We would like to acknowledge the technical assistance of Kristina Jurcic in the University of Western Ontario MALDI MS facility, as well as the National Sciences and Engineering Council (NSERC) for funding this work. The authors would also like to acknowledge the Caprioli group (Vanderbilt University, TN) and Chaurand group (Universit&#233; de Montreal, QC) for their advice in optimizing the sublimation technique presented in this manuscript.

All animal handling procedures described below adhere to the University of Western Ontario&#39;s animal care committee (2016-014).
1. Tissue Preparation and Sectioning
<ol>
	<li>Extract brain tissue from the rat through a process known as &#34;Fresh Frozen Extraction&#34; (FFE).
	<ol>
		<li>Euthanize the rat with an overdose of pentobarbital sodium and monitor reflexes in the limbs. Once all reflexes have ceased, sever the head of the rat using a guillotine.</li>
		<li>Carefully separate the muscles and other tissues from the skull using a scalpel. Use bone-cutting forceps to break the skull to expose the brain, starting from the base of the skull (foramen magnum) to the anterior portion.</li>
		<li>Once the brain is exposed, carefully scoop it out with a surgical spatula and immediately place on crushed dry ice for flash freezing. 
		NOTE: The brain will be very malleable. Therefore, take extreme care when placing the brain on dry ice. If the brain is crushed or pressed against a surface, it will alter its shape and freeze in that position.</li>
	</ol>
	</li>
	<li>Remove fresh frozen tissue (not perfused with formaldehyde or embedded in OCT) from - 80&#176; C freezer and place on dry ice.</li>
	<li>Place several drops of water on a cryostat holder and place on either dry ice or cryostat freeze bar. Press tissue lightly onto cryostat holder as it begins to freeze and hold until water freezes around the base of the tissue and anchors it in place. Add additional water to further secure the tissue on the holder if necessary. 
	NOTE: Water is used instead of OCT media for mounting tissue in order to avoid contamination of the tissue with compounds which can affect the detection of the desired signal.</li>
	<li>Position tissue in cryostat. Section tissue up to the desired anatomical location. Ensure that the cutting thickness is between 8 - 12 &#181;m. 
	NOTE: When sectioning tissue in communal cryostat devices, it is imperative that separate materials such as blades, brushes, anti-roll bars, and holders be used in order to avoid contamination with embedding compounds. The inside of the cryostat should also be cleaned thoroughly with ethanol before use.</li>
	<li>Using the cryostat anti-roll bar, slowly section flattened tissue sections. Holes or markings on the tissue can occur if the roll bar placement is incorrect or blade is dull. Move the tissue to the center of slicing platform using clean paintbrushes.</li>
	<li>Mounting
	<ol>
		<li>Freeze conductive slides or metal plates by placing them in the cryostat while slicing. When the slide is completely frozen, carefully move the sectioned tissue onto the conductive surface of the slide using clean paintbrushes. Once all tissue sections are positioned correctly on the slide, place a finger under the slide, opposite the tissue, and press until the section thaws. 
		NOTE: Sections may fold or curl during the thawing process when using this mounting method. This can be reduced by keeping the slide in the cryostat when thawing the tissue. If these issues become problematic, an alternative, warm-mount method is listed below.</li>
		<li>Alternatively, take a room temperature Indium-tin Oxide (ITO) slide (or metal plate) and lightly press down on frozen tissue section on the slicing platform surface, conductive side down. This will lead to the tissue section thawing evenly onto the surface of the slide with little curling or folding of the tissue. 
		NOTE: Condensation may appear below the section when mounting using this method which may lead to loss of certain proteins and lipids.</li>
	</ol>
	</li>
	<li>Place slides with tissue in a desiccator for 5 - 10 min.</li>
</ol>
2. Sublimator Apparatus Set-up
NOTE: Perform these steps in a fume hood.
<ol>
	<li>Place a sand bath in an aluminum container onto a hot plate, with the hot plate on a metal scissor lift of appropriate surface area (i.e. larger than the hot plate). Turn on the hot plate and set the temperature to 140 &#176;C. 
	NOTE: The melting point for DAN matrix is between 187 - 190 &#176;C. Do not exceed this temperature on the hotplate.
	<ol>
		<li>If the hot plate is equipped with a temperature feedback probe, use it to monitor sand temperature throughout the experiment and ensure temperature consistency. This feature can assist with experiment reproducibility across various sublimation experiments. 
		NOTE: The sand bath should be contained within an aluminum container as a glass container may shatter at high temperatures.</li>
	</ol>
	</li>
	<li>Place 300 mg of DAN matrix onto the bottom surface of sublimation apparatus. Place the matrix in the center of the apparatus and spread out in an even layer in the approximate width and length of the slide being sublimated. 
	CAUTION: DAN matrix is toxic. Therefore, it is important wear gloves, masks, and safety goggles at all times when handling the powdered matrix. DAN should be stored in the dark as it is light sensitive.</li>
	<li>Tape a metal plate onto the inner surface of the apparatus with the plate making direct contact with the bottom of the condenser in order to ensure even distribution of temperature cooling across the entire surface of the slide during sublimation. Alternatively, sand the bottom of the condenser to ensure a flat surface.
	<ol>
		<li>Place the tape along the outer edges of the plate and adhere to the sides of the inner glassware. If the tape is placed under the plate and adheres to the bottom of the inner glass surface, the temperature distribution may not be even and could result in uneven matrix distribution across the surface of the slide.</li>
		<li>Tape a blank (test) slide diagonally across the surface of the metal plate with the tape again placed on the outer edges of the slide.</li>
	</ol>
	</li>
	<li>Connect the top and bottom portions of the apparatus, with a rubber O-ring in the middle to ensure a complete seal.</li>
