Yuki X. Chen, Kelly Veerasammy, and Mayan Hein are supported by the Sloan Foundation CUNY Summer Research Program (CSURP). Jun Yin is supported by the intramural research program of the National Institutes of Health Project Number 1ZIANS003137. Support for this project was provided by a PSC-CUNY Award to Ye He and Rinat Abzalimov, jointly funded by The Professional Staff Congress and The City University of New York.

1. Fly head embedding
NOTE: The whole procedure takes ~45-60 min.
<ol>
	<li>Prepare the optimal cutting temperature compound (OCT compound) stage with a flat surface.
	<ol>
		<li>Add OCT into a plastic cryomold (15 mm x 15 mm x 5 mm) to half of the depth of the cryomold and avoid bubble formation. Leave the mold on a flat surface for several minutes, and then transfer it onto dry ice.</li>
		<li>Keep the cryomold flat on the dry ice and allow the OCT to form a flat and even surface. Wait until the OCT is fully solidified in the mold. Use the frozen OCT stages immediately or store them at &#8722;80 &#176;C.</li>
	</ol>
	</li>
	<li>Anesthetize adult flies using CO2 (i.e., a CO2 pad).
	<ol>
		<li>Prepare a Petri dish containing a piece of laboratory wipe. Use water to moisten part of the wipe to reduce the static electricity. Keep the wipe half wet and half dry.</li>
		<li>Under dissecting scope 1, use forceps to cut the fly head. Collect 4-5 heads each time and put them onto the dry area of the laboratory wipe. 
		NOTE: The collection of 4-5 heads takes ~ 2 mins.</li>
	</ol>
	</li>
	<li>Transfer the OCT stage from dry ice to the dissection scope.
	<ol>
		<li>Take the OCT stage from dry ice to microscope 2, immediately transfer the heads to the OCT stage, and arrange them quickly, which takes ~30 s to avoid OCT melting. Leave ~1 mm of blank space around each fly brain to ensure adequate support from the OCT and 4-5 mm of blank space from the edge of the block to provide adequate room to handle the section. If the proboscis is too long, remove the tip; if it is not too long, keep it intact. Put the OCT stage back onto the dry ice and let it stay on for ~3 min to make sure the OCT stage remains frozen and solid.</li>
		<li>When the OCT stage is back on the dry ice and waiting for solidification, collect another round of heads. Repeat steps 1.2 and 1.3 to transfer additional samples onto the remaining space on the stage. 
		NOTE: Eight heads are usually prepared for each genotype, and four genotypes are used in one OCT stage in this laboratory.</li>
		<li>Use two dissection microscopes, side by side, to avoid changing focus and increasing the time taken to transfer and arrange the heads on the OCT stage and to lower the risk of the OCT stage melting.</li>
	</ol>
	</li>
	<li>After all the fly heads are aligned, let the OCT stage sit on the dry ice for another 5-10 min.</li>
	<li>Take the OCT stage away from the dry ice, put it on a flat surface (bench), and then, quickly, add a large amount of OCT compound to cover all the samples and fill the whole cryomold, which takes ~3 s.</li>
	<li>Immediately transfer the cryomold back to dry ice and freeze the whole OCT block containing the embedded tissues. Let the OCT stage sit on the dry ice for another 5-10 min. Label the samples on the margin of the cryomold.</li>
	<li>Store the frozen samples at &#8722;80 &#176;C until ready for sectioning.</li>
</ol>
2. Cryosectioning the tissue
NOTE: When handling the indium tin oxide (ITO) slides, wear gloves at all times to avoid tissue contamination. Wearing a mask is also recommended to avoid breathing directly onto the slide.
<ol>
	<li>Confirm the ITO-coated side by testing the conductivity of the ITO slides using a voltmeter set to resistance. Mark the side with a resistance measurement as the side to adhere the tissue to. Label it and always set a laboratory wipe on the bottom of the slide to avoid slide contamination.</li>
	<li>Allow the tissues to equilibrate in the cryostat chamber for 30-45 min.
	<ol>
		<li>To avoid the melting of the OCT, place all the necessary tools such as forceps and a thin-tipped artist brush in the cryostat chamber ahead of time to precool them.</li>
	</ol>
	</li>
	<li>Clean the cryostat, preferably with 70% ethanol. Wipe the roll plate and stage and remove used blades. Use additional clean wipes to ensure that the ethanol has evaporated and that all the surfaces are dry before sectioning begins.</li>
	<li>Adjust the temperature of the cryostat chamber and specimen head according to the type of the tissue (e.g., &#8722;14 &#176;C for liver, &#8722;20 &#176;C for muscle, and &#8722;25 &#176;C for skin10; &#8722;18 &#176;C for fly heads in this protocol).</li>
	<li>Mount the tissue onto the specimen holder using OCT. Be careful to use enough OCT to cover the base of the OCT block and mount the block as flat as possible.</li>
