Markers for Correlated Light and Electron Microscopy

CHAPTER 23

Markers for Correlated Light and Electron Microscopy Gina E. Sosinsky, Ben N. G. Giepmans, Thomas J. Deerinck, Guido M. Gaietta, and Mark H. Ellisman National Center for Microscopy and Imaging Research and Center for Research in Biological Systems University of California, San Diego, La Jolla, California 92093

I. Introduction II. How Do LM and EM Complement Each Other? III. Fluorescence Photooxidation A. Principles B. Small, Organic Fluorophores C. Eosin-Based Fluorescent Probes D. Genetic-Based Tags IV. Enzymatic-Based Methods V. Particle-Based Methods for Protein Localization A. Gold Particles B. Quantum Dots VI. Concluding Remarks References

The dynamic behavior of cells is a consequence of the coordinated and elaborate interactions between complexes of macromolecules that constitute their formed structures or organelles. Live-cell imaging and high-resolution determination of the location of nucleic acids, proteins, metabolites, and ions can now be made within subcellular domains down to the molecular level, revealing the important information required to understand complex cellular functioning. Light and electron microscopic methods have proved extremely powerful in providing information about cells, and techniques such as immunolabeling have been applied successfully to identify molecules in situ. Recently, developments in recombinant methods of modern molecular biology and synthetic chemistry have lead to new probes and targeting techniques for these METHODS IN CELL BIOLOGY, VOL. 79 Copyright 2007, Elsevier Inc. All rights reserved.

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0091-679X/07 $35.00 DOI: 10.1016/S0091-679X(06)79023-9

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correlated modalities. Here we review whole cell, organelle, and protein labeling methods for correlated observations in the spatial domains of both light and electron microscopy.

I. Introduction A continuing challenge to structural biologists is the elucidation of large macromolecular complexes and organelles, including multicomponent combinations of proteins, membranes, and/or nucleic acids. We have coined and have popularized the term ‘‘mesoscale’’ (from the Greek ‘‘mesos,’’ middle) as the imaging range we mine to obtain new knowledge about the structure and dynamics of the macromolecular aggregates that compose the functional complexes within cells, tissues, and organs. These mesoscale structures are imaged with light microscopy (LM) and with more detail with electron microscopy (EM) at moderate resolutions, achievable with numerous preparatory methods, as opposed to high resolution, atomiclevel, 3D structural methods, such as nuclear magnetic resonance (NMR), x-ray crystallography, and molecular microscopy (e.g., electron cryomicroscopy). In addition, along with the task of creating systems biology computational approaches to protein–protein interactions (the ‘‘interactomes’’ Rual et al., 2005), one would ideally like to place macromolecular complexes within their cellular environments in order to fully understand their functional interactions (‘‘visual proteomics,’’ Nickell et al., 2006). The focus of this chapter is the visualization of cells and organelles, and the identification of molecules for correlative LM and EM.

II. How Do LM and EM Complement Each Other? One way of highlighting proteins of interest is to attach probes to them. For LM, this most often involves the tagging of molecules with fluorophores (Giepmans et al., 2006) or nonfluorescent stains, such as peroxidase-mediated precipitates. EM markers must be highly electron dense in order to be distinguished from background constituents. Such markers can be visualized by EM tomography to provide information about the 3D ultrastructure of tissues, cells, and macromolecular complexes (as reviewed in Baumeister et al., 1999; Frey et al., 2006; McEwen and Marko, 2001; Sosinsky and Martone, 2003; Subramaniam, 2005). The combination of EM tomographic reconstructions with labeling of specific components provides a powerful method for localizing proteins within their cellular components. The application of correlated LM and EM is leading the way in revolutionizing our understanding of the structural basis of cell biology. Better fluorescent probes have been and are being developed that highlight a particular protein of interest, and better LM instrumentation allows us to monitor these probes in three dimensions (3D) and real time (4D-imaging). Developments in fluorescence LM provide great opportunities to determine protein colocalization, turnover, and function (reviewed in

