Research reported in this publication was supported by the National Institute of Aging of the National Institutes of Health under award numbers R01AG062840 (to RKS) and R01AG072896 (to RKS). The content is solely the authors&#39; responsibility and does not necessarily represent the official views of the National Institutes of Health. Approximately $200k (100%) of federal funds supported this project. We would also like to thank Dr. Joshua Yang for his assistance with manuscript editing.

All animal experiments were conducted in compliance with the University Laboratory Animal Resources under protocols approved by the University of California, Irvine, Institutional Animal Care and Use Committee. The experiments were performed with male B6C3-Tg(APPswe, PSEN1dE9)85Dbo/Mmjax (APP/PS1) mice (13-month-old, n = 35). The mice were obtained from commercial sources (see Table of Materials).
1. Brain tissue preparation
<ol>
	<li>Anesthetize the mice using a lethal dose of a phenytoin/pentobarbital-based anesthetic (150 mg/kg) injected intraperitoneally (see Table of Materials) following approved animal protocols. Perform cardiac perfusion with ice-cold phosphate-buffered saline (1x PBS) for 5 min at a speed of 5 mL/min to clear the brain vasculature14.</li>
	<li>Harvest brain tissue following the previously published report15, separate into the left and right brain hemisphere and place the right hemi-brain of each mouse into a 15 mL conical tube containing 5 mL of freshly prepared 4% paraformaldehyde&#160;(PFA)&#160;solution in 1x PBS for 72 h at 4 &#176;C. 
	NOTE: The brains may be immersion-fixed from 24-72 h depending on the antigen being studied. The left hemi-brain, without the cerebellum, can be snap-frozen in liquid nitrogen and then stored at -80 &#176;C, followed by processing for biochemical assays such as ELISA and biochemical A&#946; detection16. 
	CAUTION: PFA is a probable carcinogen, and skin contact with PFA may lead to allergic skin symptoms. Use nitrile or butyl gloves, masks, and eye protection for handling it, and prepare under a chemical fume hood.</li>
	<li>After incubation in 4% PFA, incubate the hemi-brains sequentially in 5 mL of 10%, 20%, and 30% sucrose solutions prepared in 1x PBS for 24 h, each at 4 &#176;C until the brain tissue sinks to the bottom of the conical tube.</li>
	<li>After incubation in the 30% sucrose solution, remove the hemi-brain, gently dab the brain on a filter paper to remove the excess sucrose solution, and freeze the fixed hemi-brain in powdered dry ice for 30 min. Store the frozen hemi-brain in well-labeled aluminum foils at -80 &#176;C until cryosectioning. 
	NOTE: In the current protocol, the hemi-brains were stored at -80 &#176;C for 6-8 months.</li>
	<li>Section the frozen hemi-brain into 20 &#181;m thick sections using a cryostat (see Table of Materials). 
	&#8203;NOTE: For the current protocol, the hemi-brains were sectioned into sagittal sections, and if needed, coronal sections can also be prepared17. Step 2 is for A&#946; immunofluorescent staining on fixed and cryoprotected brain tissue samples.</li>
</ol>
2. Immunofluorescence
<ol>
	<li>Place the sagittal brain tissue sections (step 1.5) into a 24-well plate (up to six mouse brain sections per 300 &#181;L per well). Wash for 5 min with 1x PBS three times at room temperature (23 &#176;C) by placing the plate on a shaker with gentle swirling.</li>
	<li>Incubate the brain tissue sections with 70% formic acid in dH20 at room temperature for 10 min. 
	CAUTION: Formic acid is corrosive, so avoid skin and eye contact.</li>
	<li>Wash the brain tissue sections for 5 min with dH20 three times at room temperature.</li>
	<li>Block non-specific binding with 0.5% bovine serum albumin (BSA) and 0.3% TritonX 100 (see Table of Materials) in 1x PBS at room temperature for 1 h.</li>
	<li>Incubate the brain tissue sections with a fluorophore-labeled primary antibody (6E10, see Table of Materials) diluted (1:1000) in 1x PBS containing 0.3% TritonX 100 at 4 &#176;C for 24 h. 
	NOTE: Since the primary antibody is fluorophore-conjugated, the plate needs to be covered from this step onward, or all work must be done in a dark room to maintain fluorophore effectiveness.</li>
	<li>Wash the brain tissue sections for 10 min with 1x PBS three times at room temperature.</li>
	<li>Mount the brain tissue sections onto positively-charged glass slides (see Table of Materials) that are well labeled (label information on the slide is based on preference), after a brief wash with dH20 to remove remaining salts. Let the slides air dry in the dark. 
	NOTE: The brain sections must be mounted carefully to avoid folds and rips, impacting data quantification. In case of rips and/or folds that lie in the region of interest and can interfere with data quantification, re-staining and re-mounting are recommended.</li>
	<li>Mount the brain tissue sections with an aqueous mounting media (see Table of Materials) and place the glass coverslip over the tissue. Seal the ends of the coverslip with clear nail polish and store the slides in a slide box at 4 &#176;C until imaging. 
	&#8203;NOTE: In the current protocol, the slides were imaged within 1 month after staining.</li>
</ol>
3. Imaging
<ol>
	<li>Image the 6E10-stained brain sections using a fluorescence (epifluorescence or confocal) microscope (see Table of Materials), that has a 2x objective to capture the entire brain tissue section in one image and is equipped with the appropriate filter (GFP in this work). 
	NOTE: Imaging settings must be consistent across different slides.</li>
	<li>Save the captured images as a TIFF file or as required and open them in the image analysis software (see Table of Materials) as described in step 4.2 below for 6E10 quantification. 
	&#8203;NOTE: Include a scale bar before capturing the image to quantify in the image analysis software, ImageJ.</li>
</ol>
4. Full-region of interest analysis
NOTE: The present work&#39;s two regions of interest are the hippocampus and the iso-cortex. Full-region of interest analysis represents the analysis of the entire iso-cortex (referred to as the cortex going forward) or the hippocampus in the imaged brain tissue section.
<ol>
	<li>Download the image analysis software (see Table of Materials) and start the software once installed.</li>
	<li>Once the software runs, click on File | Open | Choose the image to be analyzed.</li>
	<li>Click on Analyze | Set scale| Click to remove scale.&#160;Select the&#160;Straight&#160;tool from the software tool bar and draw a straight line along the length of the scale bar. Click on&#160;Analyze | Measure. Note the length or distance of the scale bar in pixels. Click on&#160;Analyze | Set scale.
	<ol>
		<li>In the pop-up window, enter the distance in pixels, the known distance of the scale bar (in &#181;m in this case), and the unit of length as &#181;m. Check&#160;Global&#160;to apply the new scale setting to all following images if multiple images are processed. Click&#160;OK&#160;to apply the settings. 