	<li>Place a clean blade in the stage and lock it. Position the head of the specimen toward the stage as needed to achieve the desired cutting angle.
	<ol>
		<li>Place the specimen block in an orientation where all genotypes/treatment groups are positioned vertically to the blade. 
		NOTE: This ensures a consistent cutting plane and avoids cross-contamination from different groups. If cutting a different block of tissue, switch to a new blade between samples to prevent cross-contamination.</li>
	</ol>
	</li>
	<li>Begin cutting in thick sections (50-100 &#181;m) until the region of interest (e.g., the desired region of the brain) is found.
	<ol>
		<li>Constantly brush off extra pieces with a precooled artist brush to keep the stage clean.</li>
	</ol>
	</li>
	<li>Change the thickness of the sections to 10-12 &#181;m once the desired region is reached.
	<ol>
		<li>Adjust the chamber temperature slightly as necessary. For example, set a higher temperature if the section tends to flake or fall apart easily. 
		NOTE: The recommended temperature for OCT blocks is &#8722;18 &#176;C.</li>
	</ol>
	</li>
	<li>Carefully collect the desired section and adhere it to the ITO slide. Perform this operation in the cryostat chamber.
	<ol>
		<li>Take a room-temperature ITO slide and position it over the section, approach&#160;the section gently and quickly for the section to adhere to the slide without leaving traces on the cryostat stage. 
		NOTE: The OCT will melt and cling to the slide.</li>
	</ol>
	</li>
	<li>Place the slide aside in a rack or laboratory wipe outside the cryostat between the collections of multiple sections.</li>
	<li>If comparison across different samples of the same cohort is desired, place the sections from multiple samples onto a single slide for simultaneous analysis and minimal&#160;variation. If necessary, separate to two slides, as the MALDI target holder can accommodate two slides in a single run.</li>
	<li>Transport the slides in a vacuum box to a desiccator with desiccant as the bottom layer. Dry the slides under a vacuum for 30-60 min. 
	NOTE: Alternatively, if the lab is not equipped with a vacuum desiccator, the slides should be kept at &#8722;20 &#176;C throughout the process until storage at &#8722;80 &#176;C within 24 h or shipping on dry ice to avoid the deterioration of lipids or metabolites.</li>
	<li>Proceed to matrix deposition. Use 2,5-dihydroxybenzoic acid (DHB) in methanol/water (70/30, v/v) as the matrix.</li>
	<li>If slides are not run immediately, either store the slides at &#8722;80 &#176;C (up to 1 month for fly brain sections and 6 months for rodent brain sections), or immediately ship the sample to MALDI core facilities with adequate dry ice. For optimal storage and shipping, place the slides into a slide transporter and securely seal the opening with wax film. Seal with a zip bag, place it into another zip bag containing desiccant, and label the outside. 
	NOTE: Refer to previous work for the followed steps of slide scanning, matrix deposition, and MALDI imaging procedures11.</li>
	<li>Perform matrix deposition using the automatic HTX M5 matrix sprayer and a 40 mg/mL solution of DHB in methanol/water (70/30, v/v). Spray the matrix at a customized flow rate of 0.12 mL/min and a nozzle temperature of 85 &#176;C for 10 passes. Use an N2 gas pressure of 10 psi. 
	NOTE: If the N2 gas pressure in the sprayer is lowered below 5 psi, a safety mechanism in the sprayer will shut the heater off at once to avoid any damage to the sprayer with a 1,300 mm/min spray velocity, 2 mm track spacing, 3 L/min flow rate, and 40 mm nozzle height.</li>
	<li>Use a MALDI time of flight (TOF) MS instrument (see the Table of Materials) in positive ion mode to acquire a mass spectrum within the mass ranges from m/z 50-1,000.
	<ol>
		<li>To calibrate the instrument, spot 0.5 &#181;L of red phosphorus emulsion in acetonitrile onto the ITO slides and use its spectra to calibrate the instrument in the 50-1,000 m/z mass range by applying a quadratic calibration curve2.</li>
		<li>Set the laser spot diameters to Medium, as it is the modulated beam profile for 40 &#181;m raster width, and gather imaging data by summing 500 shots at a laser repetition rate of 1,000 Hz per array position.</li>
		<li>Use software (see the Table of Materials) to record and process the spectral data. Perform imaging data analysis using root mean square (RMS) normalization to generate ion images at a bin width of &#177;0.10 Da. Align the spectra of both the OCT and brain tissue from the same experiment using the software to evaluate the overlapping peaks and the ion suppression interference of the OCT (Supplemental Figure S1). After the experiment, process the MALDI slides containing the tissue samples by hematoxylin and eosin (H&#38;E) staining, as previously described11.</li>
	</ol>
	</li>
</ol>

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The authors have no conflicts of interest to disclose.