	<li>Place a metal U-joint around the center of the apparatus and tighten vices until the top and bottom half of the apparatus are sealed tightly together. Place in the metal O-ring above the sand bath.</li>
	<li>Take a handful of crushed ice and place it in the condenser. Fill condenser &#188; to &#189; full with cold water to create ice slush. The ice slush will cool the metal plate on the inside of the apparatus and subsequently the slide adhered to it. Wait at least 5 min for the temperature to reach a steady state.</li>
	<li>Pour 300 mL of ethanol in the cold trap container and place the glassware into the container. Drop 2 - 3 small pieces of dry ice into the ethanol in the bottom of the cold trap container. The cold trap input has a glass tube running to the bottom of the cylinder, while the output does not.</li>
	<li>Connect the vacuum pump to the cold trap output using rubber tubing. Use another piece of rubber tubing to connect the cold trap input to the sublimation apparatus. Ensure that the tubing is tightly secured using metal clamps if available.</li>
	<li>Use the vacuum pump to deliver a vacuum of 30 - 50 mT. Allow the pump to run for at least 5 min for pressure equilibration. Vices on U-ring of sublimation apparatus may have to be tightened again once the vacuum pump is turned on because of decreased pressure in the apparatus. 
	NOTE: It is highly recommended that a vacuum gauge be attached to the pump to monitor the pressure of vacuum during the experiment and to test for leaks in pressure in order to achieve the highest possible reproducibility between experiments. However, most pumps are designed to maintain a constant pressure, therefore the gauge may not be essential for experienced users. Additionally, vacuum seal grease can be used to help maintain vacuum pressure.</li>
</ol>
3. Sublimation
<ol>
	<li>Ensure that the sand bath temperature has stabilized at 140 &#176;C and that the apparatus is secured in the metal O-ring above the sand bath with all tubing connected and the vacuum turned on.</li>
	<li>Set timer for 7 min but do not start timer. Slowly raise the scissor lift and sand bath up to the sublimation apparatus until the U-joint of the sublimator is well above the metal O-ring. This extra space allows for adjustment of the apparatus on the sand.
	<ol>
		<li>Quickly press the sublimator gently on the sand surface to ensure that the apparatus is sitting evenly on the sand, and then immediately start the timer.</li>
	</ol>
	</li>
	<li>When the timer sounds, turn off the vacuum pump and carefully lower the scissor lift until the sublimator is no longer touching the sand bath and is sitting securely in the metal O-ring.
	<ol>
		<li>Slowly loosen the micro-vent valve to release pressure in the apparatus. Loosen the metal clamp around the rubber tubing of the sublimation apparatus and slowly begin to loosen the tube. Once the rubber tube has been loosened slightly, bend the tube to one side to allow residual pressure to escape.</li>
		<li>When ambient pressure returns, carefully remove the rubber tubing from the sublimation apparatus.</li>
	</ol>
	</li>
	<li>Loosen the vices on the U-joint of the apparatus and remove. Carefully separate the two halves of the sublimation apparatus and pull off the slide from the top of the inner glassware. Examine slide to confirm even matrix distribution. 
	NOTE: If matrix is uneven, the powdered matrix in the bottom of the sublimator can be repositioned or the sublimator apparatus can be repositioned in the sand for the next slide. The sublimation process can be repeated several times using the same matrix, however, the matrix will eventually become darker from repeated heat exposure and the quantity of powder will decrease such that the sublimation time will have to be adjusted slightly to compensate. For this reason, it is important to monitor the amount of matrix being sublimated after each experiment to ensure consistency in matrix deposition between slides</li>
	<li>If the distribution and amount of matrix sublimated is sufficient, tape a new slide with tissue onto the inside of the sublimator and repeat Section 3. 
	NOTE: For quality control (QC) purposes, the amount of matrix sublimated can be measured after each experiment by weighing the slide before and after sublimation and dividing the weight of sublimated matrix with the surface area of the slide 2, it should be noted that due to the lack of precision of most scales beyond 4 decimal points and variability between scales, these measurements should be only be used a guide for optimal matrix deposition as opposed to a fully quantifiable means of QC unless a high precision balance is used (see Figure 3).</li>
</ol>
4. Tissue Storage/Rehydration
<ol>
	<li>After sublimation, store slides in a sealed container or small cassette in sealed plastic bag.</li>
	<li>Incubate slides in -20 &#176;C freezer for 2 h&#160;or overnight. 
	NOTE: The goal of the freezing process is to act as a rehydration step for the desorption of tissue materials into the matrix. The freezing process has also been shown to prevent degradation of lipid signals for up to 1 week when stored at -80 &#176;C 12. For this reason, the amount of time the samples are stored in the freezer can be varied to a certain degree to suit the needs of the experimenter without significantly altering signal detection (as observed in our lab). However, we have noticed some discoloration of matrix when samples are frozen for longer than 24 h at -20 &#176;C. Thus, we recommend imaging the sublimated tissue before that time or storing tissue at -80 &#176;C for longer incubation periods.</li>
	<li>Remove slides from freezer (and container) and place in a desiccator for 5 - 10 min.</li>
</ol>
5. Imaging and Analysis
<ol>
	<li>Remove slides from desiccator and apply instrument standards to ensure mass accuracy of MALDI instrument during imaging. The type of standards will vary depending on instrumentation. Standards are generally applied evenly across entire slide, surrounding tissue to be imaged (Figure 3D).</li>
	<li>Insert slide into MALDI instrument and follow manufacturer&#39;s instructions for imaging experiments (instrument methods should be optimized for a 1,000 - 2,000 mass range). Representative result (Figure 4) was acquired in reflectron negative mode with a 70 &#181;m raster and 20 shots/spectrum (acquisition time ~ 2 h).</li>
</ol>