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Giepmans et al., 2006). However, imaging at the resolution of the LM provides only a general cellular location of the probe, a size and shape, and a sense of how the probe moves during sequential data acquisition. The relatively low resolution it provides might lead to a conclusion of colocation of two probes or the localization of a marker to a specific organelle that does not reflect the actual in situ situation. For example, a molecule or complex that sits just under the plasma membrane could be misinterpreted in LM imaging as being localized in the plasma membrane. While technological advances in LM imaging are pushing the resolution beyond the classical limit of 0.1–0.2 mm (Garini et al., 2005), there are great additional benefits from probes that can also be imaged at the EM level. The shorter wavelength of the electron particle beam oVers a substantial increase in resolution, and it also provides valuable information about the location of probes within the full cellular context; in addition to the probe, one sees many subcellular structures (e.g., organelles), membrane systems (e.g., plasma membrane), and macromolecular complexes (e.g., ribosomes), all at high resolution. Theoretically, this resolution increase when based only on instrumentation parameters is >1000-fold, but practically speaking, for biological specimens this ratio is
40- to 100-fold improvement. EM samples are inanimate, static structures. A well-known neuroscientist, Floyd Bloom, once went so far as to say ‘‘the gain in the brain lies mainly in the stain’’ (personal communication), but it is worth noting that this concept of improved protein detection is applicable to many fields of biological EM. Therefore, new probes to localize molecules at both the LM and EM levels have great potential to combine the benefits of the two techniques. In this chapter, we divide the correlative LM and EM probes aimed at obtaining mesoscale structure and dynamics into three categories: (1) fluorescence photoconversion, (2) enzymatic-based detection, and (3) particle-based methods for protein localization. Each of these techniques has its own advantages and drawbacks; in reality, an investigator needs to decide ahead of the experiment as to the kind of information that s/he wishes to obtain. A flowchart of the experimental approaches employed in determining the localization of proteins in diVerent samples and conditions is shown in Fig. 1.

III. Fluorescence Photooxidation A. Principles Photooxidation is a powerful and versatile technique for LM and EM whereby the oxygen radicals generated by a fluorescent compound under illumination are used to drive the oxidation of diaminobenzidine (DAB; see schematic in Fig. 2). Oxidized DAB can be made electron dense through osmification. Table I summarizes the photooxidation experiments with small fluorophores that are described in the following sections. Photooxidation of DAB was first shown with Lucifer Yellow (Maranto, 1982) in whole cells. Photooxidation of highly permeant

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Fig. 1 Flowchart of the methods for correlative LM and EM described in this chapter. These include photooxidation, enzymatic-based detection, and particle-based methods. See text for details.

Singlet excited state

Triplet excited state

Xenon lamp

Localized polymer stainable with OsO4

Reactive oxygen EOSIN-5-ITC Br HO

Br O

O

Br

H2N

Br

NH2 NH2

H2 N

=

C-OH O

Ordinary O2

DAB

N=C=S

Fig. 2 Schematic of the photooxidation process. The process reads left to right, first with illumination by a xenon source that excites the photooxidizer to create reactive oxygen species that reacts with DAB to form an osmiophilic polymer.

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Table I Major Classes of Small Fluorophores for Photoconversion and LM and EM Analysis Dye

Target

Lucifer Yellow

Entire cell (microinjection)

Fluoro-Ruby DiI PI BODIPY

Anterograde axon transport Membranes Nucleic acids Conjugates (like BODIPY ceramide: Golgi)

FM1-43

Plasma membrane and transport vesicles

Eosin

Immuno-conjugates to identify proteins

ReAsH

Oligonucleotides conjugates to identify nucleic acids Ligand conjugates: Phalloidin-conjugate to identify F-actin a-Bungarotoxin to nictonic acetylcholine receptor Genetically targeted to identify proteins

