The answer is no. Although the use of buffered solutions prior to staining and sometimes after staining is feasible, both the in vivo tracers and the histochemical tracers should be reconstituted in either distilled water or saline, as indicated.
Yes, all tracers are compatible with all of these tissue processing procedures. However, formalin fixed tissue typically reveals the more detailed morphology. Paraffin processing can shift the emission spectrum of Fluoro-Gold from a light yellow to a light blue color.
No, making them suitable for multiple labeling studies.
No, all tracers are quite stable and exhibit only negligible fading as a result of prolonged excitation or archival storage.
Most tissues can be dehydrated by either by passing them through graduated concentrations of ethanol, or else air dehydrated for a few min on a slide warmer or hot plate at 60-70°C. We find the latter method to be the simplest, fastest and to result in optimal contrast and resolution staining. Unless counter-indicated, dehydrated slides are then cleared in xylene for 1-2 min and then coverslipped with a non-polar non-fluorescent mounting media such as DPX.
By the proper gelling and drying of the slides, the tissue sections can be prevented from falling off the slide even through relatively harsh solutions. The following gelling procedure is strongly recommended:
The rate for both anterograde and retrograde axonal transport is around 1 cm/day in warm blooded animals. Therefore, an appropriate survival time for small animals such as rodents would typically be in the 2-4 day range, while large animals might require a 1-2 week survival interval, depending on the length of the pathway being studied. It is worth noting that both tracers are quite resistant to degradation and therefore long survival intervals can be used to optimize labeling.
No, although Fluoro-Gold is typically used to demonstrate retrograde axonal transport while Fluoro-Ruby is generally used to demonstrate anterograde transport, both tracers are capable of undergoing bidirectional transport. However, Fluoro-Gold must bind to cytoplasmic nucleic acids to become highly fluorescent, making anterograde transport difficult to detect. However, antibodies to Fluoro-Gold have been developed which will allow localization of anterograde transport as well. Conversely, Fluoro-Ruby is generally used to demonstrate anterograde axonal transport, however some retrogradely labeled cells can also be seen. One limitation to using this tracer as a retrograde tracer is that the degree of labeling is system specific, with some neurons labeling more conspicuously than others.
The stereotaxic pressure injection of these tract tracers is typically accomplished via a small micro-syringe (e.g. 1 ul microsyringe). Small animals typically receive a 0.05 - 0.2 ul injection while large animals could receive an injection volume between 0.2 - 1 ul. Fluoro-Gold is usually used at a concentration between 1-2%, while Fluoro-Ruby is used at a concentration of 10% in saline vehicle. Higher concentrations of Fluoro-Gold can cause local degeneration at the site of injection. Both tracers may also be delivered iontophoretically when highly discrete injections are wanted.
Yes, it has been injected into muscles to label motor neurons in the spinal cord and brain and has also been administered systemically to label hypothalamic cells with projections to the pituitary. Systemic injection has also been reported to result in labeling of the lumen of all vascular endothelial cells.
The first sign that the stain is becoming depleted is an increase in staining time. The staining time can be increased by up to 20% to compensate for this, however even longer times are generally ineffective. Another sign of dye depletion is the appearance of a fine black precipitate at the bottom of the staining container. Still, the presence of a limited amount precipitate does not mean solution is completely depleted. Another sign that the solution is being depleted of stain is it turning from a straw color to an almost colorless solution.
The absence of virtually all staining is usually attributable to one of two causes. One reason for total absence of staining is the use of unfixed vs fixed tissue. The second most common cause is the use of tissue that has been lipid extracted, such as tissue processed for paraffin embedding. Thus, Black-Gold II staining is only seen when using formaldehyde fixed, non-embedded tissue sections.
Although staining at room temperature can be both convenient and can also result in useful staining of the myelin, there are two reasons why staining at elevated temperatures (60-65 0C) are generally recommended. One reason is because it is faster (15-20 min vs 1 hr.) and the other reason is that it generally results in a stain of noticeably better the contrast.
The most common myelin pathologies seen involve either internodal edema, or fragmentation. Total demyelination will simply appear as a pale region of the tissue section.
After 15 min in the staining solution, the slide should be removed and examined under the microscope every few minutes. When fully impregnated, both the fiber bundles and the fine individual fibers (like the parallel fibers in layer I of the cerebral cortex) will be stained. If these fine myelinated fibers are not seen, the slide should be replaced in the warm staining solution until such fibers are observed.
If left in the staining solution for too long, the sections can acquire a lavender background staining of the neuropil.
No structures within the normal brain will be labeled, although if not removed, there may be some labeling of red blood cells and collagen fibrils within the meninges.
Although the labeling of degenerating neurons is by far the most conspicuous, in terms of size, number and brightness, sometimes hypertrophied astrocytes immediately surrounding the region of neuronal degeneration will label. Also, in both transgenic mouse models of Alzheimer’s disease as well as human autopsy tissue, amyloid plaques will be stained by these tracers.
Both tracers will label degenerating neurons in high resolution and contrast. Although both tracers have their proponents, in our lab, Fluoro-Jade C gives the optimal stain. But direct comparisons of the two tracers is complicated by the fact that the FJ-B is typically used at a higher concentration (.0004% - .0003%), while the FJ-C is used at a lower concentration (.0001% - .00015%).