As demonstrated in the study on the variations in lipid composition in mutant and wild-type Drosophila brains, MALDI MSI can be a valuable label-free imaging technique for in situ analysis of molecular distribution patterns within organs of small insects. Indeed, because lipids are distributed in both the brain tissue and fat bodies of Drosophila heads, conventional lipidomics approaches based on liquid-chromatography and mass-spectrometry (LC-MS) can detect only combined signals from both regi...

Lipids are involved in a wide range of biological processes and can be broadly classified into five categories based on their structural diversity: fatty acids, triacylglycerols (TAGs), phospholipids, sterol lipids, and sphingolipids1. The fundamental functions of lipids are to provide energy sources for biological processes (i.e., TAGs) and form cellular membranes (i.e., phospholipids and cholesterol).&#160;However, additional roles of lipids have been noted in development and diseases, and have been extensively studied in the biomedical field. For instance, reports have shown that fatty acids of different lengths may have unique therapeutic roles. Short fatty acid chains can be involved in defense mechanisms against autoimmune diseases, medium-length fatty acid chains produce metabolites that can mitigate seizures, and long fatty acid chains generate metabolites that can be used to treat metabolic disorders2. In the nervous system, glia-derived cholesterol and phospholipids have been shown to be vital for synaptogenesis3,4. Other types of lipids have shown promise in medical applications, including&#160;sphingolipids utilized in drug delivery systems and saccharolipids&#160;used to support the immune system5,6. The numerous roles and potential therapeutic applications of lipids in the biomedical field have made lipidomics&#8212;the study of the pathways and interactions of cellular lipids&#8212;a critical and increasingly important field.
Lipidomics makes use of analytical chemistry to study the lipidome on a large scale. The main experimental methods utilized in lipidomics are based on mass spectrometry (MS) coupled with various chromatography and ion-mobility techniques7,8. The use of MS in the area is advantageous due to its high specificity and sensitivity, speed of acquisition, and unique capabilities to (1) detect lipids and lipid metabolites occurring even at low and transient levels, (2) detect hundreds of different lipid compounds in a single experiment, (3) identify previously unknown lipids, and (4) distinguish between lipid isomers. Among the developments in MS, including desorption electrospray ionization (DESI), MALDI, and secondary ion mass spectrometry (SIMS), MALDI MSI has emerged as a powerful imaging technique that complements conventional MS-based approaches by providing unique information on the spatial distribution of lipids within tissue compartments9,10.
The typical workflow of lipidomics consists of sample preparation, data acquisition using mass-spectrometry technology, and data analysis11. The study of lipids and metabolites in samples has led to the emergence of techniques to understand the physiological and pathological conditions of metabolic processes in organisms. While understanding biological interactions is important, the sensitivity of lipids and metabolites makes them difficult to image and identify without dyes or other modification. Changes in metabolite levels or distribution may lead to phenotypic changes. One tool used for metabolomic profiling is MALDI MSI, a label-free, in situ imaging technique capable of detecting hundreds of molecules simultaneously. MALDI imaging allows for the visualization of metabolites and lipids in samples while preserving their integrity and spatial distribution. Previous technology for lipid profiling involved the use of radioactive chemicals to individually map lipids, while MALDI imaging forgoes this and allows for the detection of a range of lipids simultaneously.
Lipid metabolism and homeostasis play important functions in cell physiology, such as the maintenance and development of the nervous system. One essential aspect of nervous system lipid metabolism is the lipid shuttling between neurons and glial cells, which is mediated by molecular carrier lipoproteins, including very-low-density lipoprotein (VLDL), low-density lipoproteins (LDL), and high-density lipoproteins (HDL)12. Lipoproteins contain apolipoproteins (Apo), such as ApoB and ApoD, which function as structural blocks of lipid cargo and as ligands for lipoprotein receptors. The neuron-glia crosstalk of lipids involves multiple players such as glia-derived ApoD, ApoE, and ApoJ, and their neuronal LDL receptors (LDLRs)13,14. In Drosophila, apolipophorin, a member of the ApoB family, is a major hemolymph lipid carrier15. Apolipophorin has two closely related lipophorin receptors (LpRs), LpR1 and LpR2, which are homologs of mammalian LDLR15,16. In previous studies, the astrocyte-secreted lipocalin Glial Lazarillo (GLaz), a Drosophila homolog of human ApoD, and its neuronal receptor LpR1 were discovered to cooperatively mediate neuron-glia lipid shuttling, thus regulating dendrite morphogenesis17. Therefore, it was speculated that the loss of LpR1 would cause a decrease in overall lipid content in the Drosophila brain. MALDI MSI would be a suitable tool for profiling the lipid contents in small tissues of LpR1&#8722;/&#8722; mutant and wild-type Drosophila brains, as demonstrated in this study.
Despite the growing popularity of MALDI MSI, the instrument&#39;s high cost and experimental complexity often impede its implementation in individual laboratories. Thus, most MALDI MSI studies are conducted using shared core facilities. As with other applications of MALDI MSI, a careful sample preparation process for lipidomics is critical to achieve reliable results. However, because sample slide preparation is typically performed in individual research laboratories, there is a possibility of variation in MALDI MSI acquisition. To combat this, this paper aims to provide a detailed protocol for the sample preparation of small biological samples prior to MALDI MSI measurement using lipid analysis of a large group of adult Drosophila brains in positive ion mode as an example11,17. However, some phospholipid classes and the majority of small metabolites are favorably detected by MALDI imaging in negative ion mode, which was described previously11. Therefore, with these two example studies, we hope to provide detailed sample preparation protocols of various combinations: free-standing large tissue versus embedded small tissue, thaw-mounting versus warm-slide mounting, and positive ion mode versus negative ion mode.