Reference(s) Maranto, 1982 Bushong et al., 2002 Schmued and Snavely, 1993 von Bartheld et al., 1990 Balercia et al., 1992 Pagano et al., 1989; Takizawa et al., 1993 Brumback et al., 2004 Henkel et al., 1996 Harata et al., 2006 Richards et al., 2000 Schikorski and Stevens, 2001 Fomina et al., 2003 Deerinck et al., 1994; Hand et al., 2001 Huang et al., 1994 Capani et al., 2001 Martone et al., 1999 Gaietta et al., 2002 Sosinsky et al., 2005 Martin et al., 2005 Beahm et al., 2006

Typical applications of the diVerent dyes are summarized. See text for further details.

fluorophores (Lucifer Yellow, BODIPY ceramide) or reagents (FlAsH/ReAsH), in combination with DAB and reactive oxygen, possesses several advantages over other methods for high-resolution protein localization studies: (1) due to the small size of the reagents, the penetration of labels into the tissue is substantial, even without the use of harsh pretreatments such as detergents; (2) it can be used for correlated LM and EM; (3) due to the limited spread of reaction product, the spatial resolution under the EM is vastly superior to enzymatic methods; and (4) whereas photoconversion specifically stains the target protein, particle methods [gold or quantum dots (QDs)] decorate it. Moreover, in case of tetracysteine (Cys4) labeling the photooxidation product is directly on the protein. For gold labeling or eosinmediated photoconversion, the ‘‘tree’’ of antibodies or other targeting reagents is a complex typically >10 nm in diameter. Following treatment with osmium, the photooxidized DAB reaction product exhibits uniform staining with little granularity. The positively stained, photoconverted material often has a ‘‘negative-stain’’-like appearance such that macromolecules appear light, but are surrounded by electron density. This technique provides

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specifically labeled, thick (>0.1 mm) specimens for high-resolution, 3D correlated confocal and high voltage microscopies (e.g., EM tomography). Standard EM techniques such as colloidal gold (CG) immunolabeling or immunoperoxidase are usually suboptimal for thick sections, because the label cannot penetrate thick sections unless the cell ultrastructure is highly compromised. B. Small, Organic Fluorophores Small organic dyes and compounds that either diVuse easily, or naturally intercalate into membranes or organelles, have been useful for generating the reactive oxygen species critical to good photooxidation (Table I). Maranto (1982) demonstrated the usefulness of photooxidation with neurons injected with the fluorescent tracer dye, Lucifer Yellow. In that study, horseradish peroxidase (HRP) was used to label synapses in combination with Lucifer Yellow. Lucifer Yellow (molecular weight 457 D) diVuses easily through entire cells, and thus delineates the cell’s expanse. This technique was recently used to show that astrocyte processes establish exclusive territories and are much more expansive than had been shown with standard immunolabeling of an astrocyte marker (Bushong et al., 2002). Similarly, Fluoro-Ruby, a fluorescent tetramethylrhodamine-dextran amine, used to demonstrate anterograde axon transport, has been successfully photoconverted and examined by EM (Schmued and Snavely, 1993). The membrane stain, 1,10 -dioctadecyl-3,3,30 ,30 tetramethylindocarbocyanine perchlorate (DiI), is a fluorescent dye that diVuses within cell membranes and can be photoconverted (von Bartheld et al., 1990). Propridium iodide (PI), a fluorescent compound that intercalates into nucleic acid structures stains the nucleus and can be photoconverted (Balercia et al., 1992). Fluorescent conjugates of ceramide, such as BODIPY ceramide, accumulate at the Golgi apparatus of fixed cells and can be photoconverted to selectively stain this organelle (Pagano et al., 1989; Takizawa et al., 1993). Some classes of fluorescent lipid dyes intercalate into smaller vesicles and can then be photoconverted for EM analysis. One such dye is FM1-43, a dye that becomes highly fluorescent when partitioning into a hydrophobic environment, such as membranes; it has therefore been instrumental in monitoring exocytosis, endocytosis, and endosomal traYc (Brumback et al., 2004; Henkel et al., 1996). The charged FM1-43 is applied to the outside of the cell; it partitions into the outer leaflet of the plasma membrane, but it cannot flip-flop or diVuse across the membrane. Because surfaceexposed FM1-43 can be washed out, the endosome-trapped dye can be used to specifically stain endosomes at the LM level and, following photoconversion, at the EM level. This technique has proven useful for identifying intermediates of the synaptic vesicle cycle and in the recycling of these membrane components (Harata et al., 2006; Henkel et al., 1996; Richards et al., 2000; Schikorski and Stevens, 2001). FM1-43 has also been used to investigate the kinetics of constitutive endocytosis and to visualize the fate of endocytic organelles in resting and activated human T lymphocytes. Internalized cargo was carried to lysosome-like compartments in resting T cells and to multivesicular bodies in activated T cells (Fomina et al., 2003).

