This study was supported in whole or in part by the Canadian Consortium of Neurodegeneration and Aging (CCNA), the Canadian Institute of Health Research (CIHR), the ALS Society of Canada (ALS Canada), and the Alzheimer Society of Canada (ASC). The authors would like to thank Sultan Darvesh and Andrew Reid from the Maritime Brain Tissue Bank for providing the FTLD brain tissues, Milan Ganguly from the Spatio-Temporal Targeting and Amplification of Radiation Response (STTARR) program and its affiliated funding agencies for tissue embedding and sectioning services, and the Advanced Optical Microscopy Facility (AOMF) for providing microscopy instruments. Kevin Hadley is thanked for critical review of the manuscript.

Note: The work presented was performed in compliance with recognized international standards, including the International Conference on Harmonization (ICH), the Council for International Organizations of Medical Sciences (CIOMS), and the principles of the Declaration of Helsinki. Use of human tissue was with the approval of University Health Network Research Ethics Board. The human brain samples were collected as a part of the Maritime Brain Tissue Bank. At the time of collection, informed consent was obtained from all patients.
1. Construction of photobleaching apparatus and solutions
<ol>
	<li>Prepare stock solutions.
	<ol>
		<li>Prepare 1 L of 1x stock Tris-buffered saline (1x TBS) solution (150 mM NaCl, 50 mM Tris-Cl, pH 7.4) by dissolving 8.77 g of NaCl and 6.06 g of Tris base in 800 mL of ddH2O and adjust the pH to 7.4 using HCl. Bring up the volume to 1 L and autoclave.</li>
		<li>Prepare 10% (200x stock) sodium azide by dissolving 1 g of sodium azide in 10 mL of ddH2O (10%, 200x stock).</li>
	</ol>
	</li>
	<li>For 1-3 standard size slides, use a single 100 mm x 100 mm transparent, square petri dish as a slide chamber. For a single slide chamber, add 0.25 mL of 200x sodium azide stock to 50 mL of 1x TBS to make a 0.05% azide-TBS solution. Stack 2-3 slide chambers vertically to process additional slides. Prepare an additional 50 mL of azide-TBS solution for each chamber.</li>
	<li>Create a scaffold to elevate the slide chamber(s) such that a lamp head can fit underneath.
	<ol>
		<li>For a 100 mm x 100 mm slide chamber, cut openings in the bottom and sides of a 100 mm x 100 mm x 30 mm plastic food container and invert the container. Ensure the side openings are large enough to fit an LED light source and ensure the bottom opening is large enough such that light from the light source reaches the sample chamber without impediment.</li>
		<li>Apply electrical tape to the scaffold to increase the grip between the scaffold and the sample chamber/benchtop. Use any alternative materials to construct the scaffold so long as it securely elevates the slide chamber without impeding the light from reaching the sample.</li>
	</ol>
	</li>
	<li>Remove any diffusers or opaque plastic from the desk lamp that may impede the LED light from directly reaching the sample (if possible) and orient the LED array upwards. Place the scaffold and slide chamber(s) above the LED array. Use a lamp with a flexible neck for easy manipulation.</li>
	<li>Construct a reflective dome cover for the apparatus by lining the inside of a box large enough to cover the slide chamber and scaffold with aluminum foil. Use a 1 mL pipette tip box for a single chamber or a 150 mm x 150 mm x 150 mm cardboard box for multiple, vertically stacked chambers.</li>
</ol>
2. Photobleaching pre-treatment of tissue sections
NOTE: Tissue section preparation may vary depending on the source of tissue and fixation and embedding methods used. Here, brain tissue (orbitofrontal gyri) from a case of FTLD-T was fixed for ~2 days in formalin, run through a sucrose gradient, embedded in OCT, and cut to 10 &#181;m thick sections using a cryostat.
<ol>
	<li>In a 4 &#186;C cold room, cold cabinet, or refrigerator, orient the lamp under the scaffold and place the sample chamber on the scaffold. Pour 50 mL of azide-TBS solution into the sample chamber.</li>
	<li>Submerge tissue sections mounted on standard glass microscope slides into the slide chamber containing azide-TBS using clean forceps. For multiple slides, ensure that the slides are placed in the chamber on a single layer.</li>
	<li>Cover the apparatus with the reflective dome, turn on the LED lamp, and incubate for 48 h at 4 &#186;C.</li>
</ol>
3. Immunofluorescence
<ol>
	<li>To stain the tissue for phosphorylated tau using DAPI counterstain and Alexa 488- and Texas Red-conjugated secondary antibodies, first prepare solutions for antigen retrieval, permeabilization, blocking, and primary antibody binding.
	<ol>
		<li>Prepare 500 mL of antigen retrieval buffer (10 mM citric acid, 2 mM ethylenediaminetetraacetic acid, 0.05% Tween 20; pH 6.2) by dissolving 0.92 g of citric acid and 0.37 g of ethylenediaminetetraacetic acid (EDTA) in 500 mL of ddH2O. Adjust the pH to 6.2 with NaOH and add 0.25 mL of Tween 20.</li>