		NOTE: It is always recommended to check if the accurate scale is applied to the image before further analysis.</li>
	</ol>
	</li>
	<li>To set the desired measurement to the area of the section, go to Analyze | Set measurements | Select the Area and Display Label boxes. Check that the image being analyzed is selected under Redirect to.</li>
	<li>For ease of hippocampus or cortex visualization, go to Image | Adjust | Brightness/Contrast. Drag the Maximum slide bar gradually to the left to increase tissue clarity until the brain regions of interest are identifiable. 
	NOTE: Do not apply this setting to avoid false measurements during analysis, instead proceed to the next step.</li>
	<li>Use the Polygon selection or Freehand selection tool to outline the hippocampus region. Click on the Reset option of the Brightness/Contrast settings once the hippocampus is outlined to revert to the original brightness. 
	NOTE: The steps must be repeated separately for the cortical region.</li>
	<li>To measure the total tissue area of the selected region, click on Edit | Clear outside. Once the selected region is the only image on-screen, click on Analyze | Measure to obtain the total tissue area analyzed in a pop-up window. Save the data in an Excel file for later use.</li>
	<li>To measure the 6E10-positive area, go to Image | Adjust | Color threshold. A pre-made filter under the Thresholding method usually provides desired results highlighting the strongest signals in red. 
	NOTE: The optimal threshold selection will depend on the background of the image and the staining intensity. Select a threshold setting that picks up the stain and not the background.</li>
	<li>After selecting the appropriate threshold, check Dark background. This will highlight the A&#946; spots (the stain of interest) on a black background. Click on Select | Original | Select, giving the dark signals (the A&#946; deposits) on a white background. Click on Analyze | Analyze particles and click OK when the pop-up window generates.</li>
	<li>Copy the summary output generated by the software by clicking on Edit | Copy. Paste into the previously started Excel file with respective labels. This is the 6E10-positive area in the selected regions of interest (either hippocampus or cortex). 
	&#8203;NOTE: In Excel, there will be a column for the total 6E10-positive area (step 4.10) and the total tissue area (step 4.7).</li>
	<li>Calculate the 6E10-positive area (%) as follows16: (Total 6E10-positive area/Total tissue area analyzed) x 100.</li>
</ol>
5. Sub-region of interest analysis
NOTE: Sub-region of interest analysis represents the analysis of a part of the cortex or the hippocampus in the imaged brain tissue section.
<ol>
	<li>Download the image analysis software and start the software once installed.</li>
	<li>Once the software runs, click on File | Open | Choose the image to be analyzed.</li>
	<li>Click on&#160;Analyze | Set scale| Click to remove scale.&#160;Select the&#160;Straight&#160;tool from the software toolbar and draw a straight line along the length of the scale bar. Click on&#160;Analyze | Measure. Note the length or distance of the scale bar in pixels. Click on&#160;Analyze | Set scale.
	<ol>
		<li>In the pop-up window, enter the distance in pixels, the known distance of the scale bar (in &#181;m in this case), and enter the unit of length as &#181;m. Check&#160;Global&#160;to apply the new scale setting to all following images if multiple images are processed. Click&#160;OK&#160;to apply the settings. 
		NOTE: It is always recommended to check if the accurate scale is applied to the image before further analysis.</li>
	</ol>
	</li>
	<li>To set the desired measurement to the area of the section, go to Analyze | Set measurements | select the Area and Display Label boxes. Check that the image being analyzed is selected under Redirect to.</li>
	<li>Adjust the brightness and contrast if the image is too dim and the brain regions (e.g., the hippocampus or the cortex in this case) cannot be easily identified. Use the software toolbar and click on Image | Adjust | Brightness/Contrast and drag the Maximum sliders to the left as needed to increase the tissue visibility. 
	NOTE: Do not apply this setting to avoid false measurements during analysis, instead proceed to the next step.</li>
	<li>Using the Rectangle tool, select the region of interest in the cortex or the hippocampus. Use the toolbar and click on Edit | Selection | Specify, changing the height and width to a pre-defined value. Adjust the box, so it is completely covered by tissue. Reset the brightness/contrast to revert to the original brightness. 
	NOTE: The size of the box used to select the regions of interest must be consistent for all the images. For the present analysis, the box size was either 300 pixels x 300 pixels (equivalent to 1177 &#181;m x 1177 &#181;m) or 400 pixels x 200 pixels (equivalent to 1569 &#181;m x 784 &#181;m).</li>
	<li>Duplicate the selected region of interest by right-clicking the box and clicking on Duplicate. A new window with the selected region will open. Rename the duplicated image to display the region it is located in (e.g., cortex or hippocampus).</li>
	<li>Adjust the duplicated image type by using the toolbar and clicking on Image | Type | 8-Bit to convert the duplicated RGB image to 8-bit to best analyze the plaques. Invert the image by clicking on Edit | Invert.</li>
	<li>To measure the 6E10-positive area, go to Image | Adjust | Threshold. A pre-defined filter under the Thresholding method usually provides the desired results by highlighting the strongest signals in red. 
	NOTE: The optimal threshold selection will depend on the background of the image and the staining intensity. Select a threshold setting that picks up the stain and not the background.</li>
	<li>After selecting the appropriate threshold, select Apply.</li>
	<li>To analyze the 6E10-positive area, use the toolbar and click on Analyze | Analyze particles, ensuring the &#34;Summarize Results&#34; is checked.</li>
	<li>Copy the Summary output with the %Area generated by the software by clicking on Edit | Copy. Paste into the previously started Excel file with respective labels.</li>
	<li>Repeat steps 5.3-5.12 for the different regions in the tissue. Ensure the placement of each box to outline the region of interest is consistent between each image. 
	NOTE: The rotation tool can be utilized if the Rectangle toolbox dimensions cannot fit into the specified region due to tissue curvature.</li>
	<li>To rotate the rectangle, use the toolbar and click on Edit | Selection | Rotate and adjust the rotation degree as needed. Duplicate the image as mentioned in step 5.7 and clear the outside by using the toolbar and clicking on Edit | Clear outside, which clears the 6E10 stains outside the specified rectangle. Proceed with step 5.8 described above.</li>
</ol>