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C. Eosin-Based Fluorescent Probes Eosin, a tetrabrominated derivative of fluorescein, is a potent generator of singlet oxygen while still possessing reasonable fluorescence. Eosin has a quantum yield of 0.57 for singlet oxygen generation, however the fluorescence quantum yield of eosin is typically only about 10–20% that of fluorescein (Deerinck et al., 1994). Eosin can also be conjugated to various molecules, such as antibodies, phalloidin, and a-bungarotoxin, to achieve high-resolution protein localization at the fluorescence LM and EM levels (Martone et al., 1999). Capani et al. (2001) used phalloidin conjugated to eosin and photooxidation to investigate the actin cytoskeleton in dendritic spines. Phalloidin selectively distinguished among diVerent populations of dendritic spines, both within and between brain regions (Capani et al., 2001). In addition, streptavidin–eosin conjugates enabled the photoconversion of RNA probes (Huang et al., 1994) for EM analysis of the distribution of poly(A)-RNA in mammalian cells. Despite its advantages, photoconversion suVers from a lack of sensitivity compared to enzyme-based methods—which ‘‘amplify’’ the signal by churning out an often diVusible reaction product. In fact, for most immunolabeling procedures, the standard fluorescent dyes fluorescein isothiocyanate (FITC) and rhodamine cannot be used because they are poor generators of singlet oxygen and thus are too ineYcient at driving the polymerization of a reactive compound like DAB (Deerinck et al., 1994). As outlined above, eosin oVers a reasonable compromise between fluorescence yield and oxidation of DAB. However, even with this more potent oxidizer, the fluorescent signal (local concentration of target molecules) may not be strong enough to drive adequate photooxidation when stronger aldehyde fixation is used for maintaining ultrastructure for EM imaging (Hand et al., 2001). We found that the immunofluorescence of many commercial connexin43 antibodies decreased dramatically even with small amounts of glutaraldehyde. Using tyramide signal amplification methods can modulate some of the antibody fixation sensitivity. These tyramide amplification techniques have also been successfully implemented for enhanced postembedding immuno-based CG labeling of proteins in pancreatic and liver tissues (Mayer and Bendayan, 1999). While there is a small decrease in the precise localization of labeling, due to the enzymatic nature of the amplification and the larger size of the immunolabeling reagents, we demonstrated that the labeling for connexin43 gap junctions was specific and provided specimens suitable for 3D localization by EM tomography (Hand et al., 2001).

D. Genetic-Based Tags A focus of further development for the fluorescence photooxidation technology is the application and invention of fluorescent genetic probes with photooxidation capabilities that are appended to or inserted into proteins of interest (see also Giepmans et al., 2006). Fluorescent proteins (FPs) have provided a wealth of LM data, but genetically encoded tags for EM are just being developed.