		<li>Prepare 500 mL of 0.025% Triton X-100 in TBS solution (TBS-Triton) by adding 0.125 mL of Triton X-100 to 500 mL 1x TBS.</li>
		<li>Prepare a 1% bovine serum albumin (BSA) solution in TBS (BSA-TBS buffer) by dissolving 0.1 g BSA in 10 mL TBS.</li>
		<li>Prepare a blocking solution by adding 0.2 mL of normal goat serum to 1.8 mL of 1% BSA/TBS.</li>
		<li>For each slide, prepare 150 &#181;L of primary antibody solution (1:100 dilution) by pipetting 1.5 &#181;L of anti-phospho-PHF-tau pSer202 + Thr205 (AT8) antibody into 148.5 &#181;L of 1% BSA-TBS buffer and leave on ice.</li>
	</ol>
	</li>
	<li>To perform antigen retrieval, submerge the photobleached slides vertically in a slide collector containing 25 mL of antigen retrieval buffer. Secure the collector with tape and/or strings such that the collector does not fall into the water bath. Heat the collector in a water bath at 90 &#186;C for 30 min and allow the collector to cool to room temperature for 30 min before removing the slides. Do not remove the slides immediately as it will cause the sections to dry out.</li>
	<li>Transfer the slides from the antigen retrieval collector into a staining jar filled with 30 mL of TBS-Triton and wash the sections for 5 min on an orbital shaker with gentle shaking. Repeat the wash once with fresh TBS-Triton. Wick away excess buffer with a lint-free tissue and outline the tissue with a hydrophobic pen. Take care not to let the slides dry out.
	<ol>
		<li>For each slide, block the tissue by pipetting 200 &#181;L of blocking solution onto the tissue and place the slide in a humidified chamber. Construct the chamber by placing a slide rack inside a pipette tip box containing a wet paper towel. Incubate at room temperature for 2 h on a level surface. Ensure that the blocking solution fully covers the tissue.</li>
	</ol>
	</li>
	<li>Remove the blocking solution by aspiration and pipette 100-150 &#181;L of primary antibody solution onto the tissue. Ensure sufficient volume of antibody is present and that the section is on a level surface to avoid pooling of antibody solution to one side. Incubate at 4 &#186;C overnight in a humidified chamber.</li>
	<li>Prepare the secondary antibody mixture and the DAPI nuclear counterstain.
	<ol>
		<li>For each slide, prepare 150 &#181;L of secondary antibody mixture (1:100 dilutions) by adding 1.5 &#181;L of goat anti-mouse Alexa 488 and 1.5 &#181;L of goat anti-mouse Texas Red to 147 &#181;L of BSA-TBS and leave on ice.</li>
		<li>Prepare 0.1 &#181;g/mL DAPI counterstain by serial dilution. Mix the stock solution thoroughly and dilute 1 &#181;L of stock 5 mg/mL DAPI in 999 of TBS to make 1 mL of 5 &#181;g/mL solution. For each slide, dilute 3 &#181;L of 5 &#181;g/mL solution with 147 &#181;L of TBS to a final concentration of 0.1 &#181;g/mL. 
		CAUTION: DAPI is a known mutagen and should be handled with care.</li>
	</ol>
	</li>
	<li>Remove the primary antibody by aspiration. Submerge the slides in a glass staining jar containing 30 mL of TBS-Triton and wash for 5 min with gentle mixing on an orbital shaker. Repeat the wash step with fresh TBS-Triton. Wick away excess TBS-Triton and pipette 100 to 150 &#181;L of secondary antibody mixture to each slide.
	<ol>
		<li>Ensure the tissue is fully covered by the antibody mixture. Incubate for 2 h at room temperature in the humidified chamber in the dark.</li>
	</ol>
	</li>
	<li>Remove the secondary antibody mixture by aspiration and transfer the slide into a glass staining jar containing 30 mL of TBS (no Triton). Wash for 5 min with gentle mixing on an orbital shaker. Repeat the wash step with fresh TBS. Apply 100 to 150 &#181;L of 0.1 &#181;g/mL DAPI counterstain to each slide and incubate for 10 min at room temperature in the dark.</li>
	<li>Transfer the slides to a glass staining jar containing TBS and wash 3 times with gentle mixing for 5 min each, using fresh TBS for each wash. Wick away excess buffer.</li>
	<li>Apply 3 drops of aqueous mounting medium to the tissue. Using forceps, gently lower a clean glass coverslip onto the tissue, starting with one edge and slowly lowering the other edge to avoid trapping of air bubbles. Take care not to dislodge the coverslip if imaging immediately. Otherwise, allow the mounting medium to dry before storing at 4 &#186;C in the dark.</li>
</ol>
4. Fluorescence microscopy
<ol>
	<li>Turn on the fluorescence lamp, the microscope and the computer and allow the lamp to warm up for 15 min. Place the stained tissue slides in the fluorescence microscope stage. Use the bright field image to locate the tissue at 10x magnification.</li>
	<li>Apply a drop of ddH2O to the coverslip surface and use a 20x water immersion objective lens (NA=1.0). Select the 4-line average plane scan setting. Set the pinhole size to 1 Airy unit that gives an optical slice of ~3 microns. Select the laser excitation and emission wavelengths for each fluorophore in separate tracks for best signal. 
	NOTE: Alexa 488: &#955;ex = 488 nm (argon laser) &#955;em = 493-570 nm; Texas Red: &#955;ex = 561 nm (DPSS 561 nm laser), &#955;em = 601-635 nm; DAPI: &#955;ex = 405 nm (Diode 405 laser), &#955;em = 410-507 nm.