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S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Amyloid-beta+and+tau:+the+trigger+and+bullet+in+Alzheimer+disease+pathogenesis.">Amyloid-beta and tau: the trigger and bullet in Alzheimer disease pathogenesis.</a> JAMA Neurology. 71 (4), 505-508 (2014).</li><li>Gremer, L., 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=Fibril+structure+of+amyloid-beta(1-42)+by+cryo-electron+microscopy.">Fibril structure of amyloid-beta(1-42) by cryo-electron microscopy.</a> Science. 358 (6359), 116-119 (2017).</li><li>Ferrero, J., 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=First-in-human,+double-blind,+placebo-controlled,+single-dose+escalation+study+of+aducanumab+(BIIB037)+in+mild-to-moderate+Alzheimer's+disease.">First-in-human, double-blind, placebo-controlled, single-dose escalation study of aducanumab (BIIB037) in mild-to-moderate Alzheimer's disease.</a> Alzheimer's &amp; Dementia Translational Research &amp; Clinical Interventions. 2 (3), 169-176 (2016).</li><li>Poon, C. 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The authors have nothing to disclose.

The protocol described herein outlines the procedure for hemi-brain preparation for sagittal sectioning, immunofluorescent staining of A&#946; deposits using the 6E10 antibody on free-floating sections, imaging of the A&#946;-stained brain sections followed by quantification of the A&#946; deposits in the cortex and the hippocampus of mouse brain tissue using an image analysis software. While there are published protocols to quantify A&#946; load in brain tissue sections8,10, this protocol describes the steps involved in the quantification of A&#946; load in the entire iso-cortex (referred to as cortex) and hippocampus in comparison with the A&#946; load in a sub-region of interest in the cortex and the hippocampus, when this may be desired. The correlation between the full- versus sub-regions of interest analyses is also provided.
There are several critical steps in the protocol. First, the protocol described is for 20 &#181;m thick brain tissue sections subjected to free-floating immunostaining, which results in optimal antibody penetration within the tissue section18. The free-floating technique may require the tissue sections to be manually transferred between the different solutions during the immunofluorescent staining, and careful handling of the tissue sections throughout the procedure. This is especially crucial when the tissue sections are immersed in the 70% formic acid solution for antigen retrieval in the current protocol, increasing tissue fragility for thin sections. Alternate approaches to the described protocol include using thicker tissue sections (e.g., 30-40 &#181;m) or using tissue sections that are directly mounted onto positively-charged slides before the immunofluorescent staining. Second, the protocol described herein uses a fluorescently-labeled 6E10 antibody. Besides using the fluorescently-labeled 6E10 antibody, non-fluorescent 6E10 antibodies (e.g., horseradish peroxidase-conjugated 6E10 antibody) can also be used to detect A&#946; load in brain tissue sections, and the current protocol can be adapted to quantify A&#946;-positive immunochemical stains in the brain tissue sections as described previously8. Third, the accuracy of the results for A&#946; load quantification will depend on the appropriate threshold selection in the analysis software, which is dependent on the tissue background and signal intensity. Threshold selection must be performed by the end-user such that only A&#946;-positive stains are selected for quantification. End-user intervention is required to optimize the specific threshold that can be applied to all the images to assure the accuracy of the threshold setting. Fourth, since the sub-region of interest analysis requires selecting a small region of interest in the tissue section, two independent readers were used for this analysis. To maintain independence and blinding during data collection, all the images were number coded; the image analysis sequence was randomized between the readers such that the different readers analyzed different images at any given time, and the data was submitted at the end of each week. Due to the increased likelihood of inter-reader variability in the region of interest selection in the sub-region of interest analysis, the readers were trained using several sample images to optimize region selection in the cortex and the hippocampus before beginning the data collection. This training is crucial to reduce inter-reader variability, and as can be seen (Figure 5A,B), the 6E10-positive area reported by both the readers shows a strong relative agreement in the current study.
The current protocol and the results provide valuable information about the impact of the region of interest size on A&#946;-positive area quantification. A larger region of interest is expected to represent the tissue more than a smaller region of interest. Therefore, sampling a larger tissue is desirable to accurately quantify the A&#946; load in tissues. However, in the case of homogenous A&#946; load distribution within the tissue, a smaller sampling region is generally considered a good representation of the larger tissue under analysis. The current study results confirm this, and the A&#946; load in the entire cortex and the hippocampus was a strong correlate of A&#946; load in a selected sub-region of the cortex and the hippocampus (Figure 5C,D). To further confirm the agreement between the full- and sub-regions of interest analyses, the mean 6E10-positive area in the cortex and the hippocampus were compared, and no difference between the two methods (Figure 5E) was found. This confirms that either of these methods (full- or sub-region analysis) yield comparable A&#946; load measurements.
The current protocol has some limitations. The two methods (full-region versus sub-region analysis) may not always be used interchangeably. The choice of using the full- or sub-region analysis will depend on the regional distribution of A&#946; within the tissue, which is impacted by the age, sex, and strain of the AD mouse model. At 13 months, the A&#946; load is distributed throughout the cortex and hippocampus of the APP/PS1 mice. However, at 6 months, A&#946; deposits are limited to the cortex, and minimal deposits are observed in the hippocampus12. Under such conditions, the full-region of interest analysis may be the desired approach to increase the tissue sampling area and thereby the A&#946; signal. On the other hand, sub-region analysis may be the method of choice when A&#946; load in a specific brain region is of interest, (e.g., the somatosensory cortex). Additionally, at 13 months, the APP/PS1 male mice demonstrate intense 6E10-positive stains, and the immunofluorescent staining results in an excellent signal with a very low background, making the current protocol very suitable for quantification under the given conditions. It is unclear if this quantification method can be applied successfully to less intense staining, and future work will be required to answer this question. The immunofluorescent and image quantification method presented herein detects all forms of A&#946;, including the precursor form13. As a result, if there is an interest in the detection of a specific A&#946; specie (e.g., A&#946;1-40 or A&#946;1-42), antibodies specific to these A&#946; isoforms can be used. Therefore, though the 6E10 immunofluorescent staining and detection method correlated with the measurements of A&#946;1-42 measurements in whole-brain homogenates using ELISA (Figure 5F,G), the correlation was only modest. This can be attributed to the measurement of only A&#946;1-42 using the ELISA and detection of all A&#946; species using the 6E10 immunostaining. The current study uses three readers to assess the agreement and correlation between the full- and sub-regional analyses. Having additional readers may improve the robustness of the study and can further validate the agreements between the two methods presented here. Further, we use the Pearson correlation as a measure of agreement, which is widely used among other methods to describe the agreement between continuous variables19. However, one limitation of using the Pearson correlation to determine agreement is that the two methods used herein may provide related results, but one method may result in overall higher values than the other due to systematic bias. Therefore, Pearson correlation is a good measure of relative agreement19. To increase the robustness of the protocol, additional methods to confirm the absolute agreement, such as comparing the mean 6E10-positive area by the two methods (Figure 5E), can be used19. Taken together, the current protocol compares the A&#946; load detected by immunofluorescent staining and analyzes the full- and sub-regions of interest in brain tissue sections. The results show a strong correlation between these two methods for brain tissue sections derived from 13-month-old APP/PS1 male mice that show abundant A&#946; deposits.