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Green fluorescent protein (GFP) is the most widely used fluorescent genetic tag, but GFP has been used for photoconversion of DAB in only a couple of situations (Grabenbauer et al., 2005; Monosov et al., 1996) because the reactive oxygen generation is much lower than that of ReAsH and may only work for situations where it is expressed at extremely high concentrations and by forced packing, changing its own properties, such that singlet oxygen can be generated. In addition, FPs (
26 kDa) are often larger than the protein of interest. While not necessarily a significant limitation in cell biology, the large size of FPs can interfere with the distribution, function, and fate of some recombinant proteins (reviewed in Giepmans et al., 2006). Recombinant proteins in intact cells can also be labeled by genetically appending or inserting a small motif (6–20 residues) that encodes a Cys4 sequence -Cys-Cys-Pro-Gly-Cys-Cys-, then exposing the cells to a membrane-permeant, nonfluorescent, biarsenical derivative of fluorescein, FlAsH. Roger Y. Tsien and his colleagues (Adams et al., 2002; GriYn et al., 1998) developed this system originally for live-cell imaging. FlAsH binds with high aYnity and specificity to the Cys4 motif and thereby becomes strongly fluorescent, emitting green light when excited with blue light. These fluorescent complexes can survive for days in the absence of excessive (mM) concentrations of competing 1,2-ethanedithiol (EDT) (Martin et al., 2005). A biarsenical resorufin derivative, ReAsH, has been shown to photoconvert DAB (Gaietta et al., 2002). Because the labeling is carried out in live cells, as described above for FlAsH, preparation for EM allows stringent fixation and does not require permeabilization. This preserves the ultrastructural aspects of the specimen. In principle, application of cryofixation methods to these labeled specimens would be highly advantageous. In practice, however, cells or tissues being photoconverted cannot be fast frozen and then photooxidized. Nonetheless, we have demonstrated that aldehyde fixation used prior to rapid freezing provides very good ultrastructural preservation (Sosinsky et al., 2005). The combination of aldehyde fixation prior to high pressure freezing is one aspect of future development of photoconversion methods. The Cys4/biarsenical ligand technique allows pulse-chase labeling at the fluorescence LM and EM level (Gaietta et al., 2002) to address questions about how proteins move through the various cell compartments during their life cycle. Examples of optical pulse-chase applications include the traYcking of the gap junction protein, connexin43, (Gaietta et al., 2002) or the localization of protein synthesis for the a-amino-5-hydroxy-3-methyl-4-isoxazole propionic acid (AMPA) receptor, glutamate receptor 1, GluR1 (Ju et al., 2004). AMPA receptors are named for their synthetic agonist; they include glutamate receptors (GluR). This class of receptors is the most commonly found receptor in the nervous system, and therefore, proper traYcking of their subunits (e.g., GluR1) is an important consideration in synaptic remodeling. The proper traYcking of gap junction proteins has great relevance to the various connexin diseases, because many of these mutations cause mistraYcking of the protein.

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A peptide with higher aYnity for ReAsH (Martin et al., 2005) and GFP combined with ReAsH have both been used to increase the specificity of ReAsH labeling to Cys4 and to increase specific photooxidation at the ReAsH-labeled chimeric protein by excitation of ReAsH through FRET from the donor GFP (Beahm et al., 2006; Martin et al., 2005). These samples are amenable to EM tomographic imaging (Sosinsky et al., 2003) and can be used to follow Golgi components through mitosis (Gaietta et al., 2006). Whichever method is used for photoconversion, DAB precipitation is most easily detectable on structural compartments or proteins that are concentrated in the cell. While there are no published quantifications of the concentration of label and size, we have some estimates based on the Cys4-tagged specimens we have developed. For example, 180 copies of a viral protein tagged with a Cys4 domain and labeled with ReAsH are enough for good photoconversion of a
30-nm diameter virus (Lanman et al., personal communication). In addition, individual actin filaments made from native and Cys4-tagged actin monomers are also easily photoconverted (Giepmans et al., personal communication). Since this technique does ‘‘paint’’ individual labeled macromolecules like hemichannels with as few as six Cys4–ReAsH molecules, making them directly visible in the TEM—there is hope for further improvement in sensitivity, if the signal-to-noise ratio (SNR) can be improved. FRET-based photoconversion of ReAsH has promise here (Martin et al., 2005). For more diVuse proteins, enzymatic or particle-based markers might be preferable.

