Einfache Beseitigung von Hintergrundfluoreszenz in Formalin fixiert menschliche Gehirngewebe für die Immunfluoreszenz-Mikroskopie

A common problem in immunofluorescence microscopy is the presence of endogenous background fluorescence that can interfere with your signal. In many cases, this autofluorescence can make your results uninterpretable. In this protocol we will demonstrate how to remove autofluorescence from human brain tissue samples using photobleaching with a commercial LED desk lamp.

The main advantages of this methods are the significant cost-savings over current techniques without sacrificing effectiveness and that no exogenous compounds are introduced into the sample. Before you begin, prepare stock solutions of one times tris-buffered saline, TBS, and 10%sodium azide. For every three standard size microscope slides that you would like to process, you will need a single 100 millimeter by 100 millimeter square Petri dish as a slide chamber.

For each slide chamber, prepare 50 milliliters of 05%azide in TBS by adding 25 milliliters of 10%sodium azide to 50 milliliters of one times TBS. Create a scaffold to elevate your slide chamber such that a lamp head can fit underneath. For a 100 millimeter square Petri dish, a plastic food container of similar size can be cut to shape.

Anything that can securely elevate the slide chamber without impeding the light from reaching the samples can be used. Acquire a white phosphor LED desk lamp and remove any diffusers or opaque plastic that cover the light source. Orient the LED array upwards.

For this purpose a lamp with a flexible neck is recommended. Construct a reflective dome cover for the apparatus by lighting the inside of a box large enough to cover the slide chambers and scaffold with aluminum foil. For a single chamber, a one millimeter pipette tip box can be used.

For this video, formalin-fixed human FTLD-T orbital frontal gyri brain tissue was subjected to the photobleaching pre-treatment. Tissue preparation and amino staining procedures may vary based on the source, fixation, and embedding of your tissue sections. In a four degree Celsius cold room, cold cabinet, or refrigerator, place the lamp under the scaffold and the sample chamber on top of the scaffold.

Fill the sample chamber with 50 milliliters of azide TBS. Submerge 10 micrometer thick tissue sections mounted on standard glass microscope slides into the sample chamber using clean forceps. Cover the apparatus with the reflective dome and turn on the LED lamp.

Allow the samples to photobleach for 48 hours. This incubation time may vary, depending on the lamp intensity and the amount of autofluorescent material in the tissue. After photobleaching pre-treatment, the slides can be immunostained with a standard protocol for your specific tissue and proteins of interest.

For demonstration purposes, FTLD-T brain tissue was stained for phosphorylated tau using secondary antibodies conjugated to Alexa 488 and Texas Red. DAPI was used as a nuclear counterstain. Prepare 500 milliliters of antigen retrieval buffer, 500 milliliters of 0.025%Triton X-100 in TBS, 10 milliliters of 1%bovine serum albumin, BSA, in TBS, and two milliliters of blocking solution, consisting of goat serum added to BSA TBS.

Perform antigen retrieval by submerging the photobleached slides into antigen retrieval buffer in a slide collector. Heat the collector at 90 degrees Celsius for 30 minutes. Once the collector has been removed from the heat treatment, it is important to allow the chamber to cool at room temperature for 30 minutes.

Removing the slides immediately will cause sections to dry out. Prepare your primary antibody mixture according to manufacturer recommendations. In this case, anti-phospho PHF tau p-serine 202 plus threonine 205 AT8 antibody was diluted one in 100 with BSA TBS and left on ice.

150 microliters of antibody mixture is required per section. After the sections have cooled for 30 minutes, transfer the slides from the antigen retrieval collector to a staining jar filled with TBS Triton. Wash the slides in this solution for five minutes with gentle shaking.

Empty and replace the TBS Triton to repeat this wash step a second time. After washing, remove the slides from the staining jar and wick away any excess buffer. Using a hydrophobic pen, draw an outline around the tissue.

This will prevent your solutions from running off of the slide. Take care to not let the slides dry out. Block the tissue by pipetting 200 microliters of blocking solution onto the slides and place them inside of a humidified chamber.

This chamber can be something as simple as a slide rack inside of pipette tip box containing a wet paper towel. Ensure that the entire tissue sample is covered with blocking solution. Incubate at room temperature for two hours.

Remove the blocked solution by aspirator or pipette. Pipette 100 to 150 microliters of primary antibody mixture onto the slides. Ensure that the entire sample is covered with antibody solution and that the sections are kept on a level surface to avoid uneven pooling of antibody.

Incubate with primary antibody at four degrees Celsius overnight in a humidified chamber. The next day, prepare the secondary antibody mixture. In this case, goat anti-mouse Alexa 488 and goat anti-mouse Texas Red were diluted one in 100 using BSA TBS.