Alzheimer&#39;s disease (AD), the sixth leading cause of death in the United States, continues to be a public health threat, with an estimated 6.2 million Americans living with AD. This is expected to reach 13.8 million by 20601. To date, symptomatic management through medications such as cholinesterase inhibitors and memantine is the primary course of treatment2. AD is characterized by neuropathologic manifestations such as extracellular deposition of amyloid-beta (A&#946;) plaques and intracellular hyperphosphorylated tau accumulation in the form of neurofibrillary tangles3,4. Formed by an endoproteolytic cleavage of the amyloid precursor protein (APP) via beta- and gamma-secretase, A&#946; aggregates to form oligomers and fibrils, leading to neurotoxic effects5. A&#946; has been hypothesized to serve a primary pathological role since the 1980s, and is the therapeutic target of the only FDA-approved disease-modifying therapy for AD6. As a result, transgenic AD mouse models harboring mutations in genes resulting in robust cerebral A&#946; accumulation have been widely used for preclinical AD research since the early 1990s7.
Detection of A&#946; species in these AD transgenic mouse brains is generally done using two immunoassays: enzyme-linked immunosorbent assay (ELISA) and immunostaining. The former assay enables quantitative determination of different A&#946; species and is less time-consuming compared to immunostaining, which requires several sequential tissue processing and imaging steps, including tissue sectioning, immunostaining, imaging, and quantification8. Further, the results obtained following immunostaining are semi-quantitative8. However, the ability to spatially localize A&#946; makes immunostaining an attractive approach for A&#946; detection in brain tissues8.
While using A&#946; immunostaining, several different quantification paradigms have been employed by different research groups. For example, some research groups quantify the A&#946; load in the entire region of interest (cortex or hippocampus), while others quantify the A&#946; load in a specified sub-region of interest (a part of the cortex or the hippocampus)9,10,11. Though the methods for A&#946; detection and quantification have been well established and documented, the impact of the size of the region of interest on A&#946; load measurements following immunostaining has not been reported. Therefore, the current protocol aimed to compare the A&#946; load measurements across the full- and sub-regions of interest using an image analysis software, ImageJ.
The current study used 13-month-old APP/PS1 double transgenic male mice, which express a chimeric mouse/human APP and a mutant presenilin 1, to model early-onset of AD12. A&#946; deposits start developing by 6-7 months of age, and abundant A&#946; accumulation is observed both in the cortex and the hippocampus of these mice by 9-10 months of age12. The transgenic amyloid peptides and holoprotein can be detected by 6E10-immunostaining13, making it a desirable animal model for the present protocol. The procedure covered herein includes brain tissue preparation, immunostaining of free-floating sections, imaging, and quantification of A&#946; load in full- versus sub-regions of interest. The analysis shows a strong correlation between the full- and sub-regional quantification, indicating robust agreement between these two methods in the brain tissue sections derived from 13-month-old APP/PS1 male mice that show abundant A&#946; deposits.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>15 mL conical tubes</td>
			<td>ThermoFisher Scientific, MA, USA</td>
			<td>339650</td>
			<td>https://www.thermofisher.com/order/catalog/product/339650</td>
		</tr>
		<tr>
			<td>24-well plates</td>
			<td>Fisher Scientific, NH, USA</td>
			<td>FB012929</td>
			<td>https://www.fishersci.com/shop/products/jet-biofil-surface-treated-steriletissue-culture-plates-3/FB012929</td>
		</tr>
		<tr>
			<td>Amyloid beta 42 human ELISA kit</td>
			<td>ThermoFisher Scientific, MA, USA</td>
			<td>KHB3441</td>
			<td>https://www.thermofisher.com/elisa/product/Amyloid-beta-42-Human-ELISA-Kit/KHB3441</td>
		</tr>
		<tr>
			<td>Aqueous mounting media</td>
			<td>Vector laboratories, CA, USA</td>
			<td>H-5501-60</td>
			<td>https://vectorlabs.com/products/mounting/vectamount-aq-aqueous-mounting-medium</td>
		</tr>
		<tr>
			<td>Bovine serum albumin</td>
			<td>Sigmaaldrich, MO, USA</td>
			<td>A2153-50G</td>
			<td>https://www.sigmaaldrich.com/US/en/product/sigma/a2153?gclid=CjwKCAjw9aiIBhA1EiwAJ_G 
			TSiZ9B3YGz3RvSpWqCH4CPW78 
			Dj4WlgxmzPs631z5IHmy5XLV 
			TdC_jBoC9zQQAvD_BwE</td>
		</tr>
		<tr>
			<td>BZ-X710 Keyence all-in-one fluorescence microscope</td>
			<td>Keyence, IL, USA</td>
			<td>BZ-X710</td>
			<td>https://www.keyence.com/products/microscope/fluorescence-microscope/bz-x700/models/bz-x710/</td>
		</tr>
		<tr>
			<td>Clear nail poilsh</td>
			<td>User preference</td>
			<td>NA</td>
			<td>None</td>
		</tr>
		<tr>
			<td>Cryostat</td>
			<td>Leica Biosystems, IL, USA</td>
			<td>Leica CM1860 Cryostat</td>
			<td>https://www.leicabiosystems.com/us/histology-equipment/cryostats/leica-cm1860/</td>
		</tr>
		<tr>
			<td>Formic acid</td>
			<td>Sigmaaldrich, MO, USA</td>
			<td>F0507-500ML</td>
			<td>https://www.sigmaaldrich.com/US/en/product/sigald/f0507?gclid=CjwKCAjw9aiIBhA1EiwAJ_G 
			TSheH6JMGnla50C3Ag0cLzXE8 
			BObxvDApl0udjYAPZmBGe7a 
			8PRUv1RoCt34QAvD_BwE</td>
		</tr>
		<tr>
			<td>Glass coverslips</td>
			<td>VWR, PA, USA</td>
			<td>48393-081</td>
			<td>https://us.vwr.com/store/product/4645817/vwr-micro-cover-glasses-rectangular</td>
		</tr>
		<tr>
			<td>GraphPad Prism</td>
			<td>GraphPad Software, CA, USA</td>
			<td>Version 8</td>
			<td>https://www.graphpad.com/scientific-software/prism/</td>
		</tr>
		<tr>
			<td>ImageJ 1.51k</td>
			<td>National Institutes of Health, MD, USA</td>
			<td>Version 1.53e</td>
			<td>https://imagej.nih.gov/ij/download.html</td>
		</tr>
		<tr>
			<td>Mice</td>
			<td>Jackson Laboratories, ME, USA</td>
			<td>034829-JAX</td>
			<td>https://www.mmrrc.org/catalog/sds.php?mmrrc_id=34829</td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Sigmaaldrich, MO, USA</td>
			<td>P6148-500G</td>
			<td>https://www.sigmaaldrich.com/US/en/product/sial/p6148?gclid=CjwKCAjw9aiIBhA1EiwAJ_G 
			TShtLb9Ax9MmRyrFn6Rfmmg 
			1l52_5XZFXOeXT24ik8Lkw 
			GH7fvlDoHBoChzYQAvD_BwE&#160;</td>
		</tr>
		<tr>
			<td>Phenytoin/pentobarbital based anesthetic (Euthasol)</td>
			<td>Patterson Veterinary, MA, USA</td>
			<td>07-805-9296</td>
			<td>https://www.pattersonvet.com/Supplies/ProductFamilyDetails/PIF_32818</td>
		</tr>
		<tr>
			<td>Phosphate-buffered saline</td>
			<td>Fisher Scientific,&#160; NH, USA</td>
			<td>BP661-50</td>
			<td>https://www.fishersci.com/shop/products/pbs-1x-powder-concentrate-white-granular-powder-fisher-bioreagents-2/BP66150</td>
		</tr>
		<tr>
			<td>Plus (+) microscope slides</td>
			<td>Ted Pella, Inc., CA, USA</td>
			<td>260100</td>
			<td>https://www.tedpella.com/histo_html/slides.htm#260384</td>
		</tr>
		<tr>
			<td>Primary antibody (6E10)</td>
			<td>Biolegend, CA, USA</td>
			<td>803013</td>
			<td>https://www.biolegend.com/en-us/products/alexa-fluor-488-anti-beta-amyloid-1-16-antibody-10833</td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Sigmaaldrich, MO, USA</td>
			<td>47289</td>
			<td>https://www.sigmaaldrich.com/US/en/product/supelco/47289?gclid=CjwKCAjw9aiIBhA1EiwAJ_ 
			GTSuwlymWL_PUl2KIMHymi 
			GLOWluZdPjf3pRcjMEjQD 
			siItfWiG-C2-RoCxyoQAvD_BwE</td>
		</tr>
		<tr>
			<td>Triton X 100</td>
			<td>Sigmaaldrich, MO, USA</td>
			<td>T8787-100ML</td>
			<td>https://www.sigmaaldrich.com/US/en/product/sigma/t8787?context=product</td>
		</tr>
	</tbody>
</table>