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The authors have nothing to disclose.

This work details a standardized protocol of matrix sublimation on tissue for the detection of negatively charged lipids, such as gangliosides, in MALDI IMS experiments. Sample preparation for MALDI IMS is highly variable and must be customized to suit the unique properties of the analyte of interest. The application of matrix onto the tissue sample surface is a crucial aspect of the sample preparation process with regards to the quality of results in IMS. Particular care should be taken when selecting matrices, ensuring that the matrix is compatible with the analyte of interest and is free of matrix-derived artifacts in the spectrum. Sublimation is a dry matrix deposition technique which provides a high degree of matrix crystal homogeneity with little delocalization of lipids on tissue. DAN matrix in particular shows a high degree of sensitivity for the desorption of a group of negatively charged membrane lipids called gangliosides, as demonstrated in rat brain sections, and is stable under vacuum for extended periods of time, allowing for compatibility with the majority of imaging applications. DAN matrix has the additional benefit of also having a high affinity for positively charged lipid species, thus increasing the versatility of this matrix for MALDI IMS applications 2. It should also be noted that multiple lipid species, including phospholipids, can be detected simultaneously when using this IMS protocol.
A thorough examination of the properties of DAN matrix as compared to 9 other commonly used matrices has been previously published 2. In this article, the authors highlight the increased sensitivity of DAN matrix in negative ion mode as compared to the other matrices examined as well as the capability for high resolution imaging using DAN. DAN matrix was found to have the highest overall performance for negative polarity ions. Interestingly, it performed equally well for positive polarity ions. Other common matrices which have demonstrated high affinity for negative polarity ions include DHA, DHB, and 9-AA. The efficiency of DHA as a matrix is limited by its low stability under vacuum which makes it impractical for most imaging applications 2. 9-AA has the benefit of producing low background noise and demonstrated enrichment of sulfatide signals in the 750 - 950 mass range in negative mode compared to DHB matrix 13. Although DHB has been shown to be very effective in positive mode, it yields lower negative polarity ion signal as compared to DAN matrix 2. DAN matrix also demonstrated increased sensitivity in negative ion mode in the lower mass range (650 - 950) compared to DHB 2. Additionally, it has been reported that DHB can form significant matrix clusters in negative ion mode up to 750 Da 2.
Another method of dry matrix deposition has been described by Puolitaival et al., in which ground matrix is passed through a fine sieve and deposited onto the sample surface. The authors compared this dry method of matrix deposition to a wet matrix manual spray technique and found that they gave comparable signal for phospholipids in the 450 - 1,200 mass range 14. One study used both sublimation and dry-coating with a sieve to examine phospholipid expression, however, the authors did not comment on how the two techniques compared in terms of image resolution or ion signal yield 15.
In order to ensure the widest possible range of applicability, the protocol presented was a basic workflow that will allow both new and more specialized users of MALDI IMS to achieve highly reproducible results for a wide variety of lipid imaging applications. However, sublimation protocols can be modified to meet the unique circumstances of the experiment. For example, the fine matrix crystal deposition provided by sublimation allows for the option to reverse the order of sample preparation by pre-coating slides/plates with matrix prior to mounting the tissue. The matrix can be sublimated onto the conductive surface in advance and stored in a dark environment for later use thus reducing sample preparation time post-sectioning while still maintaining high sensitivity for lipid detection 16,17. Another modification that can be made to this protocol to enhance signal for the detection of lipids is a 2 min wash with ammonium formate 8.
Although sublimation can circumvent many of the drawbacks of most wet deposition techniques, such as smaller, more homogenous matrix crystal deposition 1, decreased delocalization of analytes, and low cost as compared to automated sprayers, some variability in the sample preparation process is unavoidable. In the case of sublimation, there are several factors which may cause minute differences from one experiment to another and can include the following which have been observed in our lab. There can be differences in the position of the sublimation apparatus on the sand. When running several sublimation experiments back-to-back, the sand in the sand bath may begin to shift its position which can affect the dispersion of heat throughout the sand bath. Flattening the sand before each round of sublimation can also change the dispersion of heat as sand that has been displaced may take time to return to a uniform temperature. Sublimation time may have to be adjusted slightly between each round to compensate for shifting sand in the sand bath. Alternatively, use of an oil bath may lead to more homogenous heat dispersion.
Secondly, the amount of matrix at the bottom of the sublimator can vary. DAN matrix has a clumpy grey appearance and has a tendency to form large clusters. These clusters can be manually broken up and arranged in the bottom of the sublimator but cannot be completely eliminated. Large clusters of matrix will take longer to heat and sublimate than smaller pieces which can affect the end product of the sublimation.
The vacuum output on the sublimation apparatus is another critical factor. Most sublimators are made with a single vacuum output onto which tubing is attached to connect it to the vacuum pump during sublimation. Because the pressure is being regulated on that side of the apparatus, there may be a slight unevenness of the matrix deposition, with more being deposited on the side with the vacuum output. This can be somewhat compensated for by shifting either the position of the apparatus in the sand, the position of the matrix in the bottom of the sublimator, or by changing the position of the slide/plate on the condenser. These factors are the main drawbacks of the sublimation technique and for this reason it is crucial to run a test slide through the sublimation process before running experimental sample so that time can be adjusted to compensate for these variables.
Another reported drawback of sublimation is that the detection of higher mass analytes is limited by insufficient analyte extraction and/or mixing with matrix. A rehydration step after sublimation can help pull out these higher mass analytes to circumvent this issue. This is typically achieved using a humidity chamber 18. However, in this protocol, a freezing step is added after sublimation which achieves the same purpose. The freezing and subsequent thawing (in a desiccator) of the samples before insertion in the MALDI instrument causes condensation to form on the surface of the sample which temporarily rehydrates the sample to allow for the extraction of higher mass lipids while preserving the tissue for analysis 12,17.
Overall, sublimation is a highly sensitive and cost effective method of matrix application for the detection of lipids in MALDI IMS experiments. Indeed, our group has noticed significant improvements to IMS results when we shifted from using an air-spray method 3 to a sublimation method 4 for matrix deposition. The present protocol is appropriate for a number of lipid imaging applications but can be modified to suit the needs of the experimenter.