	<ol>
		<li>Adjust the laser power and gain settings to optimize the signal intensity for each track. Collect the composite image and save. Use the same laser settings to compare fluorescence intensities in a different slide.</li>
	</ol>
	</li>
	<li>For visualization of fluorescence intensity in each channel, install the RGB profile tools macro for ImageJ10. Save the macro from the webpage as a text file (https://imagej.nih.gov/ij/macros/tools/RGBProfilesTool.txt). From the ImageJ menu, select Plugins -&#62; Macros -&#62; Install; select the text file to install the RGB profiles tool.
	<ol>
		<li>Open the confocal image file in ImageJ and convert the composite images from the 3 stacks to RGB by performing the following: Image -&#62; Color -&#62; Channel Tool. Select &#34;Composite&#34; from the dropdown menu and check all three channels. Then, select Image -&#62; Type -&#62; RGB color.</li>
		<li>&#160;Select the RGB profile tools icon and draw a line across the section in the image to be profiled. Save the intensity data as a spreadsheet for plotting.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Banerjee, B., Miedema, B. E., Chandrasekhar, H. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Role+of+basement+membrane+collagen+and+elastin+in+the+autofluorescence+spectra+of+the+colon.">Role of basement membrane collagen and elastin in the autofluorescence spectra of the colon.</a> J Investig Med. 47 (6), 326-332 (1999).</li><li>Terman, A., Brunk, U. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lipofuscin+.">Lipofuscin .</a> Int J Biochem Cell Biol. 36 (8), 1400-1404 (2004).</li><li>Del Castillo, P., Llorente, A. R., Stockert, J. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Influence+of+fixation,+exciting+light+and+section+thickness+on+the+primary+fluorescence+of+samples+for+microfluorometric+analysis.">Influence of fixation, exciting light and section thickness on the primary fluorescence of samples for microfluorometric analysis.</a> Basic Appl Histochem. 33 (3), 251-257 (1989).</li><li>Ottis, P., Koppe, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Human+and+rat+brain+lipofuscin+proteome.">Human and rat brain lipofuscin proteome.</a> Proteomics. 12 (15-16), 2445-2454 (2012).</li><li>Sun, Y., Chakrabartty, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cost-effective+elimination+of+lipofuscin+fluorescence+from+formalin-fixed+brain+tissue+by+white+phosphor+light+emitting+diode+array.">Cost-effective elimination of lipofuscin fluorescence from formalin-fixed brain tissue by white phosphor light emitting diode array.</a> Biochem Cell Biol. 94 (6), 545-550 (2016).</li><li>Davis, A. S., Richter, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterizing+and+Diminishing+Autofluorescence+in+Formalin-fixed+Paraffin-embedded+Human+Respiratory+Tissue.">Characterizing and Diminishing Autofluorescence in Formalin-fixed Paraffin-embedded Human Respiratory Tissue.</a> J Histochem Cytochem. 62 (6), 405-423 (2014).</li><li>Zimmermann, T., Rietdorf, J., Pepperkok, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spectral+imaging+and+its+applications+in+live+cell+microscopy.">Spectral imaging and its applications in live cell microscopy.</a> FEBS Lett. 546 (1), 87-92 (2003).</li><li>Duong, H., Han, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+multispectral+LED+array+for+the+reduction+of+background+autofluorescence+in+brain+tissue.">A multispectral LED array for the reduction of background autofluorescence in brain tissue.</a> J Neurosci Methods. 220 (1), 46-54 (2013).</li><li>Albeanu, D. F., Soucy, E., Sato, T. F., Meister, M., Murthy, V. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=LED+arrays+as+cost+effective+and+efficient+light+sources+for+widefield+microscopy.">LED arrays as cost effective and efficient light sources for widefield microscopy.</a> PLoS One. 3 (5), 1-7 (2008).</li><li>Schneider, C. A., Rasband, W. S., Eliceiri, K. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=NIH+Image+to+ImageJ:+25+years+of+image+analysis.">NIH Image to ImageJ: 25 years of image analysis.</a> Nat Methods. 9 (7), 671-675 (2012).</li><li>Robertson, J. B., Zhang, Y., Johnson, C. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Light-emitting+diode+flashlights+as+effective+and+inexpensive+light+sources+for+fluorescence+microscopy.">Light-emitting diode flashlights as effective and inexpensive light sources for fluorescence microscopy.</a> J Microsc. 236 (1), 1-4 (2009).</li><li>Mackenzie, I. R. A., Neumann, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Molecular+neuropathology+of+frontotemporal+dementia:+insights+into+disease+mechanisms+from+postmortem+studies.">Molecular neuropathology of frontotemporal dementia: insights into disease mechanisms from postmortem studies.</a> J Neurochem. 138 (S1), 54-70 (2016).</li><li>Kouri, N., Whitwell, J. L., Josephs, K. A., Rademakers, R., Dickson, D. 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The authors have nothing to disclose.