full- versus sub-regional quantification of amyloid-beta load on mouse brain sections

Extracellular accumulation of amyloid-beta (A&#946;) plaques is one of the major pathological hallmarks of Alzheimer's disease (AD), and is the target of the only FDA-approved disease-modifying treatment for AD. Accordingly, the use of transgenic mouse models that overexpress the amyloid precursor protein and thereby accumulate cerebral A&#946; plaques are widely used to model human AD in mice. Therefore, immunoassays, including enzyme-linked immunosorbent assay (ELISA) and immunostaining, commonly measure the A&#946; load in brain tissues derived from AD transgenic mice. Though the methods for A&#946; detection and quantification have been well established and documented, the impact of the size of the region of interest selected in the brain tissue on A&#946; load measurements following immunostaining has not been reported. Therefore, the current protocol aimed to compare the A&#946; load measurements across the full- and sub-regions of interest using an image analysis software. The steps involved in brain tissue preparation, free-floating brain section immunostaining, imaging, and quantification of A&#946; load in full- versus sub-regions of interest are described using brain sections derived from 13-month-old APP/PS1 double transgenic male mice. The current protocol and the results provide valuable information about the impact of the size of the region of interest on A&#946;-positive area quantification, and show a strong correlation between the A&#946;-positive area obtained using the full- and sub-regions of interest analyses for brain sections derived from 13-month-old male APP/PS1 mice that show widespread A&#946; deposition.

Here, two different methods are compared to quantify the 6E10-positive area in the hippocampus and the cortex of mouse brain tissues. The two methods are the full-region and sub-region of interest analyses (Figure 1). The full-region of interest analysis, as the name suggests, involves outlining the entire region of interest (in this case, either the iso-cortex or the hippocampus) to determine the 6E10-positive area (Figure 1A,B). The sub-region of interest analysis involves selecting a pre-defined region within the region of interest to determine the 6E10-positive area (Figure 1C,D). The stepwise ImageJ protocol for the two methods is shown in Figure 2, Figure 3, and Figure 4.
This study used three readers; two independent readers performed the sub-region of interest analysis, and the third reader performed the full-region of interest analysis. As seen in Figure 5A,B, there was a strong significant positive correlation (p &#60; 0.0001) between the 6E10-positive area reported by the two readers performing the sub-region of interest analysis (Pearson correlation coefficient r = 0.97 for the cortex and r = 0.96 for the hippocampus). The 6E10-positive areas reported by the two readers for the sub-region of interest analysis were averaged, and the averaged sub-region of interest 6E10-positive area shared a strong significant positive correlation (p &#60; 0.0001) with the 6E10-positive area obtained using the full-region of interest analysis for both the cortex (Pearson correlation coefficient r = 0.96; Figure 5C) and the hippocampus (Pearson correlation coefficient r = 0.95; Figure 5D). The mean cortical- and hippocampal-6E10-positive area obtained by the full-region and sub-region of interest analyses were comparable with no significant difference, confirming the agreement between the two methods (Figure 5E). Further, the insoluble A&#946;1-42 was measured in whole-brain homogenates in a subset of mice and the cortical (Figure 5F) and hippocampal (Figure 5G) 6E10-positive area determined by the full-region of interest analysis was significantly (p &#60; 0.01) correlated with insoluble A&#946;1-42 load using ELISA (see Table of Materials).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63669/63669fig01.jpg" /> 
Figure 1: Full- versus sub-regions of interest selection. Representative images showing the full iso-cortex (cortex) and hippocampus outlined for the full-region of interest analysis in (A) and (B), respectively. Representative images show the selection of sub-region/s of the cortex and hippocampus for the sub-region of interest analysis in (C) and (D), respectively. Scale bar = 200 &#181;m. <a href="https://www.jove.com/files/ftp_upload/63669/63669fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/63669/63669fig02.jpg" /> 
Figure 2: Protocol for the full-region of interest 6E10-positive area quantification. Image analysis steps showing the original image (A), the image after brightness/contrast adjustment (B), selection of the area of interest (C), clearing (D), threshold adjustment (E), and the final image ready for analysis (F). The numbers in the figure designate the step numbers in the protocol. Scale bar = 200 &#181;m. <a href="https://www.jove.com/files/ftp_upload/63669/63669fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/63669/63669fig03.jpg" /> 
Figure 3: Protocol for the sub-region of interest 6E10-positive area quantification. Image analysis steps showing the original image (A), the image after brightness/contrast adjustment (B), selection of the area of interest (C), duplication of the image of the region of interest (D), changing the image to 8-Bit and inverting the image (E), threshold adjustment (F), and final image ready for analysis (G). The numbers in the figure designate the step numbers in the protocol. Scale bar = 200 &#181;m. <a href="https://www.jove.com/files/ftp_upload/63669/63669fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/63669/63669fig04.jpg" /> 
Figure 4: Protocol for the region of interest rotation. Image analysis steps showing the selection of the area of interest and rotation of the selection box to fit the tissue curvature (A), the image of the region of interest after duplication (B), and the image after clearing the outside (non-region of interest) area (C). The numbers in the figure designate the step number in the protocol. Scale bar = 200 &#956;m. <a href="https://www.jove.com/files/ftp_upload/63669/63669fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/63669/63669fig05.jpg" /> 
Figure 5: Correlation between the full- and sub-regions of interest analyses. Scatter plots show the correlation between the 6E10-positive area by the two independent readers performing the sub-region of interest analysis for the cortex (A) and the hippocampus (B). A strong positive correlation is observed between the 6E10-positive area resulting from the sub-region of interest analysis and the full-region of interest analysis for both the cortex (C) and the hippocampus (D). There is no statistically significant difference in the mean 6E10-positive area by the full-region of interest and the sub-region of interest analyses in the cortex and the hippocampus (E). A significant correlation is observed between whole-brain homogenate insoluble A&#946;1-42 measurements using ELISA and the full-region of interest analysis for both the cortex (F) and the hippocampus (G). Data were analyzed using the Pearson correlation coefficient, r, in (A-D) and (F-G), and using two-way repeated-measures ANOVA in (E) using a graphing and statistics software. Data are presented as mean &#177; standard error of the mean (SEM) of n = 35 mice in (E), and a two-tailed p &#60; 0.05 was considered statistically significant. <a href="https://www.jove.com/files/ftp_upload/63669/63669fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Full- versus Sub-Regional Quantification of Amyloid-Beta Load on Mouse Brain Sections at JoVE.com