IV. Enzymatic-Based Methods A more traditional technique for visualizing proteins within cells involves the use of enzyme-based systems, typically in combination with immunolabeling (Etemadi, 1980; Farr and Nakane, 1981; van Leeuwen, 1981). Enzyme-based methods were first introduced in the mid-1960s and continue to be used today. The benefits of these methods are that they are straightforward and cheap, they penetrate well into the sample, and they have higher sensitivity than eosin-based methods. Commercial kits are available for ease of use (e.g., from Pierce, Clinisciences, or Molecular Probes, Inc.). These methods are still a valuable complementary approach to fluorescence photooxidation. The two enzymes most commonly used for this work are HRP and alkaline phosphatase. These enzymes can be targeted to specific proteins by antibodies or by ligands conjugated to an enzyme. HRP catalyzes a reaction that uses DAB and H2O2 as substrates; as with fluorescence photooxidation, the DAB forms an amorphous polymer that is readily stained with OsO4 (see above). With this method, one can observe the same preparation with nonfluorescent LM and EM. However, the diVusion of the reaction product away from the reaction site and onto membranous structures is a significant drawback, decreasing the resolution of the labeling (Deerinck et al., 1994). The enhanced resolution of the photoconversion reaction product versus that generated by peroxidase is probably the result of two factors.

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A

Eosin conjugated secondary antibody

DAB + O2 + light

B

C

DAB + H2O2

Streptavidin peroxidase complex

PPT B

PPT Primary antibody

PPT Antigen Eosin photoconversion

B

DAB + O2 + light Protein of interest

Biotinylated secondary antibody

Tetracysteine domain/ReAsH complex

Tetracysteine/ReAsH photoconversion

Antigen Peroxidase-mediated DAB deposition

Fig. 3 Comparison of relative labeling sizes for three methods discussed in this chapter that use DAB polymerization as a labeling strategy. In each of these labeling procedures, DAB is used in combination with reactive oxygen species to form an osmiophilic precipitate that in combination with osmium tetroxide forms an electron dense substance. (A) Eosin photoconversion involves the binding of an eosin-conjugated ligand. In this case, eosin is bound to a primary antibody, but this method can also be used with eosin-conjugated toxins or ligands. (B) A Cys4 domain is genetically appended to a protein of interest and the biarsenical ligand, ReAsH, is bound to the Cys4 domain. The ReAsH serves as the singlet oxygen generator. (C) Schematic of peroxidase-mediated DAB polymerization (ABC methods) involving the use of biotin, streptavidin, and HRP. While this method may provide more sensitivity than the photooxidation methods (A or B), the resolution (size of the precipitate) is not nearly as precise.

First, the photoconversion reaction is carried out after postfixation with high concentrations of glutaraldehyde. This fixation increases the relative cross-linking of the cytoplasm and reduces the ability of the reaction product to diVuse. In addition, when singlet oxygen is generated by a point source (the fluorophore), the radius of influence is limited in our estimations to the single nanometer range, and the DAB appears to form a polymer in that limited domain. This was shown with eosin-based photoconversion of antibody labeled microtubules in 1994 (Deerinck et al., 1994). Finally, photoconversion is done on ice. This also reduces the likelihood of diVusion of the reaction product. These interpretations are supported by the images shown in Fig. 4C and H. Figure 3 contains a schematic comparing the relative sizes and EM staining processes for (A) eosin-based photooxidation (B) Cys4/ReAsH-based photooxidation and (C) peroxidase (enzymatic)-based reactions.