Matrix-assisted Laser Desorption/Ionization (MALDI) Imaging Mass Spectrometry (IMS) is becoming a highly sought after technique for visualization of the spatial distribution of lipids, peptides and proteins across intact sample surfaces. MALDI IMS was previously known as an analytical technique for pre-purified analytes, but in recent years, it has been drawing attention in many other disciplines because of the ability to combine the accuracy of mass spectrometry with high resolution visual/anatomical reference points without the need for any external labelling. As the scientific pool of researchers utilizing this technique continues to grow, there is increased need for standardized, easy-to-follow protocols to assist in the development and optimization of IMS experiments. Gangliosides, a group of membrane lipids highly abundant in the central nervous system, are ideal for MALDI IMS experiments as their location, embedded within the membrane, makes certain species impossible to detect using conventional Immuno-labelling. Additionally, we have shown, using MALDI IMS, that these lipids, which function as modulators of cell signaling, among other things, have unique anatomical distribution patterns in the healthy rodent brain that are altered after brain injury 3,4,5. Gangliosides are located at a higher mass range compared to most lipid species, and are thus most suited to the MALDI imaging platform.
<img alt="Figure 1" src="/files/ftp_upload/55254/55254fig1.jpg" /> 
Figure 1: Workflow of MALDI IMS Experiment. Diagram of the general workflow of a MALDI IMS experiment using sublimation. Tissue frozen at -80 &#176;C is sectioned in a cryostat and 10 &#181;m sections are thaw mounted onto conductive ITO slides. The slide is then placed in a desiccator until sublimation. Slides are inserted into the sublimation apparatus and an even layer of matrix is applied to the tissue sample surface. Samples are frozen overnight in a -20 &#176;C freezer then placed in a desiccator for 10 min. Once standards have been applied, the samples are inserted into the MALDI instrument where a laser is directed across the tissue causing desorbed molecules in the matrix to ionize. The ions travel down a flight tube and separate based on their mass (time-of-flight/TOF) until they reach the detector. The information on the ionic abundance of analytes within a predetermined mass-to-charge (m/z) range is displayed as both a molecular image and mass spectrum. This data can be used to both visualize and quantify the ionic abundance of the analyte of interest within the imaged tissue. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55254/55254fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Sample preparation for MALDI IMS is highly variable as each step of the process must be customized to the analytes of interest. The defining feature of MALDI-based experiments is the use of a matrix coating deposited onto the sample surface prior to analysis. In addition to the role of absorbing and transferring radiation energy from the laser during the ablation process, the matrix also serves to isolate various analytes from the sample, thereby facilitating the analysis of compounds of interest 6,7. Homogenous application of the matrix to the sample surface is the most crucial step in the sample preparation process. Improper matrix deposition can lead to large heterogeneous matrix crystal formations and the development of artifacts, low ion-signal, and poor reproducibility 7.
Due to the affinity of certain matrices to isolate specific analytes, the type of matrix selected for an experiment can significantly alter the outcome. The matrices used for imaging of proteins and peptides often differ from those used for imaging lipids and the process is further complicated by the need for additional procedures such as washing and rehydration steps in order to successfully detect signals from tissue. Although washing steps exist for the enhancement of lipid signals 8, they are not a prerequisite for the detection of most lipid species. When selecting a matrix for a lipid imaging experiment, it is important to consider the polarity of the lipid of interest as this will narrow the range of suitable matrices. For example, gangliosides contain sialic acid residues which give them an overall negative polarity. There are a number of matrices that can effectively desorb and ionize gangliosides from tissue; however, factors such as matrix-derived peaks in the spectrum and stability of the matrix under vacuum must be taken under consideration. 1,5-diaminonapthalene (DAN) matrix is sufficiently stable under instrument vacuum conditions for the majority of imaging applications and has demonstrated a high degree of sensitivity for lipid desorption and can be used for the analysis of lipids in both positive and negative ion modes 2. DAN matrix, when compared to other negative lipid affinity matrices such as dihydroxybenzoic acid (DHB), 9-aminoacridine (9-AA), and 5-chloro-2-mercaptobenzothiazole (CMBT), was able to most efficiently desorb gangliosides from rat brain tissue in negative ion mode (manuscript in preparation).
Selecting the appropriate method of matrix deposition is of equal importance to selecting the matrix itself. Wet matrix deposition methods wherein the solid matrix is dissolved in an organic solvent, and deposited by pneumatic or automated sprayers or spotters, are particularly effective for the desorption of proteins and peptides as the liquid permeates the sample to allow for extraction of compounds and co-crystallization with the matrix. Although these techniques can also be used for lipid applications, analyte delocalization and uneven matrix crystal formations are common occurrences due to the high abundance and solubility of lipids in solvents, particularly in tissue 2,9. Because lipids are readily ionized from tissue, dry matrix deposition techniques, such as sublimation, offer a simple, cost effective alternative to sprayers while circumventing many of the drawback of these techniques. The success of sublimation in MALDI IMS experiments is attributed to features such as microcrystalline matrix morphology which increases the surface area for matrix-analyte binding, increased matrix purity, and homogenous matrix deposition leading to increased reproducibility compared to wet matrix techniques 1,10.
Sublimation involves heating a powdered matrix under vacuum immediately below a cooled sample surface resulting in solid to gas-phase transition of the powdered matrix followed by deposition onto the tissue sample surface. During sublimation, matrix deposition can be controlled by varying factors such as time, temperature and pressure to provide highly reproducible results. A single sublimation experiment can take anywhere from 5 to 20 min depending on the type of matrix selected, which can be re-used several times before disposal. The apparatus can be purchased commercially at a fraction of the price of automated sprayers and is easily taken apart for cleaning and maintenance. The low cost and relative simplicity of this matrix deposition technique make it ideal for researchers beginning or expanding upon lipid imaging applications in MALDI IMS. Although information detailing protocols for sublimation of tissues for IMS have been reported 11, few standardized protocols exist which focus on the basic workflow involved with carrying out a sublimation experiment for imaging high mass lipids in negative ion mode, making it difficult to establish the technique without extensive trial and error. The following is an experimental protocol aiming to fill that gap for the sublimation of DAN matrix onto rat brain sections for high resolution imaging and detection of gangliosides.
<img alt="Figure 2" src="/files/ftp_upload/55254/55254fig2.jpg" /> 
Figure 2: Sublimation Apparatus. Photograph (A) and schematic diagram (B) of the sublimation apparatus. The vacuum pump is connected by rubber tubing to a cold trap filled with 300 mL of ethanol. The cold trap is then connected by rubber tubing to the sublimation apparatus. The apparatus is made up of two separate pieces of glassware that are sealed together with a metal U-joint. The top half of the sublimator contains the condenser which is filled with ice slush. The sample plate is taped onto the bottom of the condenser, inside the sealed glass apparatus. The bottom half of the sublimation apparatus contains the DAN matrix, spread out evenly facing the sample plate. During sublimation, the glass apparatus is placed on a sand bath heated to 140 &#176;C by a hot plate directly below it. The temperature probe helps maintain a stable temperature throughout the sublimation experiment through feedback of the sand bath temperature compared to the preset temperature for the experiment. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55254/55254fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Sublimator</td>
			<td>Chemglass Life Sciences</td>
			<td>CG3038-01</td>
			<td></td>
		</tr>
		<tr>
			<td>1,5- Diaminonapthalene (DAN) matrix</td>
			<td>Sigma-Aldrich</td>
			<td>D21200&#160;</td>
			<td>100 G</td>
		</tr>
		<tr>
			<td>Cryostat</td>
			<td>Thermo-Fisher Scientific</td>
			<td></td>
			<td>CryoStar NX50</td>
		</tr>
		<tr>
			<td>Hot plate with temperature feedback</td>
			<td>Thermo-Fisher Scientific</td>
			<td>HP88857290</td>
			<td>Isotemp ADVD 7x7 HP 100-120v</td>
		</tr>
		<tr>
			<td>Stainless Steel Jack</td>
			<td>Thermo-Fisher Scientific</td>
			<td>2216479</td>
			<td>10x10</td>
		</tr>
		<tr>
			<td>Cold Trap</td>
			<td></td>
			<td></td>
			<td>Custom built on site</td>
		</tr>
		<tr>
			<td>Vacuum Pump</td>
			<td>Franklin Electric</td>
			<td>1102180403</td>
			<td>Savant VP100 Two Stage</td>
		</tr>
		<tr>
			<td>Indium-tin-oxide (ITO) Slides</td>
			<td>Hudson Surface Technology</td>
			<td>PSI 1111000</td>
			<td>type II, 1.1mm/25 each</td>
		</tr>
		<tr>
			<td>MALDI TOF/TOF 5800 Instrument</td>
			<td>AB Sciex</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Desiccator</td>
			<td>Sigma-Aldrich</td>
			<td>D2797</td>
			<td>tabletop desiccator</td>
		</tr>
	</tbody>
</table>

sublimation of dan matrix for the detection and visualization of gangliosides in rat brain tissue for maldi imaging mass spectrometry