Full- versus Sub-Regional Quantification of Amyloid-Beta Load on Mouse Brain Sections

The present protocol describes and compares the procedure to perform a full-region or sub-region of interest analysis of sagittal mouse brain sections to quantify amyloid-beta load in the APP/PS1 transgenic mouse model of Alzheimer's disease.

Extracellular accumulation of amyloid-beta (A&#946;) plaques is one of the major pathological hallmarks of Alzheimer's disease (AD), and is the target of ...

full-versus-sub-regional-quantification-amyloid-beta-load-on-mouse

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3.4K Views.  Keck Graduate Institute. The present protocol describes and compares the procedure to perform a full-region or sub-region of interest analysis of sagittal mouse brain sections to quantify amyloid-beta load in the APP/PS1 transgenic mouse model of Alzheimer's disease. 

Video: Full- versus Sub-Regional Quantification of Amyloid-Beta Load on Mouse Brain Sections

Preparation of Oligomeric &beta;-amyloid1-42 and Induction of Synaptic Plasticity Impairment on Hippocampal Slices

Impairment of synaptic connections is likely to underlie the subtle amnesic changes occurring at the early stages of Alzheimer s Disease (AD). &beta;-amyloid (A&beta;), a peptide produced in high amounts in AD, is known to reduce Long-Term Potentiation (LTP), a cellular correlate of learning and memory. Indeed, LTP impairment caused by A&beta; is a useful experimental paradigm for studying synaptic dysfunctions in AD models and for screening drugs capable of mitigating or reverting such synaptic impairments. Studies have shown that A&beta; produces the LTP disruption preferentially via its oligomeric form. Here we provide a detailed protocol for impairing LTP by perfusion of oligomerized synthetic A&beta;1-42 peptide onto acute hippocampal slices. In this video, we outline a step-by-step procedure for the preparation of oligomeric A&beta;1-42. Then, we follow an individual experiment in which LTP is reduced in hippocampal slices exposed to oligomerized A&beta;1-42 compared to slices in a control experiment where no A&beta;1-42 exposure had occurred.

Preparation of Oligomeric &amp;#946;-amyloid1-42 and Induction of Synaptic Plasticity Impairment on Hippocampal Slices

One feature of Alzheimer's Disease is the elevation of A&beta;1-42 peptide. Here we provide a protocol for preparing synthetic A&beta;1-42 oligomers, which impairs hippocampal Long-Term Potentiation, a cellular correlate of memory. This procedure is useful for investigating mechanisms of A&beta;-induced pathology and drug screening.

Impairment of synaptic connections is likely to underlie the subtle amnesic changes occurring at the early stages of Alzheimer s Disease (AD). ...

Double Fluorescence in situ Hybridization in Fresh Brain Sections

Here we describe a modified version of a double fluorescence in situ hybridization (dFISH) method optimized for detecting two mRNAs of interest in fresh frozen brain sections. Our group has successfully used this approach to study gene co-regulation. More specifically, we have used this dFISH method to explore the anatomical organization, neurochemical properties, and the impact of sensory experience in central sensory circuits, at single cell resolution. This protocol has been validated in brain tissue from mice, rats and songbirds but is expected to be easily adaptable to other vertebrate species, as well as to an array of non-neural tissues. In this film we provide a detailed demonstration of the main steps of this procedure.

Double Fluorescence in situ Hybridization in Fresh Brain Sections

This protocol involves a non-radioactive in-situ hybridization procedure that enables the simultaneous identification of two transcript species, at a single cell resolution, in thin sections of the vertebrate brain.

Here we describe a modified version of a double fluorescence in situ hybridization (dFISH) method optimized for detecting two mRNAs of interest in fresh ...

Vibrodissociation of Neurons from Rodent Brain Slices to Study Synaptic Transmission and Image Presynaptic Terminals

Mechanical dissociation of neurons from the central nervous system has the advantage that presynaptic boutons remain attached to the isolated neuron of interest. This allows for examination of synaptic transmission under conditions where the extracellular and postsynaptic intracellular environments can be well controlled. A vibration-based technique without the use of proteases, known as vibrodissociation, is the most popular technique for mechanical isolation. A micropipette, with the tip fire-polished to the shape of a small ball, is placed into a brain slice made from a P1-P21 rodent. The micropipette is vibrated parallel to the slice surface and lowered through the slice thickness resulting in the liberation of isolated neurons. The isolated neurons are ready for study within a few minutes of vibrodissociation. This technique has advantages over the use of primary neuronal cultures, brain slices and enzymatically isolated neurons including: rapid production of viable, relatively mature neurons suitable for electrophysiological and imaging studies; superior control of the extracellular environment free from the influence of neighboring cells; suitability for well-controlled pharmacological experiments using rapid drug application and total cell superfusion; and improved space-clamp in whole-cell recordings relative to neurons in slice or cell culture preparations. This preparation can be used to examine synaptic physiology, pharmacology, modulation and plasticity. Real-time imaging of both pre- and postsynaptic elements in the living cells and boutons is also possible using vibrodissociated neurons. Characterization of the molecular constituents of pre- and postsynaptic elements can also be achieved with immunological and imaging-based approaches.

This report demonstrates a technique for mechanical isolation of individual viable neurons retaining attached presynaptic boutons. Vibrodissociated neurons have the advantages of rapid production, excellent pharmacological control and improved space-clamp without influence from neighboring cells. This method can be used for imaging of synaptic elements and patch-clamp recording.