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			<td>2,5-Dihydroxybenzoic acid (DHB)</td>
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			<td>Andwin Scientific&#160;CRYOMOLD 15X15X5</td>
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			<td>NC9464347</td>
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			<td>Andwin Scientific&#160;Tissue-Tek CRYO-OCT Compound</td>
			<td>Fisher Scientific</td>
			<td>14-373-65</td>
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			<td>Artist brush MSC #5 1/8 X 9/16 TRIM RED SABLE</td>
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			<td>50-111-2302</td>
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			<td>autoflex speed MALDI-TOF MS system</td>
			<td>Bruker Daltonics Inc</td>
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			<td>MALDI-TOF MS instrument</td>
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			<td>BD&#160;Syringe with Luer-Lok Tips</td>
			<td>Fisher Scientific</td>
			<td>14-823-16E</td>
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			<td>BD&#160;Vacutainer General Use Syringe Needles</td>
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			<td>23-021-020</td>
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			<td>Bruker Daltonics&#160;GLASS SLIDES MALDI IMAGNG</td>
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			<td>Perfection V600</td>
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			<td>Bruker Daltonics Inc</td>
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			<td>Bruker MS imaging analysis software</td>
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			<td>HPLC Grade Methanol</td>
			<td>Fisher Scientific</td>
			<td>MMX04751</td>
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			<td>HPLC Grade Water</td>
			<td>Fisher Scientific</td>
			<td>W5-1</td>
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			<td>HTX M5 Sprayer</td>
			<td>HTX Technologies, LLC</td>
			<td></td>
			<td>Automatic heated matrix sprayer</td>
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			<td>Kimberly-Clark Professional&#160;Kimtech Science Kimwipes Delicate Task Wipers</td>
			<td>Fisher Scientific</td>
			<td>06-666A</td>
			<td></td>
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			<td>MSC&#160;Ziploc Freezer Bag</td>
			<td>Fisher Scientific</td>
			<td>50-111-3769</td>
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			<td>SCiLS Lab (2015b)</td>
			<td>SCiLS Lab</td>
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			<td>Advanced MALDI MSI data analysis software</td>
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			<td>Thermo Scientific&#160;CryoStar NX50 Cryostat</td>
			<td>Fisher Thermo Scientific</td>
			<td>95-713-0</td>
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			<td>Thermo Scientific&#160;Nalgene Transparent Polycarbonate Classic Design Desiccator</td>
			<td>Fisher Scientific</td>
			<td>08-642-7</td>
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sample preparation for rapid lipid analysis in drosophila brain using matrix-assisted laser desorption/ionization mass spectrometry imaging

Lipid profiling, or lipidomics, is a well-established technique used to study the entire lipid content of a cell or tissue. Information acquired from&#160;lipidomics is valuable in studying the pathways involved in development, disease, and cellular metabolism. Many tools and instrumentations have aided lipidomics projects, most notably various combinations of mass spectrometry and liquid chromatography techniques. Matrix-assisted laser desorption/ionization mass spectrometry imaging (MALDI MSI) has recently emerged as a powerful imaging technique that complements conventional approaches. This novel technique provides unique information on the spatial distribution of lipids within tissue compartments, which was previously unattainable without the use of excessive modifications. The sample preparation of the MALDI MSI approach is critical and, therefore, is the focus of this paper. This paper presents a rapid lipid analysis of a large number of Drosophila brains embedded in optimal cutting temperature compound (OCT) to provide a detailed protocol for the preparation of small tissues for lipid analysis or metabolite and small molecule analysis through MALDI MSI.