The photobleaching pre-treatment of tissues described in this manuscript allows for effective elimination of autofluorescence using off-the-shelf components. The protocol describes immunofluorescence imaging of phosphorylated tau aggregates in formalin-fixed human brain tissue using secondary antibodies conjugated to Alexa 488 and Texas Red, with DAPI as a nuclear counterstain. To apply the method to other tissues, we recommend performing a 48 h photobleaching pre-treatment to the sample as a starting point. After photob...

Fluorescence microscopy using antibodies targeting specific proteins is routinely used to visualize proteins of interest in cell culture and tissues. A major complication to the acquisition of clear and definitive images in immunofluorescence is autofluorescence, which can be caused endogenously in mammalian tissue by the age pigment lipofuscin and by proteins such as elastin and collagen1,2. Other sources of autofluorescence can be introduced through sample preparation steps such as aldehyde fixation3. Lipofuscin granules, composed primarily of oxidatively modified protein and lipid degradation residues, accumulate in long-living cells with increased age2. This causes difficulties in imaging postmitotic tissues such as the brain and cardiac or skeletal muscles, as the fluorescence emission spectrum of lipofuscin is broad and variable, often coinciding with the emission wavelength of common fluorophores used for labeling4. These factors make imaging of human brain tissue from cases of late-onset neurodegenerative diseases such as frontotemporal lobar degeneration (FTLD) especially challenging.
To reduce autofluorescence, we have devised a technique in which we irradiate the slide-mounted tissue sections with a white light emitting diode (LED) array using a household desk lamp5. This simple technique provides an alternative to techniques that use chemical quenchers such as CuSO4 in ammonium acetate, or commercially available quenching dyes such as Sudan Black B and Eriochrome Black T6. It also has significant cost-saving over multispectral LED lamp photobleaching techniques and avoids complications and artefacts generated from digital autofluorescence removal methods such as spectral un-mixing7,8. White phosphor LEDs have a broad emission spectrum, high luminosity and low manufacturing cost, making them ideal as an off-the-shelf component for photobleaching a variety of chromophores5,9.
In this protocol, we demonstrate the construction of a photobleaching apparatus using accessible components and apply photobleaching to a case of FTLD tissue containing tau-positive inclusions (FTLD-T) using an antibody specific for phosphorylated tau. We demonstrate the effect of photobleaching on imaging fluorescently-labeled antibodies employing two commonly-used chromophores: Alexa 488 and Texas Red. The effect of photobleaching versus untreated sections or those treated with a commercial chemical quencher are quantified and compared. This photobleaching pre-treatment can be incorporated into any standard immunofluorescence staining protocol to remove autofluorescence in a biological sample.

<table border="0" cellpadding="0" cellspacing="0" width="999">
	<tbody>
		<tr height="40">
			<td height="40" style="height:40px;width:328px;">Trizma Base</td>
			<td style="width:167px;">Sigma-Aldrich</td>
			<td style="width:104px;">T6066</td>
			<td style="width:400px;"></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Sodium Choloride</td>
			<td>Sigma-Aldrich</td>
			<td>S7653</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Hydrochloric Acid</td>
			<td>Caledon Laboratory Chemicals</td>
			<td>1506656</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Sodium Azide</td>
			<td>BioShop Canada</td>
			<td>SAZ001</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">100 mm x 100 mm x 20 mm Pitri dish</td>
			<td>Sarstedt</td>
			<td>82.9923.422</td>
			<td>All components of photobleacher can be substituted based on availability</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">6 W LED Dimmable Desk Lamp</td>
			<td>DBPower</td>
			<td>DS501</td>
			<td>All components of photobleacher can be substituted based on availability</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Citric Acid</td>
			<td>Sigma-Aldrich</td>
			<td>C-2404</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Ethylenediaminetetraacetic acid&#160;(EDTA)</td>
			<td>BioShop Canada</td>
			<td>EDT001</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Tween 20</td>
			<td>Sigma-Aldrich</td>
			<td>P-7949</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Sodium Hydroxide</td>
			<td>BioShop Canada</td>
			<td>SHY700.1</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Water bath</td>
			<td>Haake Fisons</td>
			<td>K15</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Slide collector</td>
			<td>FisherScientific</td>
			<td>12-587-17B</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Staining Jar</td>
			<td>FisherScientific</td>
			<td>E94</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Orbital Shaker</td>
			<td>Bellco Glass</td>
			<td>&#160;7744-08115</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Triton X-100</td>
			<td>Sigma-Aldrich</td>
			<td>T7878</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Bovine Serum Albumin</td>
			<td>FisherScientific</td>
			<td>BP1600-1</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Normal Goat Serum</td>
			<td>Aurion</td>
			<td>905.002</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Hydrophobic pen</td>
			<td>Sigma-Aldrich</td>
			<td>Z672548-1EA</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Phospho-Tau (Ser202, Thr205) Monoclonal Antibody (AT8)</td>
			<td>ThermoFisher</td>
			<td>MN1020</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Goat anti-Mouse Secondary Antibody, Texas Red-X</td>
			<td>ThermoFisher</td>
			<td>T862</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Goat anti-Mouse Secondary Antibody, Alexa Fluor 488</td>
			<td>ThermoFisher</td>
			<td>A-11029</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">DAPI</td>
			<td>Sigma-Aldrich</td>
			<td>D9542</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Mounting medium</td>
			<td>ThermoScientific</td>
			<td>28-600-42</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Glass soverslip</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Confocal Microscope</td>
			<td>Zeiss</td>
			<td>LSM710</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Imaging software ZEN 2012 Black Edition 11.0</td>
			<td>Zeiss</td>
			<td>LSM710</td>
			<td>Software accompanies the Confocal Microscope</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">ImageJ</td>
			<td>NIH</td>
			<td></td>
			<td>https://imagej.nih.gov/ij/download.html</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">RGB Profile Tools macro</td>
			<td>NIH</td>
			<td></td>
			<td>https://imagej.nih.gov/ij/macros/tools/RGBProfilesTool.txt</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Commercial chemical quencher</td>
			<td>Biotum</td>
			<td>23007</td>
			<td></td>
		</tr>
	</tbody>
</table>