V. Particle-Based Methods for Protein Localization A. Gold Particles Colloidal Gold (CG) bound to antibodies, soluble proteins, or ligands remains the standard method for particle-based methods for protein localization at present. As such, there are many good references for the use of CG as an EM marker (see Burry, 2000; Hayat, 1990; Roth, 1996). Here, we will discuss only recent technological

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improvements in CG labeling for correlative fluorescent LM and EM. Depending on the protein to be imaged and detectability required for EM, the size of CG typically used is 5–20 nm. While such gold particles are instantly recognizable in the EM, they are diYcult to introduce into cells and tissues unless cells are permeabilized with detergents, whereupon the cell’s ultrastructure is compromised. Exceptions to this are cell surface receptors, where permeabilization is not required for ligand binding. An example of this approach was an electron tomographic reconstruction of the ciliary ganglion in chick lens after marking of the cell surface for specific acetylcholine receptors (Shoop et al., 1999). Because of these problems, most published studies that use CG for intracellular labeling apply the tagged probe after embedding rather than using a preembedding protocol; they are stand-alone EM experiments. A major benefit of this approach is the ability to use diVerent sizes of gold simultaneously, allowing for localization of multiple distinct targets with clearly defined labels in the same specimen. Progress in gold probes for correlative microscopy involves small gold labels (
1–3 nm) that can be increased in size by gold ‘‘toning’’ or silver ‘‘enhancing’’ procedures on thin sections for better detection (Baschong and Stierhof, 1998; Hainfeld and Furuya, 1995; Robinson et al., 1998). These labels have also been made fluorescent by covalently binding FITC or Cy3 to the Fab. Probes like this, referred to as ‘‘Fluoronanogold,’’ use an
2-nm gold particle (Nanogold) (Powell and Hainfeld, 2002; Takizawa and Robinson, 2000a,b; Takizawa et al., 1998). Nanogold is a commercial product of Nanoprobes, Inc. (Yapank, NY, http:www. nanoprobes.com; Hainfeld and Powell, 2000; Powell et al., 1997, 1998). Applications of this technology for correlated LM and EM or EM tomography include that labeling of microtubules in leukocytes (Robinson and Vandre, 1997), localization of pKi-67, a putative chromatin-binding protein of unknown function (Cheutin et al., 2003), release of interleukin-4 from granule distinct vesicular compartments (Melo et al., 2005), and localization of the voltage-gated Kþ Channel, Kv4.2, in T-tubules of rat myocytes (Takeuchi et al., 2000). However, nanogold is not nearly as visible in the EM as larger sized gold particles; it typically requires an amplification step, such as silver or gold intensification, that no longer allows for precise discriminative labeling of multiple distinct of the protein targets. QDs show great promise in their preembedding multiprotein labeling abilities (see below). B. Quantum Dots QDs are inorganic nanocrystals that fluoresce at distinct wavelengths depending on their size (
4–10 nm) and shape. The core, typically a CdSe or CdTe crystal, is electron dense and enables discrimination of the distinct QDs at the EM level (Giepmans et al., 2005). A water-soluble coating allows for conjugation to targeting molecules, such as antibodies. While QDs are excellent fluorophores for LM, they are also a powerful marker for protein detection at the EM level (reviewed in Giepmans et al., 2006). The drawback of antibody-mediated targeting of QDs to cytoplasmic epitopes, as with all preembedding immuno-based techniques, is that

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B

D

E

G

H

C

F

Fig. 4 Comparison of immunolabeled microtubules stained by photooxidation, particle-based methods, and enzymatic methods. Monolayers of cultured cells were immunolabeled for the microtubule protein tubulin followed by eosin-conjugated secondary antibodies (A–C) or QD-conjugated secondary antibodies (D–F). Fluorescence LM imaging (A, D) reveals many microtubules (A: green, D: red; nuclei are counterstained in blue). (B) Eosin-mediated photooxidation of DAB leads to a fine, osmiophilic precipitate that evenly increases the density of the microtubules in EM images. A higher magnification view is shown in (C). (E) QDs are large and electron dense enough to image at the EM at near-molecular