The loss of the neuronal receptor LpR1 of the astrocyte-secreted lipocalin Glial Lazarillo (GLaz), a Drosophila homolog of human ApoD, was hypothesized to be able to cause a decrease in overall lipid content in the Drosophila brain. To test this, MALDI MSI was used to profile the lipids in LpR1&#8722;/&#8722; mutant and wild-type Drosophila brains, which is elaborated on below.
The experiment was performed according to the workflow shown in <stron...

Watch this Scientific Journal Video about Sample Preparation for Rapid Lipid Analysis in Drosophila Brain Using Matrix-Assisted Laser Desorption/Ionization Mass Spectrometry Imaging at JoVE.com

Sample Preparation for Rapid Lipid Analysis in Drosophila Brain Using Matrix-Assisted Laser Desorption/Ionization Mass Spectrometry Imaging

The aim of this protocol is to provide detailed guidance on the proper sample preparation for lipid and metabolite analysis in small tissues, such as the Drosophila brain, using matrix-assisted laser desorption/ionization (MALDI) mass spectrometry imaging.

Sample Preparation for Rapid Lipid Analysis in Drosophila Brain Using Matrix-Assisted Laser Desorption/Ionization Mass Spectrometry Imaging

Lipid profiling, or lipidomics, is a well-established technique used to study the entire lipid content of a cell or tissue. Information acquired ...

sample-preparation-for-rapid-lipid-analysis-drosophila-brain-using

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3.0K Views.  the City University of New York, Graduate Center New York. The aim of this protocol is to provide detailed guidance on the proper sample preparation for lipid and metabolite analysis in small tissues, such as the Drosophila brain, using matrix-assisted laser desorption/ionization (MALDI) mass spectrometry imaging. 

Video: Sample Preparation for Rapid Lipid Analysis in Drosophila Brain Using Matrix-Assisted Laser Desorption/Ionization Mass Spectrometry Imaging

MALDI Sample Preparation: the Ultra Thin Layer Method

This video demonstrates the preparation of an ultra-thin matrix/analyte layer for analyzing peptides and proteins by Matrix-Assisted Laser Desorption Ionization Mass Spectrometry (MALDI-MS) 1,2. The ultra-thin layer method involves the production of a substrate layer of matrix crystals (alpha-cyano-4-hydroxycinnamic acid) on the sample plate, which serves as a seeding ground for subsequent crystallization of a matrix/analyte mixture. Advantages of the ultra-thin layer method over other sample deposition approaches (e.g. dried droplet) are that it provides (i) greater tolerance to impurities such as salts and detergents, (ii) better resolution, and (iii) higher spatial uniformity. This method is especially useful for the accurate mass determination of proteins. The protocol was initially developed and optimized for the analysis of membrane proteins and used to successfully analyze ion channels, metabolite transporters, and receptors, containing between 2 and 12 transmembrane domains 2. Since the original publication, it has also shown to be equally useful for the analysis of soluble proteins. Indeed, we have used it for a large number of proteins having a wide range of properties, including those with molecular masses as high as 380 kDa 3. It is currently our method of choice for the molecular mass analysis of all proteins. The described procedure consistently produces high-quality spectra, and it is sensitive, robust, and easy to implement.

This video demonstrates the preparation of an ultra-thin matrix/analyte layer for analyzing peptides and proteins by Matrix-Assisted Laser Desorption Ionization Mass Spectrometry (MALDI-MS).

This video demonstrates the preparation of an ultra-thin matrix/analyte layer for analyzing peptides and proteins by Matrix-Assisted Laser Desorption ...

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as tissue morphogenesis, cell-cell adhesion, or cell death. Drosophila embryos expressing GFP are readily imaged using either stereoscopic or confocal microscopy. A goal of any live-imaging protocol is to minimize detrimental effects such as dehydration and hypoxia. Previous protocols for preparing Drosophila embryos for live-imaging analysis have involved placing dechorionated embryos in halocarbon oil and sandwiching them between a halocarbon gas-permeable membrane and a coverslip1-3. The introduction of compression through mounting embryos in this manner represents an undesirable complication for any biomechanical-based analysis of morphogenesis. Our method, which we call the hanging drop protocol, results in excellent viability of embryos during live imaging and does not require that embryos be compressed. Briefly, the hanging drop protocol involves the placement of embryos in a drop of halocarbon oil that is suspended from a coverslip, which is, in turn, fixed in position over a humid chamber. In addition to providing gas exchange and preventing dehydration, this arrangement takes advantage of the buoyancy of embryos in halocarbon oil to prevent them from drifting out of position during timelapse acquisition. This video describes in detail how to collect and prepare Drosophila embryos for live imaging using the hanging drop protocol. This protocol is suitable for imaging dechorionated embryos using stereomicroscopy or any upright compound fluorescence microscope.