simple elimination of background fluorescence in formalin-fixed human brain tissue for immunofluorescence microscopy

Immunofluorescence is a common method used to visualize subcellular compartments and to determine the localization of specific proteins within a tissue sample. A great hindrance to the acquisition of high quality immunofluorescence images is endogenous autofluorescence of the tissue caused by aging pigments such as lipofuscin or by common sample preparation processes such as aldehyde fixation. This protocol describes how background fluorescence can be greatly reduced through photobleaching using white phosphor light emitting diode (LED) arrays prior to treatment with fluorescent probes. The broad-spectrum emission of white phosphor LEDs allow for bleaching of fluorophores across a range of emission peaks. The photobleaching apparatus can be constructed from off-the-shelf components at very low cost and offers an accessible alternative to commercially available chemical quenchers. A photobleaching pre-treatment of the tissue followed by conventional immunofluorescence staining generates images free of background autofluorescence. Compared to established chemical quenchers which reduced probe as well as background signals, photobleaching treatment had no effect on probe fluorescence intensity while it effectively reduced background and lipofuscin fluorescence. Although photobleaching requires more time for pre-treatment, higher intensity LED arrays may be used to reduce photobleaching time. This simple method can potentially be applied to a variety of tissues, particularly postmitotic tissues that accumulate lipofuscin such as the brain and cardiac or skeletal muscles.

The photobleaching pre-treatment step can be added to a standard immunofluorescence protocol immediately prior to antigen retrieval and immunostaining (Figure 1A). Assembly of the photobleaching apparatus can also be performed using various, inexpensive, off-the-shelf components (Figure 1B). The emission spectrum of white phosphor LEDs covers a wide range of wavelengths which makes them suitable for broad-range photobleaching, ag...

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Simple Elimination of Background Fluorescence in Formalin-Fixed Human Brain Tissue for Immunofluorescence Microscopy

Background autofluorescence of biological samples often complicates fluorescence-based imaging techniques, especially in aged human postmitotic tissues. This protocol describes how autofluorescence from these samples can be effectively removed using a commercially available light emitting diode light source to photobleach the sample prior to immunostaining.

Immunofluorescence is a common method used to visualize subcellular compartments and to determine the localization of specific proteins within a tissue ...

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Biochemistry

17.1K Views.  University of Toronto. Background autofluorescence of biological samples often complicates fluorescence-based imaging techniques, especially in aged human postmitotic tissues. This protocol describes how autofluorescence from these samples can be effectively removed using a commercially available light emitting diode light source to photobleach the sample prior to immunostaining. 

Video: Simple Elimination of Background Fluorescence in Formalin-Fixed Human Brain Tissue for Immunofluorescence Microscopy

Optimized Protocol for the Extraction of Proteins from the Human Mitral Valve

Analysis of the cellular proteome can help to elucidate the molecular mechanisms underlying diseases due to the development of technologies that permit the large-scale identification and quantification of the proteins present in complex biological systems.The knowledge gained from a proteomic approach can potentially lead to a better understanding of the pathogenic mechanisms underlying diseases, allowing for the identification of novel diagnostic and prognostic disease markers, and, hopefully, of therapeutic targets. However, the cardiac mitral valve represents a very challenging sample for proteomic analysis because of the low cellularity in proteoglycan and collagen-enriched extracellular matrix. This makes it challenging to extract proteins for a global proteomic analysis. This work describes a protocol that is compatible with subsequent protein analysis, such as quantitative proteomics and immunoblotting. This can allow for the correlation of data concerning protein expression with data on quantitative mRNA expression and non-quantitative immunohistochemical analysis. Indeed, these approaches, when performed together, will lead to a more comprehensive understanding of the molecular mechanisms underlying diseases, from mRNA to post-translational protein modification. Thus, this method can be relevant to researchers interested in the study of cardiac valve physiopathology.

