 nm particles were incorporated less well, suggesting that the neck of new invaginations was too small to admit them. The EGF receptor became protected against trypsin, and therefore, it was inferred that it was sorted into the intraendosomal vesicles formed in vitro. Formation of intralumenal vesicles was also inhibited by the K173Q dominant negative mutant of hVps4, and was therefore ESCRT-dependent. Tsg101 and its partner Alix were found to control intralumenal vesicle formation, by acting as positive and negative regulators, respectively. It was concluded that budding of the limiting membrane toward the late endosome lumen leading to intraendosomal vesicle formation is controlled by the positive function of Tsg101 and the negative function of Alix.

Mechanism of enhancement of Nanogold® by GoldEnhance, showing deposition of metallic gold from solution onto gold nanoparticles. Final particle size is controlled by enhancement time.

Reference:

• Falguières, T.; Luyet, P. P.; Bissig, C.; Scott, C. C.; Velluz, M. C., and Gruenberg, J.: In vitro budding of intralumenal vesicles into late endosomes is regulated by Alix and Tsg101. Mol. Biol. Cell., 19, 4942-4955 (2008).
  • Abstract (courtesy of Molecular Biology of the Cell).

More information:

• Nanogold labeling reagents: Catalog information
• Nanogold particles (non-functional): Instructions and product information
• Nanogold reagents: technical help
• Gold enhancement: General and catalog information
• GoldEnhance EM: Instructions and product information
• Gold enhancement: Technical help
• Gold enhancement: References

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Nanoprobes: Holiday Hours and Contacts

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

We would like to extend our thanks to all of our customers who have helped make 2008 another successful year of growth for us. We look forward to bringing you more unique and exciting new products and technologies in 2009.

More information:

• Contact information
• Nanoprobes: news and press releases
• About Us: company information

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Other Recent Publications

Reference:

• Sou, Y. S.; Waguri, S.; Iwata, J.; Ueno, T.; Fujimura, T.; Hara, T.; Sawada, N.; Yamada, A.; Mizushima, N.; Uchiyama Y.; Kominami, E.; Tanaka, K., and Komatsu, M.: The Atg8 conjugation system is indispensable for proper development of autophagic isolation membranes in mice. Mol. Biol. Cell., 19, 4762-4775 (2008).
  • Abstract (courtesy of Molecular Biology of the Cell). ).

Reference:

• Schneider, G. F., and Decher, G.: From "Nano-bags" to "Micro-pouches". Understanding and Tweaking Flocculation-based Processes for the Preparation of New Nanoparticle-Composites. Nano Letters, 8, 3598-3564 (2008).
  • Abstract (courtesy of ACS Publications).

One of the most active areas in microscopy at the moment is the development of optical methods to beat the conventional resolution limit imposed by the wavelength of visible light. In another recent paper in Nano Letters, Guoxin Rong and group used plasmon coupling between individual gold nanoparticle labels to monitor subdiffraction limit distances in live cell nanoparticle tracking experiments. While the resolving power of our optical microscope is limited to about 500 nm, the new method improved upon this by more than an order of magnitude, by using a ratiometric detection scheme to detect plasmon coupling between individual gold nanoparticle labels. Plasmon coupling microscopy was successfully applied to an experiment to resolve the interparticle separations during individual encounters of gold nanoparticle labeled fibronectin-integrin complexes in living HeLa cells: cells were first incubated with a 0.2 mg/mL solution of fibronectin in imaging buffer for 10 minutes, then washed with an excess of imaging buffer and a solution of antifibronectin-functionalized 40 nm gold nanoparticles in imaging buffer. A sufficiently low concentration of gold nanoparticles was chosen sufficiently to allow the tracking of individual particles. Detection relies on the shift of the plasmon resonance to greater wavelengths as the particles approach, and fitted light scattering images obtained on two filter channels, 530 nm and 580 nm. A minimum detection range, or particle separation, of 15 nm was found, more than an order of magnitude greater than the wavelength of visible light.

Reference:

• Rong, G.; Wang, H.; Skewis, L. R., and Reinhard, B. M.: Resolving sub-diffraction limit encounters in nanoparticle tracking using live cell plasmon coupling microscopy. Nano Letters, 8, 3386-3393 (2008).
  • Abstract (courtesy of ACS Publications).

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