Sample preparation is key for optimal detection and visualization of analytes in Matrix-assisted Laser Desorption/Ionization (MALDI) Imaging Mass Spectrometry (IMS) experiments. Determining the appropriate protocol to follow throughout the sample preparation process can be difficult as each step must be optimized to comply with the unique characteristics of the analytes of interest. This process involves not only finding a compatible matrix that can desorb and ionize the molecules of interest efficiently, but also selecting the appropriate matrix deposition technique. For example, a wet matrix deposition technique, which entails dissolving a matrix in solvent, is superior for desorption of most proteins and peptides, whereas dry matrix deposition techniques are particularly effective for ionization of lipids. Sublimation has been reported as a highly efficient method of dry matrix deposition for the detection of lipids in tissue by MALDI IMS due to the homogeneity of matrix crystal deposition and minimal analyte delocalization as compared to many wet deposition methods 1,2. Broadly, it involves placing a sample and powdered matrix in a vacuum-sealed chamber with the samples pressed against a cold surface. The apparatus is then lowered into a heated bath (sand or oil), resulting in sublimation of the powdered matrix onto the cooled tissue sample surface. Here we describe a sublimation protocol using 1,5-diaminonaphthalene (DAN) matrix for the detection and visualization of gangliosides in the rat brain using MALDI IMS.

Upon completion of the sublimation experiment, the two halves of the glass apparatus are separated in the fume hood and the slide, taped to the condenser, can be removed (Figure 3A). At this point, the slide should be examined for uneven matrix distribution and the time, temperature, or placement of the apparatus in the sand bath may have to be adjusted for the next slide. A successful DAN sublimation experiment will result in an even coating of grey/brown matrix along the surface of the slide where the anatomical features of the tissue can be clearly visualized and the amount of matrix on the glass slide is similar to the amount on the tissue (Figure 3B). For example, if too much matrix has been sublimated on the tissue, the thickness of the matrix will cover the features of the tissue and only the general shape will be discernable. However, if too little matrix is sublimated, the tissue will have a darker appearance than the rest of the slide (Figure 3C). Both too much and too little matrix will result in poor signal in the MALDI instrument. For quality control purposes, Thomas et al. weighed the amount of matrix deposited on the slide and divided the weight by the surface area of the slide. They reported an optimal amount of matrix deposition for several matrices including DAN (110 &#181;g/cm&#178;) 2. Although most balances lack the precision needed to replicate such results, we have attempted this method of QC; however, due to a high degree of variability, we can only advise a suitable range of matrix deposition found to be associated successful versus unsuccessful sublimation experiments with DAN matrix as determined through signal detection in the instrument. A matrix measurement of below 100 &#181;g/cm&#178; was associated with &#34;too little&#34; matrix conditions while a measurement of above 140 &#181;g/cm&#178; was associated with &#34;too much&#34;. Matrix deposition between 100 and 140 &#181;g/cm&#178; was found to be an optimal quantity for imaging with DAN matrix. Calibrating standards should be applied on the glass slide around the tissue samples to ensure the mass accuracy of the detected IMS signals (Figure 3D)
In a MALDI IMS experiment, the molecular image allows for the visualization of the spatial distribution of the analyte of interest along with any unknown species that may be present in a given mass range. The molecular images of gangliosides in this experiment were overlaid using ImageJ software to show the anatomical distribution of several ganglioside species across the cortical and subcortical regions of the rat brain (Figure 4A). The most abundant gangliosides in the brain are the A-series species GD1a and GM1 but also contain gangliosides GM2 and GM3 which can all be found between the 1,000-2,000 m/z mass range, a higher mass range than most common brain lipids, making these gangliosides easy to identify along the spectrum (Figure 4B). The ionic abundance of each ganglioside species can be used to semi-quantify differences or changes in ganglioside species within the same image. Because a certain amount of variability in sample prep cannot be avoided in IMS, between scan comparisons are considered to be semi-quantitative.
<img alt="Figure 3" src="/files/ftp_upload/55254/55254fig3.jpg" /> 
Figure 3. DAN Sublimation. Representative results of sublimation of DAN matrix on rat brain tissue. (A) Upon completion of sublimation, the two halves of the apparatus are separated and the slide is removed from the condenser. (B) DAN matrix spreads evenly across multiple rat brain tissue sections on the surface of the slide to allow for highly reproducible results (between 100 - 150 &#181;g/cm&#178;). (C) Examples of failed sublimation experiments where too little matrix (left - &#60;90 &#181;g/cm&#178;) or too much matrix (right - &#62;150 &#181;g/cm&#178;) was deposited. Too little matrix will result in insufficient desorption of analytes, while too much matrix will not allow the laser to penetrate to the tissue during acquisition, resulting in low ion detection. (D) Slide containing rat brain tissue sections sublimated with DAN matrix and calibration standards applied is ready for insertion into MALDI instrument for imaging. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55254/55254fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/55254/55254fig4.jpg" /> 
Figure 4. MALDI IMS of Gangliosides in the Rat Brain. Representative MALDI IMS molecular image and mass spectrum of gangliosides in the rat brain after sublimation with DAN matrix. The molecular image is a composite of multiple ganglioside species in the spectrum overlaid and pseudocolored using Image J software to show the unique anatomical distribution of ganglioside species throughout the brain. Species GM1d18:1 (red, mass ~1,547 Da), GM1d20:1 (green, mass ~1,573 Da), GM3d18:1 (blue, mass ~1,178 Da), and GM3d20:1 (teal, mass ~1,207 Da) are represented. The mass spectrum displays ionic abundance of analytes within a 1,000 - 2,000 mass range. The major A-series gangliosides are labelled along the spectrum. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55254/55254fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Sublimation of DAN Matrix for the Detection and Visualization of Gangliosides in Rat Brain Tissue for MALDI Imaging Mass Spectrometry at JoVE.com

Sublimation of DAN Matrix for the Detection and Visualization of Gangliosides in Rat Brain Tissue for MALDI Imaging Mass Spectrometry

A protocol for the sublimation of DAN matrix onto rat brain tissue for the detection of gangliosides using MALDI Imaging Mass Spectrometry is presented.