The protein composition of the human mitral valve is still partially unknown, because its analysis is complicated by low cellularity and therefore by low protein biosynthesis. This work provides a protocol to efficiently extract protein for the analysis of the mitral valve proteome.

Analysis of the cellular proteome can help to elucidate the molecular mechanisms underlying diseases due to the development of technologies that permit ...

Optimal Preparation of Formalin Fixed Samples for Peptide Based Matrix Assisted Laser Desorption/Ionization Mass Spectrometry Imaging Workflows

The use of matrix-assisted laser desorption/ionization, mass spectrometry imaging (MALDI MSI) has rapidly expanded, since this technique analyzes a host of biomolecules from drugs and lipids to N-glycans. Although various sample preparation techniques exist, detecting peptides from formaldehyde preserved tissues remains one of the most difficult challenges for this type of mass spectrometric analysis. For this reason, we have created and optimized a robust methodology that preserves the spatial information contained within the sample, while eliciting the greatest number of ionizable peptides. We have also aimed to achieve this in a cost effective and simple way, thereby eliminating potential bias or preparation error, which can occur when using automated instrumentation. The end result is a reproducible and inexpensive protocol.

This protocol describes a reproducible and reliable method for the sublimation-based preparation of formalin fixed tissue destined for imaging mass spectrometry.

The use of matrix-assisted laser desorption/ionization, mass spectrometry imaging (MALDI MSI) has rapidly expanded, since this technique analyzes a host ...

High Sensitivity Measurement of Transcription Factor-DNA Binding Affinities by Competitive Titration Using Fluorescence Microscopy

Accurate quantification of transcription factor (TF)-DNA interactions is essential for understanding the regulation of gene expression. Since existing approaches suffer from significant limitations, we have developed a new method for determining TF-DNA binding affinities with high sensitivity on a large scale. The assay relies on the established fluorescence anisotropy (FA) principle but introduces important technical improvements. First, we measure a full FA competitive titration curve in a single well by incorporating TF and a fluorescently labeled reference DNA in a porous agarose gel matrix. Unlabeled DNA oligomer is loaded on the top as a competitor and, through diffusion, forms a spatio-temporal gradient. The resulting FA gradient is then read out using a customized epifluorescence microscope setup. This improved setup greatly increases the sensitivity of FA signal detection, allowing both weak and strong binding to be reliably quantified, even for molecules of similar molecular weights. In this fashion, we can measure one titration curve per well of a multi-well plate, and through a fitting procedure, we can extract both the absolute dissociation constant (KD) and active protein concentration. By testing all single-point mutation variants of a given consensus binding sequence, we can survey the entire binding specificity landscape of a TF, typically on a single plate. The resulting position weight matrices (PWMs) outperform those derived from other methods in predicting in vivo TF occupancy. Here, we present a detailed guide for implementing HiP-FA on a conventional automated fluorescent microscope and the data analysis pipeline.

Here we present a novel method for determining binding affinities at equilibrium and in solution with high sensitivity on a large scale. This improves the quantitative analysis of transcription factor-DNA binding. The method is based on automated fluorescence anisotropy measurements in a controlled delivery system.

Accurate quantification of transcription factor (TF)-DNA interactions is essential for understanding the regulation of gene expression. Since existing ...

An All-in-one Sample Holder for Macromolecular X-ray Crystallography with Minimal Background Scattering

Macromolecular X-ray crystallography (MX) is the most prominent method to obtain high-resolution three-dimensional knowledge of biological macromolecules. A prerequisite for the method is that highly ordered crystalline specimen need to be grown from the macromolecule to be studied, which then need to be prepared for the diffraction experiment. This preparation procedure typically involves removal of the crystal from the solution, in which it was grown, soaking of the crystal in ligand solution or cryo-protectant solution and then immobilizing the crystal on a mount suitable for the experiment. A serious problem for this procedure is that macromolecular crystals are often mechanically unstable and rather fragile. Consequently, the handling of such fragile crystals can easily become a bottleneck in a structure determination attempt. Any mechanical force applied to such delicate crystals may disturb the regular packing of the molecules and may lead to a loss of diffraction power of the crystals. Here, we present a novel all-in-one sample holder, which has been developed in order to minimize the handling steps of crystals and hence to maximize the success rate of the structure determination experiment. The sample holder supports the setup of crystal drops by replacing the commonly used microscope cover slips. Further, it allows in-place crystal manipulation such as ligand soaking, cryo-protection and complex formation without any opening of the crystallization cavity and without crystal handling. Finally, the sample holder has been designed in order to enable the collection of in situ X-ray diffraction data at both, ambient and cryogenic temperature. By using this sample holder, the chances to damage the crystal on its way from crystallization to diffraction data collection are considerably reduced since direct crystal handling is no longer required.

A novel sample holder for macromolecular X-ray crystallography along with a suitable handling protocol is presented. The system allows crystal growth, crystal soaking and in situ diffraction data collection at both, ambient and cryogenic temperature without the need of any crystal manipulation or mounting.

Macromolecular X-ray crystallography (MX) is the most prominent method to obtain high-resolution three-dimensional knowledge of biological macromolecules. ...