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

A simple, inexpensive, and effective method of preparing Drosophila embryos for live-imaging analysis is presented. Our protocol provides humidity and gas exchange and does not compress the Drosophila embryo. This method is suitable for GFP-based live imaging of Drosophila embryos using a stereomicroscope or upright compound microscope.

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as ...

Profiling Thiol Redox Proteome Using Isotope Tagging Mass Spectrometry

Pseudomonas syringae pv. tomato strain DC3000 not only causes bacterial speck disease in Solanum lycopersicum but also on Brassica species, as well as on Arabidopsis thaliana, a genetically tractable host plant1,2. The accumulation of reactive oxygen species (ROS) in cotyledons inoculated with DC3000 indicates a role of ROS in modulating necrotic cell death during bacterial speck disease of tomato3. Hydrogen peroxide, a component of ROS, is produced after inoculation of tomato plants with Pseudomonas3. Hydrogen peroxide can be detected using a histochemical stain 3'-3' diaminobenzidine (DAB)4. DAB staining reacts with hydrogen peroxide to produce a brown stain on the leaf tissue4. ROS has a regulatory role of the cellular redox environment, which can change the redox status of certain proteins5. Cysteine is an important amino acid sensitive to redox changes. Under mild oxidation, reversible oxidation of cysteine sulfhydryl groups serves as redox sensors and signal transducers that regulate a variety of physiological processes6,7. Tandem mass tag (TMT) reagents enable concurrent identification and multiplexed quantitation of proteins in different samples using tandem mass spectrometry8,9. The cysteine-reactive TMT (cysTMT) reagents enable selective labeling and relative quantitation of cysteine-containing peptides from up to six biological samples. Each isobaric cysTMT tag has the same nominal parent mass and is composed of a sulfhydryl-reactive group, a MS-neutral spacer arm and an MS/MS reporter10. After labeling, the samples were subject to protease digestion. The cysteine-labeled peptides were enriched using a resin containing anti-TMT antibody. During MS/MS analysis, a series of reporter ions (i.e., 126-131 Da) emerge in the low mass region, providing information on relative quantitation. The workflow is effective for reducing sample complexity, improving dynamic range and studying cysteine modifications. Here we present redox proteomic analysis of the Pst DC3000 treated tomato (Rio Grande) leaves using cysTMT technology. This high-throughput method has the potential to be applied to studying other redox-regulated physiological processes.

Reactive oxygen species level is elevated when cells encounter stress conditions. Here we show the example of 3'-3' diaminobenzidine staining as well as cysTMT labeling and mass spectrometry to profile the redox proteome in Pseudomonas syringae treated tomato leaves.

Pseudomonas syringae pv. tomato strain DC3000 not only causes bacterial speck disease in Solanum lycopersicum but also on Brassica species, as well as on ...

Dithranol as a Matrix for Matrix Assisted Laser Desorption/Ionization Imaging on a Fourier Transform Ion Cyclotron Resonance Mass Spectrometer

Mass spectrometry imaging (MSI) determines the spatial localization and distribution patterns of compounds on the surface of a tissue section, mainly using MALDI (matrix&#160;assisted laser desorption/ionization)-based analytical techniques. New matrices for small-molecule MSI, which can improve the analysis of low-molecular weight (MW) compounds, are needed. These matrices should provide increased analyte signals while decreasing MALDI background signals. In addition, the use of ultrahigh-resolution instruments, such as Fourier transform ion cyclotron resonance (FTICR) mass spectrometers, has&#160;the ability to resolve analyte signals from matrix signals, and this can partially overcome many problems associated with the background originating from the MALDI matrix. The reduction in the intensities of the metastable matrix clusters by FTICR MS can also help to overcome some of the interferences associated with matrix peaks on other instruments. High-resolution instruments such as the FTICR mass spectrometers are advantageous as they can produce distribution patterns of many compounds simultaneously while still providing confidence in chemical identifications. Dithranol (DT; 1,8-dihydroxy-9,10-dihydroanthracen-9-one) has previously been reported as a MALDI matrix for tissue imaging. In this work, a protocol for the use of DT for MALDI imaging of endogenous lipids from the surfaces of mammalian tissue sections, by positive-ion MALDI-MS, on an ultrahigh-resolution hybrid quadrupole FTICR instrument has been provided.

Dithranol (DT; 1,8-dihydroxy-9,10-dihydroanthracen-9-one) has previously been reported as a MALDI matrix for tissue imaging of small molecules; protocols for the use of DT for the MALDI imaging of endogenous lipids on the surface of tissue sections&#160;by positive-ion MALDI-MS on an ultrahigh-resolution quadrupole-FTICR instrument are provided here.

Mass spectrometry imaging (MSI) determines the spatial localization and distribution patterns of compounds on the surface of a tissue section, mainly ...