Mechanical dissociation of neurons from the central nervous system has the advantage that presynaptic boutons remain attached to the isolated neuron of ...

Multi-electrode Array Recordings of Neuronal Avalanches in Organotypic Cultures

The cortex is spontaneously active, even in the absence of any particular input or motor output. During development, this activity is important for the migration and differentiation of cortex cell types and the formation of neuronal connections1. In the mature animal, ongoing activity reflects the past and the present state of an animal into which sensory stimuli are seamlessly integrated to compute future actions. Thus, a clear understanding of the organization of ongoing i.e. spontaneous activity is a prerequisite to understand cortex function. 
Numerous recording techniques revealed that ongoing activity in cortex is comprised of many neurons whose individual activities transiently sum to larger events that can be detected in the local field potential (LFP) with extracellular microelectrodes, or in the electroencephalogram (EEG), the magnetoencephalogram (MEG), and the BOLD signal from functional magnetic resonance imaging (fMRI). The LFP is currently the method of choice when studying neuronal population activity with high temporal and spatial resolution at the mesoscopic scale (several thousands of neurons). At the extracellular microelectrode, locally synchronized activities of spatially neighbored neurons result in rapid deflections in the LFP up to several hundreds of microvolts. When using an array of microelectrodes, the organizations of such deflections can be conveniently monitored in space and time. 
Neuronal avalanches describe the scale-invariant spatiotemporal organization of ongoing neuronal activity in the brain2,3. They are specific to the superficial layers of cortex as established in vitro4,5, in vivo in the anesthetized rat 6, and in the awake monkey7. Importantly, both theoretical and empirical studies2,8-10 suggest that neuronal avalanches indicate an exquisitely balanced critical state dynamics of cortex that optimizes information transfer and information processing.
In order to study the mechanisms of neuronal avalanche development, maintenance, and regulation, in vitro preparations are highly beneficial, as they allow for stable recordings of avalanche activity under precisely controlled conditions. The current protocol describes how to study neuronal avalanches in vitro by taking advantage of superficial layer development in organotypic cortex cultures, i.e. slice cultures, grown on planar, integrated microelectrode arrays (MEA; see also 11-14).

A robust way to study neuronal avalanches, i.e. scale-invariant spatio-temporal activity bursts, indicative of critical state dynamics in cortex. Avalanches emerge spontaneously in developing superficial layers of cultured cortex which allows for long-term measurements of the activity with planar integrated multi-electrode arrays (MEA) under precisely controlled conditions.

The cortex is spontaneously active, even in the absence of any particular input or motor output.  During development, this activity is important for the ...

Morphometric Analyses of Retinal Sections

Morphometric analyses of retinal sections have been used in examining retinal diseases. For examples, neuronal cells were significantly lost in the retinal ganglion cell layer (RGCL) in rat models with N-methyl-D-aspartate (NMDA)&#x2013;induced excitotoxicity1, retinal ischemia-reperfusion injury2 and glaucoma3. Reduction of INL and inner plexiform layer (IPL) thicknesses were reversed with citicoline treatment in rats' eyes subjected to kainic acid-mediated glutamate excitotoxicity4. Alteration of RGC density and soma sizes were observed with different drug treatments in eyes with elevated intraocular pressure3,5,6. Therefore, having objective methods of analyzing the retinal morphometries may be of great significance in evaluating retinal pathologies and the effectiveness of therapeutic strategies.
The retinal structure is multi-layers and several different kinds of neurons exist in the retina. The morphometric parameters of retina such as cell number, cell size and thickness of different layers are more complex than the cell culture system. Early on, these parameters can be detected using other commercial imaging software. The values are normally of relative value, and changing to the precise value may need further accurate calculation. Also, the tracing of the cell size and morphology may not be accurate and sensitive enough for statistic analysis, especially in the chronic glaucoma model. The measurements used in this protocol provided a more precise and easy way. And the absolute length of the line and size of the cell can be reported directly and easy to be copied to other files. For example, we traced the margin of the inner and outer most nuclei in the INL and formed a line then using the software to draw a 90 degree angle to measure the thickness. While without the help of the software, the line maybe oblique and the changing of retinal thickness may not be repeatable among individual observers. In addition, the number and density of RGCs can also be quantified. This protocol successfully decreases the variability in quantitating features of the retina, increases the sensitivity in detecting minimal changes.
 	This video will demonstrate three types of morphometric analyses of the retinal sections. They include measuring the INL thickness, quantifying the number of RGCs and measuring the sizes of RGCs in absolute value. These three analyses are carried out with Stereo Investigator (MBF Bioscience &mdash; MicroBrightField, Inc.). The technique can offer a simple but scientific platform for morphometric analyses.

This video demonstrates three types of morphometric analyses of the retina, which include measuring the inner nuclear layer thickness, quantifying the number of retinal ganglion cells (RGCs) and measuring the sizes of RGCs. The technique can offer a simple but scientific platform for morphometric analyses.

Morphometric analyses of retinal sections have been used in examining retinal diseases. For examples, neuronal cells were significantly lost in the ...

Stereotaxic Infusion of Oligomeric Amyloid-beta into the Mouse Hippocampus

Alzheimer&#8217;s disease is a neurodegenerative disease affecting the aging population. A key neuropathological feature of the disease is the over-production of amyloid-beta and the deposition of amyloid-beta plaques in brain regions of the afflicted individuals. Throughout the years scientists have generated numerous Alzheimer&#8217;s disease mouse models that attempt to replicate the amyloid-beta pathology. Unfortunately, the mouse models only selectively mimic the disease features. Neuronal death, a prominent effect in the brains of Alzheimer&#8217;s disease patients, is noticeably lacking in these mice. Hence, we and others have employed a method of directly infusing soluble oligomeric species of amyloid-beta - forms of amyloid-beta that have been proven to be most toxic to neurons - stereotaxically into the brain. In this report we utilize male C57BL/6J mice to document this surgical technique of increasing amyloid-beta levels in a select brain region. The infusion target is the dentate gyrus of the hippocampus because this brain structure, along with the basal forebrain that is connected by the cholinergic circuit, represents one of the areas of degeneration in the disease. The results of elevating amyloid-beta in the dentate gyrus via stereotaxic infusion reveal increases in neuron loss in the dentate gyrus within 1 week, while there is a concomitant increase in cell death and cholinergic neuron loss in the vertical limb of the diagonal band of Broca of the basal forebrain. These effects are observed up to 2 weeks. Our data suggests that the current amyloid-beta infusion model provides an alternative mouse model to address region specific neuron death in a short-term basis. The advantage of this model is that amyloid-beta can be elevated in a spatial and temporal manner.

Here, we present a protocol for direct stereotaxic brain infusion of amyloid-beta. This methodology provides an alternative in vivo mouse model to address the short-term effects of amyloid-beta on brain neurons.