Mitochondria and Endoplasmic Reticulum Imaging by Correlative Light and Volume Electron Microscopy

Cellular organelles, such as mitochondria and endoplasmic reticulum (ER), create a network to perform a variety of functions. These highly curved structures are folded into various shapes to form a dynamic network depending on the cellular conditions. Visualization of this network between mitochondria and ER has been attempted using super-resolution fluorescence imaging and light microscopy; however, the limited resolution is insufficient to observe the membranes between the mitochondria and ER in detail. Transmission electron microscopy provides good membrane contrast and nanometer-scale resolution for the observation of cellular organelles; however, it is exceptionally time-consuming when assessing the three-dimensional (3D) structure of highly curved organelles. Therefore, we observed the morphology of mitochondria and ER via correlative light-electron microscopy (CLEM) and volume electron microscopy techniques using enhanced ascorbate peroxidase 2 and horseradish peroxidase staining. An en bloc staining method, ultrathin serial sectioning (array tomography), and volume electron microscopy were applied to observe the 3D structure. In this protocol, we suggest a combination of CLEM and 3D electron microscopy to perform detailed structural studies of mitochondria and ER.

We present a protocol to study the distribution of mitochondria and endoplasmic reticulum in whole cells after genetic modification using correlative light and volume electron microscopy including ascorbate peroxidase 2 and horseradish peroxidase staining, serial sectioning of cells with and without the target gene in the same section, and serial imaging via electron microscopy.

Cellular organelles, such as mitochondria and endoplasmic reticulum (ER), create a network to perform a variety of functions. These highly curved ...

Interactions with and Membrane Permeabilization of Brain Mitochondria by Amyloid Fibrils

A growing body of evidence indicates that membrane permeabilization, including internal membranes such as mitochondria, is a common feature and primary mechanism of amyloid aggregate-induced toxicity in neurodegenerative diseases. However, most reports describing the mechanisms of membrane disruption are based on phospholipid model systems, and studies directly targeting events occurring at the level of biological membranes are rare. Described here is a model for studying the mechanisms of amyloid toxicity at the membrane level. For mitochondrial isolation, density gradient medium is used to obtain preparations with minimal myelin contamination. After mitochondrial membrane integrity confirmation, the interaction of amyloid fibrils arising from &#945;-synuclein, bovine insulin, and hen egg white lysozyme (HEWL) with rat brain mitochondria, as an in vitro biological model, is investigated. The results demonstrate that treatment of brain mitochondria with fibrillar assemblies can cause different degrees of membrane permeabilization and ROS content enhancement. This indicates structure-dependent interactions between amyloid fibrils and mitochondrial membrane. It is suggested that biophysical properties of amyloid fibrils and their specific binding to mitochondrial membranes may provide explanations for some of these observations.

Provided here is a protocol for investigating the interactions between native form, prefibrillar, and mature amyloid fibrils of different peptides and proteins with mitochondria isolated from different tissues and various areas of the brain.

A growing body of evidence indicates that membrane permeabilization, including internal membranes such as mitochondria, is a common feature and primary ...

Production of Dynein and Kinesin Motor Ensembles on DNA Origami Nanostructures for Single Molecule Observation

Cytoskeletal motors are responsible for a wide variety of functions in eukaryotic cells, including mitosis, cargo transport, cellular motility, and others. Many of these functions require motors to operate in ensembles. Despite a wealth of knowledge about the mechanisms of individual cytoskeletal motors, comparatively less is known about the mechanisms and emergent behaviors of motor ensembles, examples of which include changes to ensemble processivity and velocity with changing motor number, location, and configuration. Structural DNA nanotechnology, and the specific technique of DNA origami, enables the molecular construction of well-defined architectures of motor ensembles. The shape of cargo structures as well as the type, number and placement of motors on the structure can all be controlled. Here, we provide detailed protocols for producing these ensembles and observing them using total internal reflection fluorescence microscopy. Although these techniques have been specifically applied for cytoskeletal motors, the methods are generalizable to other proteins that assemble in complexes to accomplish their tasks. Overall, the DNA origami method for creating well-defined ensembles of motor proteins provides a powerful tool for dissecting the mechanisms that lead to emergent motile behavior.

The goal of this protocol is to form ensembles of molecular motors on DNA origami nanostructures and observe the ensemble motility using total internal reflection fluorescence microscopy.

Cytoskeletal motors are responsible for a wide variety of functions in eukaryotic cells, including mitosis, cargo transport, cellular motility, and ...