Rapid Analysis and Exploration of Fluorescence Microscopy Images

Despite rapid advances in high-throughput microscopy, quantitative image-based assays still pose significant challenges. While a variety of specialized image analysis tools are available, most traditional image-analysis-based workflows have steep learning curves (for fine tuning of analysis parameters) and result in long turnaround times between imaging and analysis. In particular, cell segmentation, the process of identifying individual cells in an image, is a major bottleneck in this regard.
Here we present an alternate, cell-segmentation-free workflow based on PhenoRipper, an open-source software platform designed for the rapid analysis and exploration of microscopy images. The pipeline presented here is optimized for immunofluorescence microscopy images of cell cultures and requires minimal user intervention. Within half an hour, PhenoRipper can analyze data from a typical 96-well experiment and generate image profiles. Users can then visually explore their data, perform quality control on their experiment, ensure response to perturbations and check reproducibility of replicates. This facilitates a rapid feedback cycle between analysis and experiment, which is crucial during assay optimization. This protocol is useful not just as a first pass analysis for quality control, but also may be used as an end-to-end solution, especially for screening. The workflow described here scales to large data sets such as those generated by high-throughput screens, and has been shown to group experimental conditions by phenotype accurately over a wide range of biological systems. The PhenoBrowser interface provides an intuitive framework to explore the phenotypic space and relate image properties to biological annotations. Taken together, the protocol described here will lower the barriers to adopting quantitative analysis of image based screens.

Here we describe a workflow for rapidly analyzing and exploring collections of fluorescence microscopy images using PhenoRipper, a recently developed image-analysis platform.

Despite rapid advances in high-throughput microscopy, quantitative image-based assays still pose significant challenges. While a variety of specialized ...

Sample Preparation for Mass Cytometry Analysis

Mass cytometry utilizes antibodies conjugated with heavy metal labels, an approach that has greatly increased the number of parameters and opportunities for deep analysis well beyond what is possible with conventional fluorescence-based flow cytometry. As with any new technology, there are critical steps that help ensure the reliable generation of high-quality data. Presented here is an optimized protocol that incorporates multiple techniques for the processing of cell samples for mass cytometry analysis. The methods described here will help the user avoid common pitfalls and achieve consistent results by minimizing variability, which can lead to inaccurate data. To inform experimental design, the rationale behind optional or alternative steps in the protocol and their efficacy in uncovering new findings in the biology of the system being investigated is covered. Lastly, representative data is presented to illustrate expected results from the techniques presented here.

This article describes the collection and processing of samples for mass cytometry analysis.

Mass cytometry utilizes antibodies conjugated with heavy metal labels, an approach that has greatly increased the number of parameters and opportunities ...

Sample Preparation and Imaging of Exosomes by Transmission Electron Microscopy

Exosomes are nano-sized extracellular vesicles secreted by body fluids and are known to represent the characteristics of cells that secrete them. The contents and morphology of the secreted vesicles reflect cell behavior or physiological status, for example cell growth, migration, cleavage, and death. The exosomes&#39; role may depend highly on size, and the size of exosomes varies from 30 to 300 nm. The most widely used method for exosome imaging is negative staining, while other results are based on Cryo-Transmission Electron Microscopy, Scanning Electron Microscopy, and Atomic Force Microscopy. The typical exosome&#39;s morphology assessed through negative staining is a cup-shape, but further details are not yet clear. An exosome well-characterized through structural study is necessary particular in medical and pharmaceutical fields. Therefore, function-dependent morphology should be verified by electron microscopy techniques such as labeling a specific protein in the detailed structure of exosome. To observe detailed structure, ultrathin sectioned images and negative stained images of exosomes were compared. In this protocol, we suggest transmission electron microscopy for the imaging of exosomes including negative staining, whole mount immuno-staining, block preparation, thin section, and immuno-gold labelling.

This protocol describes the various techniques necessary for transmission electron microscopy including negative staining, ultrathin sectioning for detailed structure, and immuno-gold labelling to determine the positions of specific proteins in exosomes.

Exosomes are nano-sized extracellular vesicles secreted by body fluids and are known to represent the characteristics of cells that secrete them. The ...