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permeabilization is required, and these treatments have a negative impact on the cellular ultrastructure. Compared to CG, QDs have a relative low electron density, which can be increased by silver enhancement, as has been successfully applied to detect glycine receptors in synapses (Dahan et al., 2003), but now multilabeling as outlined earlier is no longer possible. Postsectioning labeling of a nuclear protein has also been shown with QD and correlated microscopies (Nisman et al., 2004). We recently reported a generic application to simultaneously label multiple endogenous proteins for correlated LM–EM in cells and tissues using preembedding immunolabeling (Giepmans et al., 2005). Proteins in cells and tissues were immuno-double or immuno-triple labeled using QDs and visualized in the LM and EM. Preembedding labeling using QDs penetrated tissues more eYciently than conventional 5-nm CG, but not nearly as well as small dyes such as FITC that are suitable solely for LM. QDs allow for labeling with several micrometer penetration, as opposed to immunogold (Giepmans et al., 2005, 2006). Therefore, QDs can be successfully used to label proteins in thick EM-sections for immuno-tomography (Giepmans et al., our unpublished observations). Since the diVerent fluorescent emissions of QDs can be correlated with diVerent sizes or shapes of QDs, which are discernable in the EM, double or triple staining of proteins followed by correlated microscopy is relatively straightforward. Moreover, labeling eYciency can be assessed by fluorescence at the LM level before continuing with more laborious EM. For high-throughput EM, large sections of labeled tissues can be screened at LM level, followed by a subsequent EM analysis of regions of interest. As an alternative to immuno-based techniques, QDs can be targeted to molecules of interest using other conjugates such as streptavidin (reviewed in Giepmans et al., 2006).

VI. Concluding Remarks In summary, new tools have been developed for correlated LM and EM (e.g., photooxidation of fluorescent probes and immuno-QD labeling); these complement more established techniques, such as peroxidase or CG labeling. A comparison of these methods has been achieved by using a standard structure, like microtubules, that were immunolabeled for tubulin and then detected by photooxidation (Fig. 4A–C), QDs (Fig. 4D–F), CG (Fig. 4G), and enzymatic labeling (Fig. 4H). Such images show the diVerent appearances of these labeling strategies. In our experiences, the SNR for enzymatic methods is better than for photooxidation, and QDs are better than standard CG, because of better penetration of the particles. However, to compare the SNR of QDs with Cys4 photooxidation is not a valid comparison, because while QDs give far better fluorescence than eosin or ReAsH, but resolution. The image in (F) is a higher magnification view. Older methods: a 5-nm immunogold microtubule labeling (G) and peroxidase-mediated enzymatic labeling (H) are shown for comparison. Black arrows in this figure point to individual microtubules. For scale, the diameter of a native microtubule is
25 nm.

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they do not label the tagged protein complex with reaction product. The applicability and implementation of these techniques to any particular biological problem depends on the exact settings and goals, like the viewing of a single versus multiple distinct targets, the desired resolution, the ease of use, desired integrity of cell architecture, and the detectability of features. While each of these techniques has advantages and disadvantages relative to the others, each method can provide important and significant information about the dynamics, localizations, and arrangements of proteins within their cellular context.

Acknowledgments We are grateful to Drs. Maryann E. Martone and Roger Y. Tsien for invaluable discussions. Some of the work described in this review was supported by grants from the National Institutes of Health grant NIH-RR004050, NS 14718, RR019701, DK54441 (M.H.E.), GM072033 (Roger Y. Tsien), GM65937 (G.E.S) GM72881 (G.E.S.), and NSF grant MCB-0543934 (G.E.S.).

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