Alzheimer&#8217;s disease is a neurodegenerative disease affecting the aging population. A key neuropathological feature of the disease is the ...

Combining Double Fluorescence In Situ Hybridization with Immunolabelling for Detection of the Expression of Three Genes in Mouse Brain Sections

Detection of gene expression in different types of brain cells e.g., neurons, astrocytes, oligodendrocytes, oligodendrocyte precursors and microglia, can be hampered by the lack of specific primary or secondary antibodies for immunostaining. Here we describe a protocol to detect the expression of three different genes in the same brain section using double fluorescence in situ hybridization with two gene-specific probes followed by immunostaining with an antibody of high specificity directed against the protein encoded by a third gene. The Aspartoacyclase (ASPA) gene, mutations of which can lead to a rare human white matter disease - Canavan disease - is thought to be expressed in oligodendrocytes and microglia but not in astrocytes and neurons. However, the precise expression pattern of ASPA in the brain has yet to be established. This protocol has allowed us to determine that ASPA is expressed in a subset of mature oligodendrocytes and it can be generally applied to a wide range of gene expression pattern studies.

Combining Double Fluorescence In Situ Hybridization with Immunolabelling for Detection of the Expression of Three Genes in Mouse Brain Sections

Localizing gene expression to specific cell types can be challenging due to the lack of specific antibodies. Here we describe a protocol for simultaneous triple detection of gene expression by combining double fluorescence RNA in situ hybridization with immunostaining.

Detection of gene expression in different types of brain cells e.g., neurons, astrocytes, oligodendrocytes, oligodendrocyte precursors and microglia, can ...

Preparation of Mouse Retinal Cryo-sections for Immunohistochemistry

Preparation of high-quality mouse eye sections for immunohistochemistry (IHC) is critical for assessing the retinal structure and function and for determining the mechanisms underlying retinal diseases. Maintaining structural integrity throughout the tissue preparation is vital for obtaining reproducible retinal IHC data but can be challenging due to the fragility and complexity of retinal cytoarchitecture. Strong fixatives like 10% formalin or Bouin's solution optimally preserve the retinal structure, they often impede IHC analysis by enhancing the background fluorescence and/or diminishing antibody-epitope interactions, a process known as epitope masking. Milder fixatives, on the other hand, like 4% paraformaldehyde, reduces background fluorescence and epitope-masking, meticulous dissection techniques must be utilized to preserve the retinal structure. In this article, we present a comprehensive method to prepare mouse ocular posterior cups for IHC that is sufficient to preserve most antibody-epitope interactions without loss of retinal structural integrity. We include representative IHC with antibodies to various retinal cell type markers to illustrate tissue preservation and orientation under optimal and sub-optimal conditions. Our goal is to optimize IHC studies of the retina by providing a complete protocol from ocular posterior cup dissection to IHC.

This report describes comprehensive methods for preparing frozen mouse retina sections for immunohistochemistry (IHC). Methods described include dissection of the ocular posterior cup, paraformaldehyde fixation, embedding in Optimal Cutting Temperature (OCT) media and tissue orientation, sectioning and immunostaining.

Preparation of high-quality mouse eye sections for immunohistochemistry (IHC) is critical for assessing the retinal structure and function and for ...

Combining Multiplex Fluorescence In Situ Hybridization with Fluorescent Immunohistochemistry on Fresh Frozen or Fixed Mouse Brain Sections

Fluorescent in situ hybridization (FISH) is a molecular technique that identifies the presence and spatial distribution of specific RNA transcripts within cells. Neurochemical phenotyping of functionally identified neurons usually requires concurrent labelling with multiple antibodies (targeting protein) using immunohistochemistry (IHC) and optimization of in situ hybridization (targeting RNA), in tandem. A &#34;neurochemical signature&#34; to characterize particular neurons may be achieved however complicating factors include the need to verify FISH and IHC targets before combining the methods, and the limited number of RNAs and proteins that may be targeted simultaneously within the same tissue section.
Here we describe a protocol, using both fresh frozen and fixed mouse brain preparations, which detects multiple mRNAs and proteins in the same brain section using RNAscope FISH followed by fluorescence immunostaining, respectively. We use the combined method to describe the expression pattern of low abundance mRNAs (e.g., galanin receptor 1) and high abundance mRNAs (e.g., glycine transporter 2), in immunohistochemically identified brainstem nuclei.
Key considerations for protein labelling downstream of the FISH assay extend beyond tissue preparation and optimization of FISH probe labelling. For example, we found that antibody binding and labelling specificity can be detrimentally affected by the protease step within the FISH probe assay. Proteases catalyze hydrolytic cleavage of peptide bonds, facilitating FISH probe entry into cells, however they may also digest the protein targeted by the subsequent IHC assay, producing off target binding. The subcellular location of the targeted protein is another factor contributing to IHC success following FISH probe assay. We observed IHC specificity to be retained when the targeted protein is membrane bound, whereas IHC targeting cytoplasmic protein required extensive troubleshooting. Finally, we found handling of slide-mounted fixed frozen tissue more challenging than fresh frozen tissue, however IHC quality was overall better with fixed frozen tissue, when combined with RNAscope.

Combining Multiplex Fluorescence In Situ Hybridization with Fluorescent Immunohistochemistry on Fresh Frozen or Fixed Mouse Brain Sections

This protocol describes a method for combining fluorescence in situ hybridization (FISH) and fluorescence immunohistochemistry (IHC) in both fresh frozen and fixed mouse brain sections, with the goal of achieving multilabel FISH and fluorescence IHC signal. IHC targeted cytoplasmic and membrane attached proteins.

Fluorescent in situ hybridization (FISH) is a molecular technique that identifies the presence and spatial distribution of specific RNA transcripts within ...

Quantification of Immunostained Caspase-9 in Retinal Tissue

The family of caspases is known to mediate many cellular pathways beyond cell death, including cell differentiation, axonal pathfinding, and proliferation. Since the identification of the family of cell death proteases, there has been a search for tools to identify and expand the function of specific family members in development, health, and disease states. However, many of the currently commercially available caspase tools that are widely used are not specific for the targeted caspase. In this report, we delineate the approach we have used to identify, validate, and target caspase-9 in the nervous system using a novel inhibitor and genetic approaches with immunohistochemical read-outs. Specifically, we used the retinal neuronal tissue as a model to identify and validate the presence and function of caspases. This approach enables the interrogation of cell-type specific apoptotic and non-apoptotic caspase-9 functions and can be applied to other complex tissues and caspases of interest. Understanding the functions of caspases can help to expand current knowledge in cell biology, and can also be advantageous to identify potential therapeutic targets due to their involvement in disease.

Presented here is a detailed immunohistochemistry protocol to identify, validate, and target functionally relevant caspases in complex tissues.