Sample preparation is key for optimal detection and visualization of analytes in Matrix-assisted Laser Desorption/Ionization (MALDI) Imaging Mass ...

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Nanyang Technological University

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JoVE Journal

Neuroscience

10.7K Views.  The University of Western Ontario. A protocol for the sublimation of DAN matrix onto rat brain tissue for the detection of gangliosides using MALDI Imaging Mass Spectrometry is presented.

Video: Sublimation of DAN Matrix for the Detection and Visualization of Gangliosides in Rat Brain Tissue for MALDI Imaging Mass Spectrometry

Visualization of Proprioceptors in Drosophila Larvae and Pupae

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other insects proprioception is provided by specialized sensory organs termed chordotonal organs (ChOs) 2. Like many other organs in Drosophila, ChOs develop twice during the life cycle of the fly. First, the larval ChOs develop during embryogenesis. Then, the adult ChOs start to develop in the larval imaginal discs and continue to differentiate during metamorphosis.
The development of larval ChOs during embryogenesis has been studied extensively 10,11,13,15,16. The centerpiece of each ChO is a sensory unit composed of a neuron and a scolopale cell. The sensory unit is stretched between two types of accessory cells that attach to the cuticle via specialized epidermal attachment cells 1,9,14. When a fly larva moves, the relative displacement of the epidermal attachment cells leads to stretching of the sensory unit and consequent opening of specific transient receptor potential vanilloid (TRPV) channels at the outer segment of the dendrite 8,12. The elicited signal is then transferred to the locomotor central pattern generator circuit in the central nervous system.
Multiple ChOs have been described in the adult fly 7. These are located near the joints of the adult fly appendages (legs, wings and halters) and in the thorax and abdomen. In addition, several hundreds of ChOs collectively form the Johnston's organ in the adult antenna that transduce acoustic to mechanical energy 3,5,17,4.
In contrast to the extensive knowledge about the development of ChOs in embryonic stages, very little is known about the morphology of these organs during larval stages. Moreover, with the exception of femoral ChOs 18 and Johnston's organ, our knowledge about the development and structure of ChOs in the adult fly is very fragmentary.
Here we describe a method for staining and visualizing ChOs in third instar larvae and pupae. This method can be applied together with genetic tools to better characterize the morphology and understand the development of the various ChOs in the fly.

Visualization of Proprioceptors in Drosophila Larvae and Pupae

A method to immunostain and visualize chordotonal organs in larvae and pupae of Drosophila melanogaster is described.

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other ...

Stereotaxic Surgery for Excitotoxic Lesion of Specific Brain Areas in the Adult Rat

Many behavioral functions in mammals, including rodents and humans, are mediated principally by discrete brain regions. A common method for discerning the function of various brain regions for behavior or other experimental outcomes is to implement a localized ablation of function. In humans, patient populations with localized brain lesions are often studied for deficits, in hopes of revealing the underlying function of the damaged area. In rodents, one can experimentally induce lesions of specific brain regions.
Lesion can be accomplished in several ways. Electrolytic lesions can cause localized damage but will damage a variety of cell types as well as traversing fibers from other brain regions that happen to be near the lesion site. Inducible genetic techniques using cell-type specific promoters may also enable site-specific targeting. These techniques are complex and not always practical depending on the target brain area. Excitotoxic lesion using stereotaxic surgery, by contrast, is one of the most reliable and practical methods of lesioning excitatory neurons without damaging local glial cells or traversing fibers.
Here, we present a protocol for stereotaxic infusion of the excitotoxin, N-methyl-D-aspartate (NMDA), into the basolateral amygdala complex. Using anatomical indications, we apply stereotaxic coordinates to determine the location of our target brain region and lower an injection needle in place just above the target. We then infuse our excitotoxin into the brain, resulting in excitotoxic death of nearby neurons. While our experimental subject of choice is a rat, the same methods can be applied to other mammals, with the appropriate adjustments in equipment and coordinates.
This method can be used on a variety of brain regions, including the basolateral amygdala1-6, other amygdala nuclei6, 7, hippocampus8, entorhinal cortex9 and prefrontal cortex10. It can also be used to infuse biological compounds such as viral vectors1, 11. The basic stereotaxic technique could also be adapted for implantation of more permanent osmotic pumps, allowing more prolonged exposure to a compound of interest.

Targeted ablation of specific brain region(s) by infusion of an excitotoxin using stereotaxic coordinates is described. This technique could also be adapted for infusion of other chemicals into the rat brain.

Many behavioral functions in mammals, including rodents and humans, are mediated principally by discrete brain regions. A common method for discerning the ...

Fiber-optic Implantation for Chronic Optogenetic Stimulation of Brain Tissue

Elucidating patterns of neuronal connectivity has been a challenge for both clinical and basic neuroscience. Electrophysiology has been the gold standard for analyzing patterns of synaptic connectivity, but paired electrophysiological recordings can be both cumbersome and experimentally limiting. The development of optogenetics has introduced an elegant method to stimulate neurons and circuits, both in vitro1 and in vivo2,3. By exploiting cell-type specific promoter activity to drive opsin expression in discrete neuronal populations, one can precisely stimulate genetically defined neuronal subtypes in distinct circuits4-6. Well described methods to stimulate neurons, including electrical stimulation and/or pharmacological manipulations, are often cell-type indiscriminate, invasive, and can damage surrounding tissues. These limitations could alter normal synaptic function and/or circuit behavior. In addition, due to the nature of the manipulation, the current methods are often acute and terminal. Optogenetics affords the ability to stimulate neurons in a relatively innocuous manner, and in genetically targeted neurons. The majority of studies involving in vivo optogenetics currently use a optical fiber guided through an implanted cannula6,7; however, limitations of this method include damaged brain tissue with repeated insertion of an optical fiber, and potential breakage of the fiber inside the cannula. Given the burgeoning field of optogenetics, a more reliable method of chronic stimulation is necessary to facilitate long-term studies with minimal collateral tissue damage. Here we provide our modified protocol as a video article to complement the method effectively and elegantly described in Sparta et al.8 for the fabrication of a fiber optic implant and its permanent fixation onto the cranium of anesthetized mice, as well as the assembly of the fiber optic coupler connecting the implant to a light source. The implant, connected with optical fibers to a solid-state laser, allows for an efficient method to chronically photostimulate functional neuronal circuitry with less tissue damage9 using small, detachable, tethers. Permanent fixation of the fiber optic implants provides consistent, long-term in vivo optogenetic studies of neuronal circuits in awake, behaving mice10 with minimal tissue damage.