Miniaturized Sample Preparation for Transmission Electron Microscopy

Due to recent technological progress, cryo-electron microscopy (cryo-EM) is rapidly becoming a standard method for the structural analysis of protein complexes to atomic resolution. However, protein isolation techniques and sample preparation methods for EM remain a bottleneck. A relatively small number (100,000 to a few million) of individual protein particles need to be imaged for the high-resolution analysis of proteins by the single particle EM approach, making miniaturized sample handling techniques and microfluidic principles feasible.
A miniaturized, paper-blotting-free EM grid preparation method for sample pre-conditioning, EM grid priming and post processing that only consumes nanoliter-volumes of sample is presented. The method uses a dispensing system with sub-nanoliter precision to control liquid uptake and EM grid priming, a platform to control the grid temperature thereby determining the relative humidity above the EM grid, and a pick-and-plunge-mechanism for sample vitrification. For cryo-EM, an EM grid is placed on the temperature-controlled stage and the sample is aspirated into a capillary. The capillary tip is positioned in proximity to the grid surface, the grid is loaded with the sample and excess is re-aspirated into the microcapillary. Subsequently, the sample film is stabilized and slightly thinned by controlled water evaporation regulated by the offset of the platform temperature relative to the dew-point. At a given point the pick-and-plunge mechanism is triggered, rapidly transferring the primed EM grid into liquid ethane for sample vitrification. Alternatively, sample-conditioning methods are available to prepare nanoliter-sized sample volumes for negative stain (NS) EM.
The methodologies greatly reduce sample consumption and avoid approaches potentially harmful to proteins, such as the filter paper blotting used in conventional methods. Furthermore, the minuscule amount of sample required allows novel experimental strategies, such as fast sample conditioning, combination with single-cell lysis for &#34;visual proteomics,&#34; or &#34;lossless&#34; total sample preparation for quantitative analysis of complex samples.

An instrument and methods for the preparation of nanoliter-sized sample volumes for transmission electron microscopy is presented. No paper-blotting steps are required, thus avoiding the detrimental consequences this can have for proteins, significantly reducing sample loss and enabling the analysis of single cell lysate for visual proteomics.

Due to recent technological progress, cryo-electron microscopy (cryo-EM) is rapidly becoming a standard method for the structural analysis of protein ...

Measurement of Energy Metabolism in Explanted Retinal Tissue Using Extracellular Flux Analysis

High acuity vision is a heavily energy-consuming process, and the retina has developed several unique adaptations to precisely meet such demands while maintaining transparency of the visual axis. Perturbations to this delicate balance cause blinding illnesses, such as diabetic retinopathy. Therefore, the understanding of energy metabolism changes in the retina during disease is imperative to the development of rational therapies for various causes of vison loss. The recent advent of commercially-available extracellular flux analyzers has made the study of retinal energy metabolism more accessible. This protocol describes the use of such an analyzer to measure contributions to retinal energy supply through its two principle arms - oxidative phosphorylation and glycolysis - by quantifying changes in oxygen consumption rates (OCR) and extracellular acidification rates (ECAR) as proxies for these pathways. This technique is readily performed in explanted retinal tissue, facilitating assessment of responses to multiple pharmacologic agents in a single experiment. Metabolic signatures in retinas from animals lacking rod photoreceptor signaling are compared to wild-type controls using this method. A major limitation in this technique is the lack of ability to discriminate between light-adapted and dark-adapted energy utilization, an important physiologic consideration in retinal tissue.

This technique describes real time recording of oxygen consumption and extracellular acidification rates in explanted mouse retinal tissues using an extracellular flux analyzer.

High acuity vision is a heavily energy-consuming process, and the retina has developed several unique adaptations to precisely meet such demands while ...

Sample Preparation for Mass-spectrometry-based Proteomics Analysis of Ocular Microvessels

The use of isolated ocular blood vessels in vitro to decipher the pathophysiological state of the eye using advanced technological approaches has greatly expanded our understanding of certain diseases. Mass spectrometry (MS)-based proteomics has emerged as a powerful tool to unravel alterations in the molecular mechanisms and protein signaling pathways in the vascular beds in health and disease. However, sample preparation steps prior to MS analyses are crucial to obtain reproducible results and in-depth elucidation of the complex proteome. This is particularly important for preparation of ocular microvessels, where the amount of sample available for analyses is often limited and thus, poses a challenge for optimum protein extraction. This article endeavors to provide an efficient, rapid and robust protocol for sample preparation from an exemplary retrobulbar ocular vascular bed employing the porcine short posterior ciliary arteries. The present method focuses on protein extraction procedures from both the supernatant and pellet of the sample following homogenization, sample cleaning with centrifugal filter devices prior to one-dimensional gel electrophoresis and peptide purification steps for label-free quantification in a liquid chromatography-electrospray ionization-linear ion trap-Orbitrap MS system. Although this method has been developed specifically for proteomics analyses of ocular microvessels, we have also provided convincing evidence that it can also be readily employed for other tissue-based samples.

Proteome characterization of ocular microvascular beds is pivotal for in-depth understanding of many ocular pathologies in humans. This study demonstrates an effective, rapid, and robust method for protein extraction and sample preparation from small blood vessels employing the porcine short posterior ciliary arteries as model vessels for mass-spectrometry-based proteomics analyses.

The use of isolated ocular blood vessels in vitro to decipher the pathophysiological state of the eye using advanced technological approaches has greatly ...