Myosin-Specific Adaptations of In vitro Fluorescence Microscopy-Based Motility Assays

Myosin proteins bind and interact with filamentous actin (F-actin) and are found in organisms across the phylogenetic tree. Their structure and enzymatic properties are adapted for the particular function they execute in cells. Myosin 5a processively walks on F-actin to transport melanosomes and vesicles in cells. Conversely, nonmuscle myosin 2b operates as a bipolar filament containing approximately 30 molecules. It moves F-actin of opposite polarity toward the center of the filament, where the myosin molecules work asynchronously to bind actin, impart a power stroke, and dissociate before repeating the cycle. Nonmuscle myosin 2b, along with its other nonmuscle myosin 2 isoforms, has roles that include&#160;cell adhesion, cytokinesis, and tension maintenance. The mechanochemistry of myosins can be studied by performing in vitro motility assays using purified proteins. In the gliding actin filament assay, the myosins are bound to a microscope coverslip surface and translocate fluorescently labeled F-actin, which can be tracked. In the single molecule/ensemble motility assay, however, F-actin is bound to a coverslip and the movement of fluorescently labeled myosin molecules on the F-actin is observed. In this report, the purification of recombinant myosin 5a from Sf9 cells using affinity chromatography is outlined. Following this, we outline two fluorescence microscopy-based assays: the gliding actin filament assay and the inverted motility assay. From these assays, parameters such as actin translocation velocities and single molecule run lengths and velocities can be extracted using the image analysis software. These techniques can also be applied to study the movement of single filaments of the nonmuscle myosin 2 isoforms, discussed herein in the context of nonmuscle myosin 2b. This workflow represents a protocol and a set of quantitative tools that can be used to study the single molecule and ensemble dynamics of nonmuscle myosins.

Presented here is a procedure to express and purify myosin 5a followed by a discussion of its characterization, using both ensemble and single molecule in vitro fluorescence microscopy-based assays, and how these methods can be modified for the characterization of nonmuscle myosin 2b.

Myosin proteins bind and interact with filamentous actin (F-actin) and are found in organisms across the phylogenetic tree. Their structure and enzymatic ...

Continuous Fluorescence-Based Endonuclease-Coupled DNA Methylation Assay to Screen for DNA Methyltransferase Inhibitors

DNA methylation, a form of epigenetic gene regulation, is important for normal cellular function. In cells, proteins called DNA methyltransferases (DNMTs) establish and maintain the DNA methylation pattern. Changes to the normal DNA methylation pattern are linked to cancer development and progression, making DNMTs potential cancer drug targets. Thus, identifying and characterizing novel small molecule inhibitors of these enzymes is of great importance. This paper presents a protocol that can be used to screen for DNA methyltransferase inhibitors. The continuous coupled kinetics assay allows for initial velocities of DNA methylation to be determined in the presence and absence of potential small molecule inhibitors. The assay uses the methyl-sensitive endonuclease Gla I to couple methylation of a hemimethylated DNA substrate to fluorescence generation.
This continuous assay allows for enzyme activity to be monitored in real time. Conducting the assay in small volumes in microtiter plates reduces the cost of reagents. Using this assay, a small example screen was conducted for inhibitors of DNMT1, the most abundant DNMT isozyme in humans. The highly substituted anthraquinone natural product, laccaic acid A, is a potent, DNA-competitive inhibitor of DNMT1. Here, we examine three potential small molecule inhibitors &#8212; anthraquinones or anthraquinone-like molecules with one to three substituents &#8212; at two concentrations to describe the assay protocol. Initial velocities are used to calculate the percent activity observed in the presence of each molecule. One of three compounds examined exhibits concentration-dependent inhibition of DNMT1 activity, indicating that it is a potential inhibitor of DNMT1.

DNA methyltransferases are potential cancer drug targets. Here, a protocol is presented to assess small molecules for DNA methyltransferase inhibition. This assay utilizes an endonuclease to couple DNA methylation to fluorescence generation and allows for enzyme activity to be monitored in real time.&#160;

DNA methylation, a form of epigenetic gene regulation, is important for normal cellular function. In cells, proteins called DNA methyltransferases (DNMTs) ...

Quantification of Subcellular Glycogen Distribution in Skeletal Muscle Fibers using Transmission Electron Microscopy

With the use of transmission electron microscopy, high-resolution images of fixed samples containing individual muscle fibers can be obtained. This enables quantifications of ultrastructural aspects such as volume fractions, surface area to volume ratios, morphometry, and physical contact sites of different subcellular structures. In the 1970s, a protocol for enhanced staining of glycogen in cells was developed and paved the way for a string of studies on the subcellular localization of glycogen and glycogen particle size using transmission electron microscopy. While most analyses interpret glycogen as if it is homogeneously distributed within the muscle fibers, providing only a single value (e.g., an average concentration), transmission electron microscopy has revealed that glycogen is stored as discrete glycogen particles located in distinct subcellular compartments. Here, the step-by-step protocol from tissue collection to the quantitative determination of the volume fraction and particle diameter of glycogen in the distinct subcellular compartments of individual skeletal muscle fibers is described. Considerations on how to 1) collect and stain tissue specimens, 2) perform image analyses and data handling, 3) evaluate the precision of estimates, 4) discriminate between muscle fiber types, and 5) methodological pitfalls and limitations are included.

A modified post-fixation procedure increases the contrast of glycogen particles in tissue. This paper provides a step-by-step protocol describing how to handle the tissue, conduct the imaging, and use stereological methods to obtain unbiased and quantitative data on fiber type-specific subcellular glycogen distribution in skeletal muscle.

With the use of transmission electron microscopy, high-resolution images of fixed samples containing individual muscle fibers can be obtained. This ...