This is typically caused by one of two general problems, either the absence of actual degenerating neurons, or some problem with the histochemical staining methodology. Often the problem is that the insult administered did not result in any degenerating neurons. It is at least worth keeping in mind that the Fluoro-Jade dyes only stain dead and degenerating neurons and not “sick” or distressed neurons. It is also worth keeping in mind that the neuronal degeneration usually occurs over a period of a few days to a few weeks. Therefore, studies with longer survival intervals may miss the window in which degenerating neurons are present. The best way to unequivocally resolve if the lack of neuronal labeling is seen because of an absence of degenerating neurons or because of a flaw somewhere in the histochemical methodology is to run a positive control. Such a control can be generated by dosing a rat with kainic acid (10 mg/kg/i.p.) and then sacrificing it 1-2 days later. Animals that exhibit seizure activity will possess degenerating neurons within the cortex and hippocampus. Seizure activity lasting more than 1-2 hours can be stopped by the administration of antiseizure medications such as Nembutal. Should labeling still not be seen, one should make sure that the staining solution is made up in 0.1% acetic acid vehicle and that the mounting media is solvent rather than water-based.
Yes, they will detect degenerating neurons, regardless of whether death was via an apoptotic or necrotic mechanism. This includes diverse classes of neurotoxins (e.g. glutamate agonists, acetylcholine agonists, glutamate antagonists, inhibitors of metabolic respiration, dopamine and serotonin agonists), physical trauma and developmental apoptosis.
There are two main approaches used to reduce high background staining. One is to reduce the concentration of the stain to half of that previously used. This may be more effective for the Fluoro-Jade B, since it is usually used at a higher concentration than the Fluoro-Jade C. Alternatively, background staining can be reduced by increasing the time (e.g. 5 more min) in the potassium permanganate solution. Also, longer post staining rinses can help reduce background staining.
One option would be to increase the concentration. This would probably be most applicable to the Fluoro-Jade C, whose concentration could be increased to .00015-.0002%. Another way to increase the brightness would be to reduce the incubation time (e.g. 5 min less) in the potassium permanganate solution.
Yes, both tracers are capable of detecting degenerating neuronal cell bodies, proximal dendrites, axons and terminals.
No, although this first step can help reduce uneven background staining, good staining can still be achieved without it. Therefore, it can be omitted if the solution is too harsh for subsequent multiple labeling procedures.
Amylo-Glo binds to misfolded proteins in both parenchymal and vascular amyloid plaques as well as intraneuronal NFTs. A similar staining pattern is observed with HQ-O, which probably label zinc bound to the A-beta aggregates. A somewhat different pattern is seen with Euro-Glo, which ehibits a more globular appearance and likely stains complex lipids contained within the plaque. Although primarily used to localize neuronal degeneration, Fluoro-Jade B and Fluoro-Jade C will also both label amyloid plaques, when present. Both vesicular and fibular type labeling can be observed which may be attributed to degenerating neurites incorporated into the plaques.
Both the stock solution and working solution of this dye is basic, resulting in yellow appearing plaques and tangles. However, transferring the slides to a neutral solution or to solvents, including those associated with paraffin processing, will shift the emission more to the blue range. Digital color cameras may also read the light yellow color as light blue.
Try decreasing the final dye concentration by reducing the amount of dye stock solution in the final staining solution (e.g. 7.5 ml stock solution: 92.5 ml water). Longer rinse times may also help reduce background staining.
Try increasing the final dye concentration by increasing the amount of dye stock solution in the final staining solution (e.g. 12.5 ml stock solution: 87.5 ml water). Shorter rinse times may also help enhance the brightness and resolution of the stain.
No, it will also stain myelinated fibers.
No, at room temperature, staining will require around 5-7 days of incubation in the staining solution. The relatively long staining time could suggest that the binding involves multiple hydrogen bonds, as is the case with immunohistochemistry.
Although the specific molecule that the tracer binds to remains to be resolved, it is possible to make some inferences about its chemical nature based on the staining conditions required. Since treatment with non-polar solvents, like ethanol and xylene, will eliminate all staining, It seems probable that Euro-Glo labels some type of complex lipid such as gangliosides or lipoproteins.
No, Euro-Glo is a chelate of the rare earth metal, Europium. This metal confers an unprecedentedly large Stokes Shift value, meaning that short wave UV excitation results in a relatively long wavelength red colored emission.
No, because non-polar solvents will extract the lipid components stained by the tracer, an aqueous mounting media, such as Ever-Glo (Histo-Chem), could be used. However the greatest resistance to fading was seen following coverslipping with mineral oil.
The staining pattern closely correlates with that seen following direct labeling with probes like Amylo-Glo as well as immunohistochemical staining with antibodies directed against A-beta.
Probably not since in vitro mechanistic studies have shown that HQ-O will not bind directly to A-beta aggregates unless zinc is also present. This suggests that HQ-O most likely binds to zinc already bound to the amyloid plaques, resulting in the formation of a highly fluorescent precipitate.
In well perfused brain tissue, HQ-O only labels amyloid plaques. However, in tissues in which the blood has not been completely flushed, labeled zinc containing leucocytes can be localized within the lumen of blood vessels.