The development of optogenetics now provides the means to precisely stimulate genetically defined neurons and circuits, both in vitro and in vivo. Here we describe the assembly and implantation of a fiber optic for chronic photostimulation of brain tissue.

Elucidating patterns of neuronal connectivity has been a challenge for both clinical and basic neuroscience. Electrophysiology has been the gold standard ...

Immunohistochemical Analysis in the Rat Central Nervous System and Peripheral Lymph Node Tissue Sections

Immunohistochemistry (IHC) provides highly specific, reliable and attractive protein visualization. Correct performance and interpretation of an IHC-based multicolor labeling is challenging, especially when utilized for assessing interrelations between target proteins in the tissue with a high fat content such as the central nervous system (CNS).
Our protocol represents a refinement of the standard immunolabeling technique particularly adjusted for detection of both structural and soluble proteins in the rat CNS and peripheral lymph nodes (LN) affected by neuroinflammation. Nonetheless, with or without further modifications, our protocol could likely be used for detection of other related protein targets, even in other organs and species than here presented.

We here present an optimized, detailed protocol for double immunostaining in formalin-fixed, paraffin-embedded rat central nervous system (CNS) and peripheral lymph node (LN) tissue sections.

Immunohistochemistry (IHC) provides highly specific, reliable and attractive protein visualization. Correct performance and interpretation of an IHC-based ...

A Dual Tracer PET-MRI Protocol for the Quantitative Measure of Regional Brain Energy Substrates Uptake in the Rat

We present a method for comparing the uptake of the brain&#39;s two key energy substrates: glucose and ketones (acetoacetate [AcAc] in this case) in the rat. The developed method is a small-animal positron emission tomography (PET) protocol, in which&#160;11C-AcAc and 18F-fluorodeoxyglucose (18F-FDG) are injected sequentially in each animal. This dual tracer PET acquisition is possible because of the short half-life of 11C (20.4 min). The rats also undergo a magnetic resonance imaging (MRI) acquisition seven days before the PET protocol. Prior to image analysis, PET and MRI images are coregistered to allow the measurement of regional cerebral uptake (cortex, hippocampus, striatum, and cerebellum). A quantitative measure of 11C-AcAc and 18F-FDG brain uptake (cerebral metabolic rate; &#956;mol/100 g/min) is determined by kinetic modeling using the image-derived input function (IDIF) method. Our new dual tracer PET protocol is robust and flexible; the two tracers used can be replaced by different radiotracers to evaluate other processes in the brain. Moreover, our protocol is applicable to the study of brain fuel supply in multiple conditions such as normal aging and neurodegenerative pathologies such as Alzheimer&#39;s and Parkinson&#39;s diseases.

Small-animal positron emission tomography enables the assessment of the brain&#39;s two main energy substrates:&#160;glucose and ketones. In the present method, 11C-acetoacetate and 18F-fluorodeoxyglucose are injected sequentially in each animal, and their uptake is measured quantitatively in specific brain regions determined from the magnetic resonance images.

We present a method for comparing the uptake of the brain&#39;s two key energy substrates: glucose and ketones (acetoacetate [AcAc] in this case) in the ...

Live Imaging of Mitosis in the Developing Mouse Embryonic Cortex

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the developing cerebral cortex, abnormal mitosis of neural progenitors can cause defects in brain size and function. Hence, there is a critical need for tools to understand the mechanisms of neural progenitor mitosis. Cortical development in rodents is an outstanding model for studying this process. Neural progenitor mitosis is commonly examined in fixed brain sections. This protocol will describe in detail an approach for live imaging of mitosis in ex vivo embryonic brain slices. We will describe the critical steps for this procedure, which include: brain extraction, brain embedding, vibratome sectioning of brain slices, staining and culturing of slices, and time-lapse imaging. We will then demonstrate and describe in detail how to perform post-acquisition analysis of mitosis. We include representative results from this assay using the vital dye Syto11, transgenic mice (histone H2B-EGFP and centrin-EGFP), and in utero electroporation (mCherry-&#945;-tubulin). We will discuss how this procedure can be best optimized and how it can be modified for study of genetic regulation of mitosis. Live imaging of mitosis in brain slices is a flexible approach to assess the impact of age, anatomy, and genetic perturbation in a controlled environment, and to generate a large amount of data with high temporal and spatial resolution. Hence this protocol will complement existing tools for analysis of neural progenitor mitosis.

Neural progenitor mitosis is a critical parameter of neurogenesis. Much of our understanding of neural progenitor mitosis is based on analysis of fixed tissue. Live imaging in embryonic brain slices is a versatile technique to assess mitosis with high temporal and spatial resolution in a controlled environment.

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the ...

Non-invasive Parenchymal, Vascular and Metabolic High-frequency Ultrasound and Photoacoustic Rat Deep Brain Imaging

Photoacoustics and high frequency ultrasound stands out as powerful tools for neurobiological applications enabling high-resolution imaging on the central nervous system of small animals. However, transdermal and transcranial neuroimaging is frequently affected by low sensitivity, image aberrations and loss of space resolution, requiring scalp or even skull removal before imaging. To overcome this challenge, a new protocol is presented to gain significant insights in brain hemodynamics by photoacoustic and high-frequency ultrasounds imaging with the animal skin and skull intact. The procedure relies on the passage of ultrasound (US) waves and laser directly through the fissures that are naturally present on the animal cranium. By juxtaposing the imaging transducer device exactly in correspondence to these selected areas where the skull has a reduced thickness or is totally absent, one can acquire high quality deep images and explore internal brain regions that are usually difficult to anatomically or functionally describe without an invasive approach. By applying this experimental procedure, significant data can be collected in both sonic and optoacoustic modalities, enabling to image the parenchymal and the vascular anatomy far below the head surface. Deep brain features such as parenchymal convolutions and fissures separating the lobes were clearly visible. Moreover, the configuration of large and small blood vessels was imaged at several millimeters of depth, and precise information were collected about blood fluxes, vascular stream velocities and the hemoglobin chemical state. This repertoire of data could be crucial in several research contests, ranging from brain vascular disease studies to experimental techniques involving the systemic administration of exogenous chemicals or other objects endowed with imaging contrast enhancement properties. In conclusion, thanks to the presented protocol, the US and PA techniques become an attractive noninvasive performance-competitive means for cortical and internal brain imaging, retaining a significant potential in many neurologic fields.