Retrieve the tissue sections and remove the primary antibody solution by aspirator or pipette. Wash the slides two times in a similar manner as previously using a staining jar filled with TBS Triton for five minutes each wash. After washing, wick away any excess buffer.

Pipette 100 to 150 microliters of secondary antibody solution, ensuring once again that the tissue sample is completely covered. Incubate for two hours at room temperature in the humidified chamber in the dark. Prepare 0.1 microgram per milliliter DAPI counterstain in TBS according to manufacturer recommendations or serial dilution.

Be sure to mix the DAPI stock solution thoroughly due to possible insolubility. Retrieve the slides and remove the secondary antibody mixture. Wash the samples two times, but instead of using TBS Triton, use plain TBS.

Aspirate or pipette the remaining wash solution off and pipette 100 to 150 microliters of DAPI stain onto your tissue sections. Incubate in the dark at room temperature for 10 minutes. Retrieve the samples and wash three times with plain TBS instead of two.

After washing, carefully remove the remaining wash solution. Apply mounting medium. Complete the slide mounting by cover the samples with glass cover slips.

Avoid bubbles by placing one edge of the cover slip down and very slowly lowering the opposite edge using forceps. Allow mounting media to dry before storing at four degrees Celsius in the dark. The slides are now ready to be visualized.

The settings for the fluorescence microscopy will vary depending on the fluorophores used for staining and the microscope. Turn on the fluorescence lamp, microscope, and computer, and allow the lamp to warm up for 15 minutes. Place the stained tissue slides in the fluorescence microscope.

Use the bright field to locate the tissue at 10 times magnification. Once a suitable region of tissue is located, add a drop of water on top of the sample's cover slip. Switch to a 20 times water immersion lens.

Select laser excitation and emission wavelengths appropriate for each fluorophore in separate tracks for best signal. Further details of the settings and wavelengths used can be found in the accompanying manuscript. Adjust the laser power and gain settings to optimize signal intensity for each track.

Once appropriate settings and field of view are found, save a composite image containing the fluorescence data from each track. For visualization of fluorescence intensity in each channel, install the RGB Profile Tools macro for ImageJ. Open the lsm confocal image file in ImageJ and convert the composite images from the three stacks to RGB by going to Image, Color, Channel Tool.

Select Composite and check all three channels. Next, go to Image, Type, and RGB Color. Select the RGB Profile Tools icon and draw a line across the section in the image to be profiled.

In this video, the steps to perform a photobleaching pre-treatment of tissue samples to produce background fluorescence have been described. This procedure can be added to any standard amino fluorescence protocol prior to antigen retrieval and amino staining. The apparatus used for photobleaching is inexpensive and easily obtainable, consisting of off-the-shelf components.

White phosphor LEDs emit light across a wide spectrum of wavelengths, which make them highly suitable for broad range photobleaching. Using a readily available LED desk lamp, unstained human FTLD-T brain tissue samples were photobleached and imaged using the wavelengths that would be appropriate if these samples were stained with Texas Red and Alexa 488. Untreated samples displayed large amounts of autofluorescence.

In contrast, photobleaching resulted in significantly reduced intensity of autofluorescence speckles that resembled lipofuscin, as well as general background fluorescence. For further validation of this technique, FTLD-T brain tissue samples were stained for hyperphosphorylated tau, a protein that misfolds and accumulates in this disease as distinct inclusion bodies. Two different secondary antibodies were applied to better distinguish between tau labeling and non-specific fluorescence.

In these composite images, colocalization of Alexa 488 and Texas Red is shown in yellow, which represents correctly stained tau protein. The appearance of red or green regions indicate either non-specific fluorescence or quenching of one of the chromophores. In the untreated control sample, significant background and non-specific signal can be seen, particularly in the Texas Red channel.

Both photobleaching pre-treatment, as well as application of a chemical quencher result in highly reduced background signal. If we split the fluorescence channels and look at the images under higher magnification, more detailed observations can be made. The untreated control possesses background fluorescence across all channels, as well as non-specific features, mainly in the Texas Red channel, that have similar intensity to the tau specific labeling.

If tau inclusion bodies did not have distinct, predictable morphology, these Texas Red results could have been misinterpreted due to the presence of these autofluorescent features. Photobleaching pre-treatment completely abolishes these Texas Red non-specific features, while preserving the majority of the tau specific Texas Red and Alexa 488 signals. Photobleaching, however, appears to have little effect on DAPI background fluorescence.

Chemical quencher reduced the background fluorescence in the Alexa 488 channel just as effectively as photobleaching. However, it also reduced the fluorescence intensities of both DAPI and Texas Red, suggesting the presence of some counterproductive quenching. Quantification of these effects can be seen in the signal profiles.

As you can see, this method produces clean images free of confounding features. This photobleaching pre-treatment can be incorporated into any standard amino fluorescence protocol to remove autofluorescence and generate high-quality, unambiguous results.