The present work describes a new protocol to perform non-invasive high-frequency ultrasound and photoacoustic based imaging on rat brain, to efficiently visualize deep subcortical regions and their vascular patterns by directing signals on skull foramina naturally present on animal cranium.

Photoacoustics and high frequency ultrasound stands out as powerful tools for neurobiological applications enabling high-resolution imaging on the central ...

Visualization of Amyloid &#946; Deposits in the Human Brain with Matrix-assisted Laser Desorption/Ionization Imaging Mass Spectrometry

The neuropathology of Alzheimer&#39;s disease (AD) is characterized by the accumulation and aggregation of amyloid &#946; (A&#946;) peptides into extracellular plaques of the brain. The A&#946; peptides, composed of 40 amino acids, are generated from amyloid precursor proteins (APP) by &#946;- and &#947;-secretases. A&#946; is deposited not only in cerebral parenchyma but also in leptomeningeal and cerebral vessel walls, known as cerebral amyloid angiopathy (CAA). While a variety of A&#946; peptides were identified, the detailed production and distribution of individual A&#946; peptides in pathological tissues of AD and CAA have not been fully addressed. Here, we develop a protocol of matrix-assisted laser desorption/ionization-based imaging mass spectrometry (MALDI-IMS) on human autopsy brain tissues to obtain comprehensive protein mapping. For this purpose, human cortical specimens were obtained from the Brain Bank at the Tokyo Metropolitan Institute of Gerontology. Frozen cryosections are cut and transferred to indium-tin-oxide (ITO)-coated glass slides. Spectra are acquired using the MALDI system with a spatial resolution up to 20 &#181;m. Sinapinic acid (SA) is uniformly deposited on the slide using either an automatic or a manual sprayer. With the current technical advantages of MALDI-IMS, a typical data set of various A&#946; species within the same sections of human autopsied brains can be obtained without specific probes. Furthermore, high-resolution (20 &#181;m) imaging of an AD brain and severe CAA sample clearly shows that A&#946;1-36 to A&#946;1-41 were deposited into leptomeningeal vessels, while A&#946;1-42 and A&#946;1-43 were deposited in cerebral parenchyma as senile plaque (SP). It is feasible to adopt MALDI-IMS as a standard approach in combination with clinical, genetic, and pathological observations in understanding the pathology of AD, CAA, and other neurological diseases based on the current strategy.

Molecular imaging with matrix-assisted laser desorption/ionization-based imaging mass spectrometry (MALDI-IMS) allows the simultaneous mapping of multiple analytes in biological samples. Here, we present a protocol for the detection and visualization of amyloid &#946; protein on brain tissues of Alzheimer&#39;s disease and cerebral amyloid angiopathy samples using MALDI-IMS.

The neuropathology of Alzheimer&#39;s disease (AD) is characterized by the accumulation and aggregation of amyloid &#946; (A&#946;) peptides into ...

Light-Induced Molecular Adsorption of Proteins Using the PRIMO System for Micro-Patterning to Study Cell Responses to Extracellular Matrix Proteins

Cells sense a variety of extracellular cues, including the composition and geometry of the extracellular matrix, which is synthesized and remodeled by the cells themselves. Here, we present the method of Light-Induced Molecular Adsorption of Proteins (LIMAP) using the PRIMO system as a patterning technique to produce micro-patterned extracellular matrix (ECM) substrates using a single or combination of proteins. The method enables printing of ECM patterns in micron resolution with excellent reproducibility. We provide a step-by-step protocol and demonstrate how this can be applied to study the processes of neuronal pathfinding. LIMAP has significant advantages over existing micro-printing methods in terms of the ease of patterning more than one component and the ability to generate a pattern with any geometry or gradient. The protocol can easily be adapted to study the contribution of almost any chemical component towards cell fate and cell behavior. Finally, we discuss common issues that can arise and how these can be avoided.

Our overall aim is to understand how cells sense extracellular cues that lead to directed axonal growth. Here, we describe the methodology of Light-Induced Molecular Adsorption of Proteins, used to produce defined micro-patterns of extracellular matrix components in order to study specific events that govern axon outgrowth and pathfinding.

Cells sense a variety of extracellular cues, including the composition and geometry of the extracellular matrix, which is synthesized and remodeled by the ...

Labeling and Imaging of Amyloid Plaques in Brain Tissue Using the Natural Polyphenol Curcumin

Deposition of amyloid beta protein (A&#946;) in extra- and intracellular spaces is one of the hallmark pathologies of Alzheimer&#39;s disease (AD). Therefore, detection of the presence of A&#946; in AD brain tissue is a valuable tool for developing new treatments to prevent the progression of AD. Several classical amyloid binding dyes, fluorochrome, imaging probes, and A&#946;-specific antibodies have been used to detect A&#946; histochemically in AD brain tissue. Use of these compounds for A&#946; detection is costly and time consuming. However, because of its intense fluorescent activity, high-affinity, and specificity for A&#946;, as well as structural similarities with traditional amyloid binding dyes, curcumin (Cur) is a promising candidate for labeling and imaging of A&#946; plaques in postmortem brain tissue. It is a natural polyphenol from the herb Curcuma longa. In the present study, Cur was used to histochemically label A&#946; plaques from both a genetic mouse model of 5x familial Alzheimer&#39;s disease (5xFAD) and from human AD tissue within a minute. The labeling capability of Cur was compared to conventional amyloid binding dyes, such as thioflavin-S (Thio-S), Congo red (CR), and Fluoro-jade C (FJC), as well as A&#946;-specific antibodies (6E10 and A11). We observed that Cur is the most inexpensive and quickest way to label and image A&#946; plaques when compared to these conventional dyes and is comparable to A&#946;-specific antibodies. In addition, Cur binds with most A&#946; species, such as oligomers and fibrils. Therefore, Cur could be used as the most cost-effective, simple, and quick fluorochrome detection agent for A&#946; plaques.

Curcumin is an ideal fluorophore for labeling and imaging of amyloid beta protein plaques in brain tissue due to its preferential binding to amyloid beta protein as well as its structural similarities with other traditional amyloid binding dyes. It can be used to label and image amyloid beta protein plaques more efficiently and inexpensively than traditional methods.

Deposition of amyloid beta protein (A&#946;) in extra- and intracellular spaces is one of the hallmark pathologies of Alzheimer&#39;s disease (AD). ...